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1984

Advances
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in Insect Rearing

Edited by E. G. King and N. C. Leppla

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Abstract

This book contains greatly elaborated and revised versions of 36 papers presented at a
conference sponsored by the U.S. Department of Agriculture and the Insect Rearing
Group in March 1980. These papers deal comprehensively with the genetics of reared in¬
sects, especially the decline in variability and performance and ways to guard against
both; diets and containers; engineering problems and solutions for insect-rearing facilities
and systems; the discovery and control of pathogens and micro-organisms in insect rear¬
ing; actual rearing systems for a diverse selection of insects intended for a variety of uses;
and the management of insect-rearing programs, including data-processing techniques,
systems management, and quality control.

Editors’ addresses: E. G. King, Southern Field Crop Insect Management Laboratory,
Agricultural Research Service, U.S. Department of Agriculture, P.O. Box 225, Stoneville,
Miss. 38776. N. C. Leppla, Insect Attractants, Behavior, and Basic Biology Research
Laboratory, Agricultural Research Service, U.S. Department of Agriculture, P.O. Box
14565, Gainesville, Fla. 32604.

This publication contains the results of research only. Mention of pesticides does not con¬
stitute a recommendation for use, nor does it imply that the pesticides are registered
under the Federal Insecticide, Fungicide, and Rodenticide Act as amended. The use of
trade names in this publication does not constitute a guarantee, warranty, or endorsement
of the products by the U.S. Department of Agriculture.

Copies of this publication can be purchased from the U.S. Government Printing Office,
Washington, D.C. 20402. When ordering by mail, ask for the publication by title. For
faster service, call the GPO order desk at (202) 783-3238 and charge the publication to
your Visa, MasterCard, or GPO Deposit Account. Discounts are available for pur¬
chases of more than 100 copies mailed to the same address.

Published by the Agricultural Research Service (Southern Region), U.S. Department of
Agriculture, New Orleans, La. January 1984.

ii

802524

Contents

Page

Preface . v

Message From the Conference Chairman . vii

Foreword. What Colonization of Insects Means to Research and Pest Management ix

Recommendations of the Conference . xiii

Section 1. Establishment and Maintenance of Insect Colonies Through Genetic
Control . 1

Genetic Changes During Insect Domestication

Alan C. Bartlett . 2

Artificial Selection of Desired Characteristics in Insects

Anita M. Collins . 9

Maintenance of Genetic Variability in Reared Insects

Dennis J. Joslyn . 20

Section 2. Diets and Containerization for Insect Rearing . 31

Insect Diets. Historical Developments, Recent Advances, and Future Prospects

Pritam Singh . 32

Ingredients for Insect Diets. Quality Assurance, Sources, and Storage and
Handling

F. D. Brewer and Oliver Lindig . 45

Containerization for Rearing Insects

Robert L. Burton and W. Deryck Perkins . 51

Section 3. Engineering for Insect Rearing . 57

Controlled Environments for Insects and Personnel in Insect-Rearing Facilities

Charles D. Owens . 58

Controlling Respiratory Hazards in Insectaries

Wayne W. Wolf . 64

General Requirements for Facilities That Mass-Rear Insects

Jack G. Griffin . 70

Automation in Insect Rearing

E. A. Harrell and C. W. Gantt . 74

Materials Handling in Insect Rearing

J. L. Goodenough . 77

The Closed-Loop System of Quality Control in Insect Rearing

J. C. Webb . : . 87

Systems Analysis and Modeling in Mass Rearing and Control of Insects

D. G. Haile and D. E. Weidhaas . 90

Section 4. Control of Pathogens and Microbial Contaminants in Insect Rearing 95

Recognition and Diagnosis of Diseases in Insectaries and the Effects of Disease
Agents on Insect Biology

Ronald H. Goodwin . 96

Micro-organisms as Contaminants and Pathogens in Insect Rearing

Martin Shapiro . 130

Microbial Contamination in Insectaries. Occurrence, Prevention, and Control

Peter P. Sikorowski . . 143

Page

Section 5. Production, Use, and Quality Testing in Insect Rearing . 155

Production of the Gypsy Moth, Lymantria dispar, for Research and Biological
Control

Thomas M. ODell et al . 156

Rearing the Tobacco Budworm, Heliothis virescens, and Com Earworm, HeUothis
zea

J. R. Raulston and E. G. King . 167

Mass Rearing the Pink Bollworm, Pectinophora gossypiella

Fred D. Stewart . 176

Production of Boll Weevils, Anthonomus grandis grandis

J. Roberson and J. E. Wright . 188

Mass Production of Screwworm Flies, Cochliomyia hominivorax

Harold A. Brown . 193

Improved Techniques for Mass Rearing Anopheles albimanus

Donald L. Bailey and J. A. Seawright . 200

Some Systems for Production of Eight Entomophagous Arthropods

E. G. King and R. K. Morrison . 206

A Laboratory Method for Mass Rearing the Eastern Spruce Budworm,

Choristoneura fumiferana

D. G. Grisdale . 223

Fractional Colony Propagation. A New Insect-Rearing System

J. David Hoffman et al . 232

Production of Insects for Industry. The Dow Chemical Rearing Program

W. R. Fisher . 234

Industrial Insect Production for Insecticide Screening

R. E. Wheeler . 240

Mass Rearing the Cabbage Looper, Trichoplusia ni

N. C. Leppla et al . 248

Section 6. Management of Insect-Rearing Systems . 255

Putting the Control in Quality Control in Insect Rearing

D. L. Chambers and T. R. Ashley . 256

Academic Training for Insect Rearing

Marion A. Brooks . 261

Health and Safety in Arthropod Rearing

Robert A. Wirtz . 263

Systems Analysis and Automated Data Processing in Insect Rearing. A System
for the Biting Gnat Culicoides variipennis and Mosquitoes

David H. Akey et al . 269

Systems Management of Insect-Population-Suppression Programs Based on
Mass Production of Biological-Control Organisms

N. C. Leppla . 292

The Insectary Manager

W. R. Fisher . 295

Management of Insect Production

Charles P. Schwalbe and O. T. Forrester . 300

Politics in Insect Rearing and Control. The Medfly Program in Guatemala and
Southern Mexico

Patrick Patton . 304

IV

Preface

This book is an elaboration of papers and discussions
from the conference “Advances and Challenges in Insect
Rearing” held in Atlanta, Ga., on March 4-6, 1980. We
have retained the original organization of the conference
in the book’s six sections. The first four sections are
basic to insect rearing, and the fifth conveys the state of
the art with descriptions of several rearing systems. The
sixth and final section discusses various topics important
to management of insect-rearing systems. In the “Mes¬
sage From the Conference Chairman,” R. F. Moore out¬
lines the questions addressed by the conference and
discusses its origins and the reasons for its organization.
And the rationale and need for insect rearing and re¬
search is elegantly presented by E. F. Knipling in the
Foreword, based on his keynote address. Finally, many
recommendations about further research and program
needs developed from the conference, and these are in¬
cluded in “Recommendations of the Conference.”

Thanks are due to many people for support of the confer¬
ence and for completion of this book. Certainly, R. F.
Moore, the conference chairman; section leaders; and
authors of the various papers deserve tribute for their
contributions. We also appreciate the support of admin¬
istrators in the U.S. Department of Agriculture, State
agricultural experiment stations, and the private sector.

E. G. King
and

N. C. Leppla,

Research entomologists,

Agricultural Research Service

v

Message From the Conference Chairman

The impetus for this conference was an item by Tom
ODell in the March 8, 1978, issue of Frass, the Insect
Rearing Group’s newsletter. He indicated that many peo¬
ple were interested in a workshop on rearing insects. The
last conference on insect nutrition and rearing was spon¬
sored by the U.S. Department of Agriculture (USDA) in
March 1963; since then, many advances and changes
have occurred. Because I aim a technical adviser for in¬
sect rearing and nutrition on the national research pro¬
gram “Non-Commodity Research for Insect Control” of
USDA’s Agricultural Research Service, it was appropri¬
ate for me to support a workshop on this topic. I con¬
tacted Tom and others to determine if plans were being
made to hold such a workshop. On learning that no active
plans were in progress, I invited 10 interested scientists
to a meeting in Washington, D.C., on April 27, 1978, to
determine if a workshop should be developed.

Their interest, enthusiasm, and concern left no doubt that
there should be a workshop; but limiting the subject and
developing a format proved difficult. What were to be the
purpose, subject matter, and scope, and how was the con¬
ference to be documented? There were concerns that in¬
sect rearing was not recognized as an important area of
entomology, that administrative support for rearing was
inadequate, and that much information was not readily
available. How could all these interests be satisfied in one
short conference?

After some prolonged discussions, we arrived at the
following objectives:

1. To assemble the scientific principles of insect rearing
that have been established in recent years. These
would include guidelines for establishing and main¬
taining colonies of insects for specific purposes.

2. To identify problem areas in insect-rearing programs
and develop specific recommendations for problem
solving. These recommendations would include pro¬
cedures for research, development, and implemen¬
tation.

3. To establish the complexity and integrity of insect
rearing as a field of scientific research.

4. To document the state of the art of insect rearing
and establish a reference for direction of the science
through publication of the conference proceedings.

Rearing of insects has been regarded as nontechnical
labor, so criticism of the research effort expended on it
has been considerable. This attitude has persisted be¬

cause early programs reared insects that were used pri¬
marily for screening pesticides and for associated
research. At that time, we depended too much on chem¬
icals for controlling insects; now, with the advent of al¬
ternate methods of control, insect rearing has assumed
new and major importance. Since most of these methods
require the ability to rear insects of specified quality, the
following highly technical questions have emerged:

1. How are laboratory -reared insects different from wild
ones?

2. What is the most economical means of meeting the
nutritional requirements of insects?

3. What are the most labor-efficient methods of rearing
insects?

4. How can we be certain these insects will perform ade¬
quately in a control program?

These questions have been addressed in the six com¬
plementary sections of the program:

Section 1, “Establishment and Maintenance of Insect
Colonies Through Genetic Control,” discusses some ge¬
netic reasons for the differences observed between labor¬
atory and wild insects. The organizing committee expects
that some principles and guildelines for colonizing insects
will be found in this section. There will also be un¬
answered questions that will require future research. For
example, can we, now or in the future, have breeding pro¬
grams to develop strains of insects for special purposes?

Section 2, “Diets and Containerization for Insect Rear¬
ing,” gives information on the ingredients of diets and on
methods of handling and using artificial diets. Quality
and cost of dietary ingredients increase in importance as
more insects are reared. Diets that are more balanced in
protein, carbohydrates, lipids, vitamins, and minerals are
needed for more efficient utilization; an economical
substitute for agar is also vital.

Section 3, “Engineering for Insect Rearing,” recognizes
the contribution of engineers to reducing the labor re¬
quired to handle insects, a major expense in rearing. In
the recent past, facilities for rearing insects were
modified rooms or buildings; today, however, insectaries
are sometimes specially designed. Special designs are
necessary because the needs of the rearing programs
vary. Some facilities produce large quantities of insects
for control purposes. Others produce small research
cultures. And still others produce intermediate amounts.

Mass-rearing programs will probably always need special¬
ly designed facilities. But, perhaps we are approaching
the time when a standardized facility can be designed for
intermediate research and small-scale production.

Section 4, “Control of Pathogens and Microbial Contam¬
inants in Insect Rearing,” demonstrates the importance
of controlling insect pathogens. There is increasing evi¬
dence that microbial contaminants affect both the quality
of laboratory-reared insects used for alternate methods of
control and the results of research obtained with these
insects.

Section 5, “Production, Use, and Quality Testing in In¬
sect Rearing,” details the rearing of several insect
species. Even though the insects discussed in this section
are mainly phytophagous, the general rearing principles
also apply to entomophages. Quality testing, for example,
is of great importance for any insects used in alternate
methods of control. Quality tests in the screwworm,
Cochliomyia hominivorax (Coquerel), program measured
the size of the last-instar larvae, survival in the labora¬
tory, and mating competitiveness. Measurements such as
rate of development to pupal or adult stages, number of
eggs produced, and yield of adults are used in other pro¬
grams. These acre rather gross indicators; and, for some
species, several diets or rearing conditions will produce
apparently equivalent insects. Presently, we are looking
at more subtle tests such as electrophysiological response
to light and sound, pheromone production, locomotor ac¬
tivity, enzyme levels, and other variables that seem likely

to affect performance. These tests should tell us how the
insects vary in these qualities from week to week. But, to
date, we have few direct relationships between laboratory
tests and field performance. A further limitation to cur¬
rent quality testing is that the test results are obtained
after the insects have been released. A goal of quality
testing should be to develop tests that will predict
performance.

Section 6, “Management of Insect-Rearing Systems,”
brings together all the complexities of the processes of in¬
sect rearing and use. This systematization is especially
apparent in the large programs designed to support alter¬
nate methods of control. These massive enterprises, re¬
quiring procedures based on current research, must be
managed by qualified professionals.

Finally, I want to acknowledge the support and assist¬
ance of the members of the Organizing and Planning
Committee, their research leaders, area directors, the Na¬
tional Program Staff, and regional administrative of¬
ficers. I especially want to thank the section leaders and
the others who assisted unofficially for their interest and
dedication. Their helpful assistance and encouragement
has made development of this conference a real pleasure.

R. F. Moore,

Supervisory research entomologist,
Agricultural Research Service

viii

Foreword

What Colonization of Insects Means
to Research and Pest Management

Since the ability to rear insects is important to virtually
every aspect of entomological research, and I am in¬
terested in all aspects of entomological research, it is a
pleasure to attend this conference to keep abreast of the
field’s problems and progress. Insect-rearing technology
can make direct contributions to the management of
many destructive pests by enabling strategies that are
sound in principle, economically advantageous, and vir¬
tually without hazard to nontarget organisms. But, some
scientists have been critical of the amount of research ef¬
fort devoted to insect rearing. So, in my address at the
1977 annual meeting of the Entomological Society of
America, I cited advances in insect rearing as being
among the most important developments in entomology
during the last few decades. I will reiterate this viewpoint
here because this field continues to warrant high priority.

Most entomologists take for granted our present capabil¬
ity for maintaining colonies of insects. But, when my
career began in the early 1930’s, colonies were limited to
a few easy-to-rear species such as fruit flies; house flies,
Musca domestica Linnaeus; cockroaches; and certain
stored-product insects. Later, in 1936, the screwworm,
Cochliomyia hominivorax (Coquerel), was reared on an ar¬
tificial diet developed by R. Melvin and R. C. Bushland
of the U.S. Department of Agriculture. This milestone
made possible the rapid screening of hundreds of can¬
didate chemicals and other formulations for effectively
treating livestock wounds. Eventually it facilitated sup¬
pression of the screwworm by genetic means.

An early experience that also impressed me with the vital
role of insect rearing in advancing applied entomology oc¬
curred during World War II. The laboratory where I
worked at Orlando, Fla., was engaged in an urgent pro¬
gram to develop insecticides and repellents for protecting
military personnel from pests and disease vectors in
various parts of the world. But methods had not been
devised for rearing the body louse, Pediculus humanus
humanus Linnaeus, and malaria-transmitting Anopheles
mosquitoes on a useful scale. Other pests (such as bed
bugs, Cimex lectularius Linnaeus; fleas; ticks; and
cockroaches) could be reared but not in appreciable
numbers. Therefore, with techniques perfected by G. H.
Culpepper, thousands of body lice were maintained by
permitting them to feed on human hosts twice each day,
and large colonies of the other pests were developed.

These colonies made it possible to screen thousands of
candidate insecticides and repellents in a short time. The
most promising materials were intensively investigated,
and this work led to new control measures for a wide
range of vectors and pests affecting man in many parts
of the world.

After World War II, scientists with the U.S. Department
of Agriculture were encouraged to undertake research on
methods for rearing various crop pests. At that time,
many laboratories were poorly designed and equipped for
this purpose. So improvement of facilities was given high
priority. Rapid progress was made; much of it could be
attributed to basic studies on insect nutrition conducted
by E. S. Vanderzant in cooperation with scientists at
Texas A&M University. These studies led to laboratory
colonization of the boll weevil, Anthonomus grandis gran-
dis Boheman; pink bollworm, Pectinophora gossypiella
(Saunders); and Heliothis spp. Today, through the com¬
bined efforts of many entomologists and engineers with
Federal and State institutions and private industry, it is
possible to produce these major pests by the hundreds of
millions. Early advances in research on insect coloniza¬
tion are described in the book “Insect Colonization and
Mass Production,” edited by C. N. Smith (Academic
Press, New York, 1966).

It is difficult to evaluate the benefits of insect rearing to
the advancement of entomological research. House fly
colonization alone has facilitated discovery of many new
insecticides and refinement of their formulations. Until
appropriate insect colonies became available, important
studies on insect physiology could not be undertaken,
research on the genetics of economic pests was handi¬
capped, and identification and synthesis of sex pher¬
omones were delayed. The ability to rear insects facilitated
research on plant resistance and made possible various
field studies on the dispersal and behavior of insects.
Finally, the field of biological control has been greatly ad¬
vanced by having methods of rearing parasites and
predators for introduction or augmentation and by the
propagation of hosts for insect pathogens.

While development of rearing methods has been vital to
the advancement of all aspects of entomology, basic and
applied, the use of this technology to produce insects for
destruction of their own kind offers an effective alter-

IX

native for controlling certain major pests. But appraisal
of this approach requires both accurate information on
the number of pest insects and the ability to inexpensively
mass-produce enough good quality insects to adequately
flood the target population. Since information on absolute
numbers of insects per unit of area is lacking for most
major pests, indirect estimates must be used. Once com¬
puted, such estimates must be related to the cost of rear¬
ing and releasing the quantities necessary to reach the
required degree of suppression. In most cases, the pests
are so abundant at normal density ranges that managing
them by releasing relatively few sterile or genetically
altered conspecifics is impractical. Therefore, other means
must first be used to suppress these populations. Despite
this requirement, I think the approach offers great prom¬
ise, particularly considering current losses and the need
for suppressive measures that avoid or minimize the
ecological disruptions and costs associated with the use
of broad-spectrum chemicals.

The advent of this two-step process, use of one technique
to reduce high population densities followed by another
that is more efficient at low levels, led to a critical
analysis of the mechanisms of various techniques of in¬
sect control and of how compatible methods might be ap¬
propriately integrated for insect suppression. I have
detailed this fundamental approach in my book, “The
Basic Principles of Insect Population Suppression and
Management” (U.S. Department of Agriculture,
Agriculture Handbook 512, 1979). I am confident that
the use of two or more methods of suppression, together
or in sequence, will eventually lead to more effective and
more acceptable insect-management systems. The use of
biological organisms may be one of the most practical
and desirable techniques to employ in such management
procedures. If so, the development of mass-rearing
technology will become increasingly more important.

When the potential of genetic engineering is fully ap¬
preciated, there should be greater interest in the practical
use of genetic control for insect suppression. In fact, sim¬
ulation models I developed in cooperation with W. Klas-
sen show that several techniques of genetic manipulation
offer prospects of greater efficiency than the conventional
sterile-insect technique. This appraisal is based on results
of genetic effects reported by several investigators. For
example, M. L. Laster of the Mississippi State Agricul¬
tural Experiment Station found that the tobacco bud-
worm, H. virescens (Fabricius), crossed with a closely
related species, H. subflexa Guenee, produces sterile
male progeny. Females, however, are fertile and can be
backcrossed with tobacco budworm males for an ap¬
parently infinite number of generations to produce more
sterile males and fertile females. This unique type of
genetic action is now under investigation at the U.S.

Agricultural Research Service’s Southern Field Crop In¬
sect Management Laboratory at Stoneville, Miss. Like¬
wise, many investigators, working on several moth
species, have shown that parent moths receiving sub¬
sterilizing dosages of irradiation produce progeny that
have higher levels of sterility than their parents and that
some adverse effects may persist for several generations.
Other promising methods of genetic engineering include
production of nondiapausing strains and conditional
lethal factors. Regardless of the genetic mechanism that
may be involved, however, the full potential of genetic
suppressive measures will not be realized until insects
that are reasonably competitive with their natural
counterparts can be mass-produced.

Even though interest in mass rearing has centered on
developing genetic-control methods, during the past 15
years entomologists have become increasingly interested
in more extensive and dependable use of biological-
control agents. To appraise both the merits and the limi¬
tations of natural biological-control organisms, I have
made extensive use of models that simulate codeveloping
parasite-host populations. While the accuracy of such ap¬
praisals can be limited, this effort yielded the following
conclusions: Self-perpetuating populations of many para¬
sites cannot consistently suppress their hosts below
damaging levels because of the actions and interactions
of several natural regulating forces: dependability of
these parasites can be greatly increased, however, by
augmenting their numbers continuously or at strategic
times in the host cycle; and, based on estimates of the
host-finding efficiency of certain parasites and on pro¬
jected costs for their mass production, using augmenta¬
tion offers outstanding potential for effectively managing
a wide range of major pests. Equally important, augmen¬
tation of natural predators and parasites should have lit¬
tle or no effect on the environment.

I believe the prospects are excellent that populations of
many of the world’s major agricultural insect pests and
disease vectors can be controlled on a regional or eco¬
system basis at levels below those that are economically
damaging: this control can be achieved if various sup¬
pression techniques that are already available or that can
be developed are properly integrated. In recent years, im¬
portant advances have been made on various methods of
suppression; these advances include more selective chem¬
icals, resistant crop varieties, and various types of attract-
ants that may be useful for suppression and may also
provide highly sensitive methods for population detection
and assessment. Biological organisms, including genet¬
ically altered insects, insect parasites, insect predators
and microbial agents, may be among the most practiced
and desirable components to use in systems for area-wide
population management. But the usefulness of biological

x

organisms for such purposes will depend not only on ef¬
ficient mass-production capabilities but also on the qual¬
ity of the organisms produced.

There is also an urgent need for research on the
ecological matters that are highly relevant to the concept
of regional management. Many entomologists have given
little consideration to this regional approach because the
research and operational problems involved in developing
and executing such a pest-management procedure are so
large. More definitive information is needed on numbers
of pest insects in natural or reduced populations, the rate
and extent of dispersal, mating behavior, and the dynam¬
ics of various pest populations. But getting enough
money for this necessary research is difficult. Yet I feel
that the alternatives should be critically analyzed.

Despite the availability of a wide variety of highly effec¬
tive insecticides, we know through several decades of ex¬
perience that a limited and uncoordinated attack on
segments of pest populations, farm to farm and crop to
crop, has had no major impact on the abundance of major
annually recurring pests. They are as numerous and pose
the same threat to production today as they did several
decades ago, especially such important pests as the boll
weevil; Heliothis complex; pink bollworm; codling moth,
Laspeyresia pomonella (Linnaeus); fall armyworm,
Spodoptera frugiperda (J. E. Smith); cabbage looper, Tri-
choplusia ni (Htibner); tropical fruit flies (Tephritidae);
sugarcane borer, Diatraea saccharalis (Fabricius); Euro¬
pean com borer, Ostrinia nubilalis (Htibner); and various

flies affecting livestock. And we still depend mostly on
ecologically disruptive insecticides for controlling these
pests when they reach damaging numbers. So I share the
opinion of more and more entomologists that there is
every justification for full exploration of the concept of
regional management. Methods of colonizing and mass
producing biological organisms of various kinds are likely
to be especially important in achieving this goal.

The success in suppressing a wide-ranging pest like the
screwworm, on a regional basis and with great economic
benefits, should not be regarded as exceptional. I feel
that the same opportunities exist for managing many
other major pests with suppression methods that would
eliminate crop losses and cost less than insecticides. And
techniques likely to be most useful in systems for manag¬
ing pest populations are generally pest specific and
should do little or no damage to the environment.
Therefore, conferences such as this are important and
necessary so that investigators engaged in research on in¬
sect colonization and mass production can report on prog¬
ress and problems that may be applicable to a wide range
of species.

E. F. Knipling,

Expert consultant in pest management,
Agricultural Research Service

xi

Recommendations of the Conference

Discussion during the conference revealed that much
critical research still must be done to achieve the ability
to mass-rear high-quality insects consistently. Section
leaders consulted with their colleagues during and after
the conference, and the following recommendations were
formulated. These recommendations can provide direction
to scientists and administrators for establishing focus
and priorities for future research on insect rearing.

Establishment and Maintenance
of Insect Colonies Through
Genetic Control

The maintenance of insect colonies has become necessary
and vital to modem pest-management strategies, includ¬
ing sterile-insect release, genetic control (such as hybrid
sterility, conditional lethal mutations, and translocation
sterility), and biological control (such as parasites, pred¬
ators, viruses, and bacteria). The establishment of col¬
onies and adaptation of insects to laboratory conditions
result in genetic changes with unknown effects on the
performance of the insect. Some effects of domestication
are probably unavoidable, but certain precautions can
minimize their impact. The following recommendations
are made to emphasize steps that could be taken to avoid
serious genetic alteration of colonized species and to sug¬
gest areas of research that should be supported.

1. How genetic selection affects performance of released
insects depends on their intended function. Neither
sterile-male releases that require insemination of wild
mates nor augmentative releases of natural enemies to
control extant populations would need the same genetic
diversity as a release to produce an in-field reproductive
population or as use of a colonized insect for viral or bac¬
terial production.

Recommendations: Support research to determine the
requirements for genetic diversity in laboratory-
reared insects used for various purposes. Determine
the environmental conditions that will select for the
best genotypic constitution. Then, develop and per¬
fect methods for maintaining these conditions, in¬
cluding ways to alter temperature and light cycles,
maintain humidity, control effects of nutrition on
selection and performance, and establish mating con¬
ditions that are as natural as possible.

2. Laboratory domestication, the restriction of popula¬
tion size, and use of artificial diets inevitably lead to dif¬
ferences in gene frequency (caused by genetic drift and
selection) between the native and colonized populations.

Therefore, there should be mechanisms to monitor such
changes, evaluate their effects, and compensate for any
that are harmful. Such criteria as isozyme variation, mor¬
phological mutations, chromosomal variation, behavioral
traits (including pheromone response, physical activity,
and reproductive processes), visual acuity, and physical
characteristics (such as body weight, wing length, bristle
number, temperature tolerance, and diapause response)
show genetic variation in insect populations.

Recommendations: Each program involving the use
of colonized insects should employ a scientist trained
in genetic techniques (chromosomal analysis, electro¬
phoresis, selection, genetic analysis, etc.) so that as
many genetic variables as possible may be measured
in the base population and during generations of
domestication. Parallel behavioral characteristics
need to be studied so that the relationship between
the genetic traits and performance of the released in¬
sect can be established. As observations on changes
in genetic variation and behavior accumulate, correla¬
tions can be made with changes in performance of do¬
mesticated populations. This information, in turn,
will increase our ability to control such changes and
predict insect performance.

3. Efficiency of laboratory -reared insects in applications
for population control is affected by at least three sig¬
nificant factors: genetic selection during domestication,
genetic changes in the native population resulting from
control procedures, and environmental and geographic
variation in the native population. Each of these prob¬
lems must be studied from the viewpoint of population
genetics.

Recommendations: Give high priority to expansion of
expertise in insect population genetics (either by em¬
ploying trained scientists or retraining existing per¬
sonnel). Support basic field studies on pest species
relating to their geographic variation and reproduc¬
tion mechanisms, resistance, and preferred host and
alternate host selection. Initiate and expand area wide
programs to measure the population dynamics of
target species.

Diets and Containerization
for Insect Rearing

1. Much progress has been made in formulating artificial
diets for rearing insects; but optimal diets need to be
developed for production of high-quality insects. An in¬
sect’s adaptability permits it to establish and reproduce

xiii

under laboratory conditions, but some genetic traits are
lost. Performance of these insects is also affected by the
interaction between genetic traits and the effects of nutri¬
tion and dietetics. The precise influence of each of these
on the insect is seldom determined. Also, many species
must still be reared on their host plants, and parasites
and predators usually require their host insect. Quality
and cost of nutrients are increasing in importance. And
we need to standardize, as much as possible, storage of
ingredients, ways to insure adequate nutritive composi¬
tion, diet preparation, and conduct of associated rearing
methods.

Recommendations: Improve and refine existing arti¬
ficial diets to meet optimal performance criteria for
insects reared in the laboratory. Develop artificial
diets for rearing representative species of beneficial
and pest insects. Support research to reduce the cost
of artificial diets, particularly for mass rearing. Alter¬
native, low-cost primary nutrient sources, especially
gelling and bulking agents, need to be evaluated and
used. Standardize diets and rearing procedures for
representative insects. Develop guidelines to insure
uniform quality of dietary ingredients, diet prep¬
aration, rearing procedures, and environmental
conditions.

2. Primary nutrients such as nonrefined protein (wheat
germ or cottonseed meal) and refined ingredients (casein,
vitamins, or agar) vary in quality because of contamina¬
tion with micro-organisms, improper storage, poor con¬
tainers, different suppliers, or other unknown factors.
Existing techniques for monitoring quality, measuring
microbial levels, etc., should be used consistently in rear¬
ing operations.

Recommendations: Develop specifications that define
the quality of diet ingredients. Establish procedures
to monitor or assay the quality of commonly used
dietary ingredients from various sources and deter¬
mine if there are significant deviations from specified
tolerances. Define and conduct research to determine
the optimal storage conditions and shelf life for diet
ingredients.

3. In each program for rearing insects, scientists develop
their own procedures for measuring and mixing diet in¬
gredients, preparing (heating) and dispensing diet, and
operating equipment. Studies to establish optimal condi¬
tions and equipment for a broad range of insects have
been inadequate.

Recommendation: Support research to determine op¬
timal methods and equipment for diet preparation to
avoid loss of nutrients and retain the desired physi¬
cal properties.

4. Insect-rearing containers are generally adapted from
commercially available cups or boxes designed for other
purposes. These are not always best suited to the insect,
are especially vulnerable to increased cost, and may be
discontinued or changed at any time. So an entire rearing
system may depend on suboptimal containers.

Recommendations: Develop specifications for im¬
proved insect-rearing containers. Evaluate the
relative merits of disposable and reusable containers.
Determine the costs and benefits of custom-made
containers.

Engineering for Insect Retiring

1. Efficient production and use of insects require engi¬
neering expertise. Joint research of an engineer and ento¬
mologist results in, for example, more efficient rearing
facilities, better environmental conditions for insects and
personnel, automation of rearing procedures and diet
handling (with major savings in cost of labor), and better
working conditions through control of insect and dietary
waste products. Those engineers with experience in these
areas have not been used effectively. And there have
been few efforts to provide them with training by ex¬
posure to existing insect facilities and programs.

Recommendations: Increase engineering support of
insect rearing by assigning additional engineers to
these programs. Provide engineers with training in
existing facilities so they will be familiar with the
many problems of rearing insects.

2. Facilities are often designed with limited knowledge of
the problems of insect rearing and of solutions that have
been devised by other engineers and entomologists.

Recommendation: Provide for consultation with
engineers and entomologists when designing or
remodeling insectaries.

3. Research in insect rearing has recognized and solved
many problems. Some of these are common to all insects,
some to certain orders, and others to certain species. Effi¬
cient use and extension of this information to all insect
rearing is needed.

Recommendations: Develop engineering standards
for the problems that are common to many insects
and for which solutions have been devised, such as
clean-air systems, environmental controls, backup
systems, traffic-flow patterns, and diet-handling
systems. Provide a standard terminology that will be
understood by both engineers and entomologists.

4. When rearing insects, one often encounters unusual

xiv

and unexpected environmental problems. For example,
maintenance of a proper level of humidity to prevent
microbial contamination in the diet and provision for a
temperature gradient from floor to ceiling may interfere
with uniformity of insect development.

Recommendation: Support research on the best en¬
vironmental conditions for rearing insects and on
methods to maintain these conditions in the facility.

Control of Pathogens
and Microbial Contaminants
in Insect Rearing

1. Widespread use of laboratory-reared insects in alter¬
nate methods of control and in support of basic and ap¬
plied research requires that they be of a consistently
good quality. Gross and lethal microbial contamination of
the insects and their diets have been recognized since the
beginning of insect rearing, and modem antimicrobial
agents now eliminate many of these problems. But sub-
lethal microbial infections are often not detected, and
there is evidence that these diseases may subtly affect
development, behavior, and reproduction. The antimicro¬
bial inhibitors may also affect insect performance.

Recommendations: Support research to devise
methods for detecting sublethal micro-organisms and
for determining how they affect the behavior of
laboratory-reared insects. Also, support research to
recognize how antimicrobial agents affect insects and
to discover new and more effective microbial in¬
hibitors. Assemble management guidelines for pre¬
venting and controlling contamination.

2. Information on diseases of laboratory-reared insects is
not widely circulated. Frequently, unknown diseases may
destroy a colony. Compilation of data on the sources,
prevention, and elimination of insect diseases is required.

Recommendations: Conduct periodic workshops to
familiarize those engaged in rearing insects with
information on disease recognition, diagnostic tech¬
niques, and therapeutic methods. Publish a manual
that would be a source of this information. Support
research to catalog and identify micro-organisms that
infect laboratory-reared insects. Discover effective
methods for prevention and control of diseases that
affect insects. These methods might include im¬
provements in the design of insect-rearing facilities
and control of micro-organisms in components of the
diet, or in the prepared diet, or in the insect and its
byproducts.

3. A colony of insects may be free of disease for a long
time and then suddenly become susceptible to and be

debilitated by organisms that were previously non-
pathogenic.

Recommendation: Determine factors in the environ¬
ment, diet, and breeding of insects that promote
biological vigor and reduce the incidence of diseases.

Production, Use, and Quality
Testing in Insect Rearing

1. Use of laboratory -reared insects for basic research and
evaluation of current and alternate methods of control
have created demand for a central source of expertise in
all phases of rearing. Expertise in engineering, pathology,
physiology, genetics, insect nutrition, etc., are required to
supply insects, solve rearing problems, and expedite de¬
velopment of new facilities and programs.

Recommendation: Determine the feasibility of
establishing a central insect-rearing research center.

2. Laboratory insects differ from their wild parent
population in behavioral, biological, and physiological
characteristics. But there are few tests of quality that
compare these characteristics in laboratory and wild
populations and even fewer that relate differences to the
performance or usefulness of the laboratory insects.

These data are essential for precise interpretation of
results of studies obtained with laboratory-reared insects.

Recommendations: Initiate an intensive research pro¬
gram to discover, evaluate, and define the biological,
behavioral, and physiological characteristics that de¬
scribe insect quality, especially those characteristics
of major insect pests. The scope of the program
should include: The identification of characteristics to
be measured; the development of laboratory tech¬
niques to assess insect quality, which can be directly
correlated with performance under natural condi¬
tions; and what threshold levels (physiological,
behavioral, ecological, physical, and genetic) to accept
and when and how to accept them.

3. Considering the worldwide multiplicity of uses for
various laboratory cultures, standardized tests must be
developed and studies must be conducted to characterize
established insect colonies reared with standard tech¬
niques. The availability of performance standards for
laboratory populations would promote communication
and cooperation between scientists working with the
same species and would enhance our understanding of
genetic, physiological, nutritional, and behavioral charac¬
teristics of economic pests.

Recommendations: Encourage and support the devel¬
opment of performance standards for well-

xv

established, standardized insect colonies. Promote
the use of performance standards in evaluating insect
consistency in standard tests of toxicity and activity.

4. Continuing increases in the use of laboratory -reared in¬
sects for research and methods development necessitates
closer communication between the insectary (producer)
and scientist (user). Routinely, producers supply insects
as a service, but they usually do not convey changes in
rearing procedures (such as dietery substitutes and
changes in environmental regime). Generally, users are
unaware of the effects such changes may have on their
research, so they do not request such information. And
the user seldom tells the producer about any changes
that may have been observed.

Recommendations: Support and encourage the de¬
velopment of procedures for the exchange of ap¬
propriate information between the insect producer
and the user. Scientists using laboratory-reared in¬
sects should be encouraged to test for and document
insect quality as a prerequisite to the conduct of
research.

5. The practical feasibility of using biological-control
agents (chiefly insects, mites, and diseases) for control of
many insect pests has been amply demonstrated. But
large-scale rearing and use of many potentially effective
entomophagous arthropods is prohibited by the Cost and
difficulty of rearing both host and agent. A concentrated
effort is needed to develop artificial diets and in vitro
rearing systems for entomophages.

Recommendations: Give high priority to the inves¬
tigation and development of laboratory techniques
for the in vitro production of entomophages. Studies
and evaluation processes necessary for determining
the competitiveness of laboratory-reared species
should also be done.

Management

of Insect-Rearing Systems

1. Rearing of biologically fit insects has become increas¬
ingly sophisticated and therefore requires local knowledge
in areas such as the nutritional requirements of insects,
disease recognition and prevention, and operation of auto¬
mated equipment. Finding people with experience in
these areas of insect rearing is difficult: most are trained
on the job.

Recommendation: Recognize the need for skilled per¬
sons to rear insects, and encourage universities to
develop formal courses that would prepare individ¬
uals for careers in this field. Such a program would
have a broad curriculum in entomology, nutrition, in¬
sect behavior, ecology, pathology, and physiology.

2. Large-scale programs for rearing insects require exten¬
sive management if the products are to be suitable for
their intended application. The facility must be main¬
tained, ample supplies made available, the required
number of insects produced on schedule, and personnel
trained. And the results of research that improves rearing
techniques must be integrated into existing programs.

Recommendations: Recognize that large-scale rearing
programs require professional managers, and develop
criteria and requirements for such positions. Encour¬
age universities to develop curriculums to prepare in¬
dividuals for careers in insect-rearing management by
providing up-to-date advice about the needs of these
positions. In the interim, plan workshops for man¬
agers of insect-rearing facilities to discuss mutual
problems and their solutions.

3. The increased number of rearing facilities and greater
quantities of insects being produced have exposed more
people to potential health hazards associated with ar¬
thropods and their byproducts. These hazards may be
either physical discomfort or some type of allergic reac¬
tion. This problem is only beginning to be recognized.

Recommendations: Identify and catalog specific
health hazards associated with arthropods. Develop
procedures and equipment to protect workers, and
improve insectary designs and environmental
controls.

4. Optimization of insect rearing requires the capability
of retrieving and analyzing extensive data on many
variables.

Recommendation: Implement available technology
for systems analysis to facilitate the collection,
analysis, and recall of essential data in programs for
insect production and use.

xvi

Section 1

Establishment and Maintenance

of Insect Colonies Through Genetic Control

The science and art of insect rearing has increased in
complexity and sophistication as the need has grown for
insects in entomological research and pest control. Early
entomologists could perform elegant studies using insects
collected from the field; or they could rear enough insects
on natural hosts. But new concepts, such as the sterile-
insect method or the mass production of parasites and
predators, require many more insects. So artificial diets
and environments have been developed. These new tech¬
niques are so successful that even further demands have
been made on the insect-rearing specialist. Not only do
entomologists want an abundant supply, but they insist
that the insects function normally when released into
their natural environment.

Unfortunately, in some cases the released insects do not
function normally. Explanations for these malfunctions
have been sought, and two biologically logical culprits
found: pathogenic contamination and genetic deteriora¬
tion. Microscopic examination will confirm or deny the
presence of pathogenic organisms in insect cultures
quickly; though eliminating the pathogens is not easy.
But the causes of genetic deterioration are not so readily
identified. Only detailed genetic studies covering two or
more generations can determine the presence or absence
of detrimental genetic traits and then only if such traits
can be defined and measured. Even so, many authors
have suggested remedial measures for assumed genetic
deterioration.

I believe an unappreciated element of this problem is that
the genetic changes taking place when an insect colony is
started are natural ones that occur whenever any bio¬
logical organism goes from one environment to another.
These processes have been well studied as evolutionary
events and involve such concepts as colonization, selec¬
tion, genetic drift, effective population numbers, migra¬
tion, genetic revolutions, and domestication theory.

In this section of the conference, we have tried to inte¬
grate some of these genetic concepts into our pool of
knowledge on insect rearing. First, I discuss what hap¬
pens to genetic variability in the process of domestica¬
tion. What factors might change it, and which ones might
be expected to have little or no effect? Then, assuming
that we can define the type of insect we want, how can
we change an insect population to conform to our expec¬
tations? We can use artificial selection; Anita Collins
reviews this well-documented field. Finally, Dennis
Joslyn discusses various strategies for maintaining ge¬
netic variability in reared insects.

Our conclusions in this section are encouraging even
though changes in genetic variability during domestica¬
tion of an insect population will be unavoidable. We must
accept the inevitability of these changes if we want to
rear insects in artificial environments. But, within budg¬
etary restraints and the limitations dictated by the in¬
tended uses of the domesticated strains, those genetic
changes need not ruin the program. We do, however, need
to carefully study the type of variation present in the
base populations and monitor changes that occur over
time in the laboratory. For example, we can measure
isozyme variation, chromosomal variation, visible mark¬
ers, behavioral variants, body weight and size, reproduc¬
tive traits, visual acuity, diapause response, and various
types of activity. If we observe genetic changes over
time, and such changes appear to be correlated with
reduced performance of the laboratory strain, then cor¬
rection can be made either by modifying procedures or by
increasing the genetic variability of the colony. These cor¬
rections will require compromises between demands of
the rearing program and the availability of appropriate
genetic techniques. Such compromises are possible only
if the insect-rearing specialist and the insect geneticist
understand each other’s problems and needs.

Alan C. Bartlett,

Research geneticist,
Agricultural Research Service

1

Genetic Changes During Insect Domestication

By Alan C. Bartlett1

Introduction

The development of artificial diets and mass-rearing tech¬
niques for insects (see Chambers 1977 for a definition of
mass rearing) has progressed rapidly during the past 20
years. Most advances have been empirical; success has
been measured in numbers of insects available for release
per unit of cost (Gast 1968). But, having been successful
in quantity, researchers are now examining insect quality
(Huettel 1976, Chambers 1977), behavior (Boiler 1972),
and genetics (Mackauer 1973, McDonald 1976). This ex¬
amination is continuing as we become more confident of
our ability to mass-produce insects, and we now desire to
have success measured in quality as well as quantity per
unit of cost. Since most entomologists accept the need for
mass-reared insects for the implementation of many
insect-control techniques (see Knipling 1966 for an early
discussion of the uses for mass-reared insects), the bio¬
logical effects of laboratory production of insects must be
examined in detail. This paper explores some of the
genetic changes that may take place in the first genera¬
tions of an insect population during its establishment in
the laboratory.

The process of establishing and maintaining insect col¬
onies has been reviewed by several investigators (for ex¬
ample, Boiler 1972, 1979; Mackauer 1972; Davidson 1974;
Hoy 1976; Huettel 1976; and McDonald 1976). In general,
they have considered the process as a colonizing episode
for the species, as a process of domestication, or as an ex¬
pression of the founder principle of species evolution.
These concepts are interrelated and their subtle distinc¬
tions need not be pointed out here. Instead, I will limit
this discussion to the process of domestication, because it
is the only concept that requires human participation. I
will consider only those species that reproduce sexually.
And I will not consider stocks that have been subjected
to extreme selection pressures and used for very
specialized purposes.

'Research geneticist, Western Cotton Research Laboratory, Ag¬
ricultural Research Service, U.S. Department of Agriculture,
Phoenix, Ariz. 85040. In cooperation with the Arizona Agri¬
cultural Experiment Station.

Genetic Changes
Changes in variability

Many discussions of the processes that take place during
the establishment of an insect colony maintain that we
change the population’s level of genetic variability by
forcing insects through a bottleneck (see, for example,
Boiler 1972, 1979). Spurway (1955), in describing the
forces that can change gene frequency during domesti¬
cation of a species, called this process a winnowing, an
analogy that is more accurate than the bottleneck com¬
parison. The idea of a restrictive bottleneck that a strain
must go through on its way to the laboratory does not
necessarily conjure up a sense of sifting and sorting of
suitable genotypes of an inanimate environment. But win¬
nowing is just such a sifting and sorting. One can visual¬
ize the force of selection blowing through the falling in¬
sects and removing the chaff of unsuited genotypes; also,
some suitable genotypes will bounce off the winnowing
board and be lost from the population (fig. 1).

Figure 1.— Winnowing insects for laboratory
domestication.

2

Understanding the force of selection on existing geno¬
types and the random loss of genotypes is vital to any
discussion of changes that take place in the domestica¬
tion of a species. We have learned much about the pro¬
cesses of natural selection through studies of artificial
selection (see “Artificial Selection of Desired Character¬
istics,” by Anita M. Collins). Here, I will emphasize how
natural selection can change a laboratory population’s
genetic variability. Lemer (1958) listed and discussed
some factors that should be considered in an experiment
in artificial selection. With some minor modifications, his
list is given below as an outline of changes that a field-
collected strain may undergo when introduced into the
laboratory.

1. Laboratory and experimental populations are usually
maintained in reasonably constant environments that
are drastically different from those where earlier
selection for stable developmental patterns has oc¬
curred. They are shielded from environmental fluc¬
tuations of various kinds and from predators; they
are provided with shelter and food. In nature, selec¬
tion may operate much more strongly in favor of
individuals able to overcome unexpected stresses.

A constant environment does not merely change the
criteria that determine fitness; it may also greatly
modify the whole genetic system of an artificially
colonized population.

2. Interspecific competition is part of natural selection
but is seldom involved in artificial colonization. Clear¬
ly, expensive rearing supplies should not be used for
rearing competitors or parasites to produce interspeci¬
fic competition in the laboratory; but we should also
realize that genetic variability may change without
this factor.

3. Members of natural populations can choose their
own environments from those available. For domesti¬
cated organisms, microenvironmental conditions are
generally made suitable for the average, or sometimes
the poorest, genotype. Within the population, all the
individuals are generally confined to the same environ¬
ment.

Laboratory conditions will also cause other changes not
directly related to the environment:

1. Density-dependent behaviors may be profoundly
affected in laboratory conditions. For instance, mate¬
searching behavior is probably restricted in the small
mating cages usually provided in the laboratory. Fe¬
male egg-laying behavior probably changes when few
deposition sites are provided. Apparently, the more
finicky the species, the more behavioral changes will
be necessary to adapt to the new conditions.

2. Mate-selection processes may be changed because
unmated or previously mated females will have re¬
stricted means of escape. Promiscuous sexual be¬
havior is an advantage in domesticated species, as is
the ability or willingness to breed in confinement.

3. Dispersal characteristics, specifically adult flight re¬
sponse and larval dispersal, may be severely restricted
by laboratory rearing conditions (Bush et al. 1976).

Kinds of change

Huettel (1976) discussed many of these possible changes
in laboratory populations and suggested methods for
monitoring and compensating for them. Some of these
factors should also be considered in predicting the kinds
and amount of changes an insect colony may experience.
The first factor is the amount of variability present in the
population. If a trait does not vary, then no genetic
change is possible without mutation. But past experience
suggests that most measurable traits are variable. In
fact, recent work has disclosed an extraordinary amount
of variation in native populations even in marginal areas
(Prakash 1973); and, in inbred populations, high amounts
can remain (Yamazaki 1972). So we may conclude that
variability for many, or even most, important traits will
be present in stocks collected from the wild.

What will happen to that variability in the process of
domestication? We can take only a small part of the
whole population to start our culture, which will now be a
closed population. In an open population, the available
genetic variation is both greater and of a different kind
because of gene migration and environmental diversity in
both time and space (Mayr 1970). In a closed population,
the common alleles originally present will be represented,
while rare alleles are likely to be missing. All genetic
changes will be made from the limited genetic variation
present in the original founders.

The number of colonizing insects will directly affect how
much variation will be taken from the native gene pool.
The lower the number of insects, the lower the number of
represented alleles and consequently the greater the
deviation of the mean phenotype of the colonized popula¬
tion from the mean phenotype of the parental population.
In an interesting experiment reported by Dobzhansky
and Pavlovsky (1957), in 17 months, lines founded with
20 individuals diverged from each other and from the
original population far more than lines founded with
2,000 individuals. But even those lines founded by 2,000
individuals diverged from each other by as much as 50%.
Powell and Richmond (1974) observed similar results for
populations started with 6 and 300 chromosomes. The
forces of natural selection imposed on a recently isolated
gene pool may also cause significant changes in fitness

3

Extraction of laboratory colony

TIME (Generations)

Figure 2.— A conceptualization of the changes in genetic variability that may take place during domes¬
tication of an insect species.

and produce what Mayr (1970) has called a genetic
revolution. Such changes are illustrated in figure 2. In a
laboratory environment, genes that have been at an ad¬
vantage may now have some disadvantages (or, converse¬
ly, disadvantages may become advantages). For example,
in northern latitudes the ability to diapause is useful to
many insect species, while fn the laboratory it may be
neutral or even harmful. Similarly, behavior such as
‘‘startle response” or the ability to react to low concen¬
trations of pheromone may change from adaptive to non-
adaptive.

Lopez-Fanjul and Hill (1973a, 1973b) have described a
series of experiments illustrating the patterns of genetic
variability in laboratory populations. Each of three lines
of Drosophila melanogaster Meigen that had been
laboratory-reared for many years and one that was newly
colonized was crossed to a strain that had been selected
for high sternoplural-bristle number. Artificial selection
was then practiced on the resulting hybrid lines. The
three long-term laboratory strains were segregating for
two similar alleles at loci controlling sternoplural-bristle
number and for seven loci controlling enzymes. And the

4

newly isolated population contained some variation not
present among those in the laboratory. But Lopez-Fanjul
and Hill concluded that all the strains probably re¬
sponded similarly to maintenance in the cages.

Such results support the theory that similar environ¬
mental conditions will select for similar genetic responses
if not for identical genetic constitutions. After the trans¬
fer of a population from the field to the laboratory, a few
individuals will be able to produce more offspring no mat¬
ter how able they were in the field. Existing genic-balance
systems that maintain homeostasis in the field may not
confer fitness in the laboratory; so new balanced gene
systems will be selected. If conditions remain the same in
the laboratory, any later introduction of native genes will
be subjected to the same process of genetic revolution;
balanced gene complexes will evolve that will act the
same as those previously selected and may, in fact, be
the same (Lopez-Fanjul and Hill 1973b).

Introducing native genes

to laboratory colonies

It has been suggested that a cure for genetic differentia¬
tion of laboratory colonies from native populations is
regular introduction of native individuals. But there is a
danger that genetic differentiation of the colony because
of adaptation to laboratory conditions may lead to genet¬
ic isolation of native and laboratory populations. Oliver
(1972), studying incompatibility in intraspecific crosses of
Lepidoptera, found that geographically isolated races
generally show heterotic vigor in the development of
hybrids and a decline in fertility, skewing of sex ratio,
and developmental abnormalities in backcross and F2
progenies. Other studies, such as Jaenson (1978) and
Jansson (1978), have found that the strength of genetic
incompatibilities seems to be proportional to the differ¬
ences between environments where the races occur. So
there is a positive correlation between geographic dis¬
tance and incompatibility of races. A similar positive cor¬
relation exists between the length of time two populations
have been separated and their incompatibility. Introduc¬
tion of native genes into a laboratory population should
be regular— every one or two generations; it should not be
delayed until problems become apparent (Richerson and
Cameron 1974, Raulston et al. 1976, Sharp 1976). One
hazard of introducing native insects into a colony is that
they may never adapt and may therefore fail to produce
enough progeny to continue the strain. Also, parasites or
pathogenic micro-organisms could be introduced with the
field-collected insects.

We can measure the effect of introducing native genes in¬
to a laboratory population by the simple formula for one
locus (from Pirchner 1969) that

a p=m(PN-PL),

where a p= change in gene frequency for a given allele at
a given locus,

m— ratio of introduced individuals to individuals
in the laboratory population,

PN = frequency of the allele in the laboratory popu¬
lation,

and PL= frequency of the allele in the laboratory
population.

So the change in gene frequency will depend on the pro¬
portion of immigrant to native alleles and on the differ¬
ences in gene frequencies between the two populations. If
PN=Pl> then no change in gene frequency will take place
no matter how many native individuals are introduced. If
the frequencies in the two populations are different, then
the direction of change will be towards the highest gene
frequency. For example, if selection has operated on the
frequency of an allele to increase it over that in the
native population, then the change will be in the direction
of the laboratory strain. So the forces of selection and of
immigration of native alleles will be antagonistic. The
amount and direction of change will depend on the
relative strengths of the two forces. If laboratory condi¬
tions remain constant, selection will reestablish the orig¬
inal gene frequency once native alleles are no longer in¬
serted into the colony.

Changes related to the size
of the breeding population

The genotypic structure of populations can also be af¬
fected by their breeding system. When mating partners
are related to each other more closely than randomly
picked individuals, the mating system is called inbreed¬
ing; outbreeding occurs when they are less closely re¬
lated. Inbreeding implies mating of relatives and the pro¬
duction of progeny more homozygous than would be ex¬
pected with random mating (panmixis). These homo¬
zygous individuals often exhibit harmful traits. The pro¬
portion of heterozygous individuals is decreased by 2 pqF
each generation (where p and q are the frequencies of
alleles at a given locus and F is the inbreeding coefficient
of the population). The inbreeding coefficient is directly
related to the size of the breeding population. Therefore,
the change in the inbreeding coefficient over one genera¬
tion is estimated as

a F=Nm+Nfl8NmNf>

where Nm = the effective number of breeding males,
and 7^= the effective number of breeding females.

If any type of selection, artificial or natural, is present
during the reproductive phase of a population, then the

5

rate of inbreeding will increase; that is, the population
may be propagated by only a few individuals. The fre¬
quency of genes identical by descent will be increased in
those families most favored by the selection process. The
process of inbreeding alone will not change gene frequen¬
cies, only genotypic frequencies. But inbreeding plus
selection, a condition often encountered in the domestica¬
tion process, will have definite and rapid effects.

Measuring genetic changes

Electrophoresis could be used to measure changes in
genotypes and gene frequencies against theoretic changes
expected from combined selection and inbreeding. Fal¬
coner (1960) defined the panmictic (random mating) index
(. P ), which complements the inbreeding coefficient
(P=l—F), as the ratio of the frequency of heterozygotes
at any time to the frequency of heterozygotes in the base
population. So, if we could measure the frequency of
heterozygotes for a pair of alleles in the base population
(H0) and of heterozygotes at that same locus at any one
time, then the panmictic index for that locus at time (t)
would be

P=HtIH,.

If we could measure heterozygosity at many isozyme loci,
we could calculate the average heterozygosity over all ex¬
amined loci (n) as

pt=iHt/iH0.

1=1 1=1

The inbreeding coefficient would be Ft=\— pr How
changes in gene frequency at the enzyme loci are related
to changes at other loci (those affecting reproductive
fitness, competitiveness, etc.) is currently being studied
with electrophoresis. Whatever the outcome of these
studies, it is already obvious that inbreeding and selec¬
tion acting on the whole genome should be correlated
with changes at specific isozyme loci.

Changes in reproductive

ability

The most discussed effect of inbreeding is the reduction
in reproductive performance of the inbred population.
Changes in certain types of performance have also been
suspected as a consequence of inbreeding depression by
Davidson (1974) and Robinson (1977). Craig (1964) and
Crystal (1967) show that this potential inbreeding can be
counteracted with outbreeding, which will produce new
variability and heterosis. Generally, native genetic ma¬
terial is incorporated into the laboratory-reared popu¬
lation before release, although Craig (1964) advocated
keeping two colonies and crossing them. In fact, Young

et al. (1975) used this method effectively for the corn ear-
worm, Heliothis zea (Boddie), when the first generation of
laboratory insects was crossed with the native population.

Outbreeding seems to offer promise for improvement of
laboratory populations, but theory suggests that only
small increases in genetic variation could be expected
(Falconer 1960). Heterosis depends, among other things,
on differences in frequencies of genes that affect the
traits in question. If two populations adapted to different
environments are crossed, the outbred individuals may be
adapted to neither of the original environments. So, when
we consider using outbreeding for colony improvement,
we must carefully choose the strains to be crossed. And
such a technique may not be successful in every case.

Conclusion

An insect species that is taken from one set of environ¬
mental conditions and placed into another cannot survive
unless the population can adapt to the new environment.
Preadaptation or prospective adaptations can be refined
or extended by imposed selection pressures. But, at first,
existing adaptabilities enable the transition (Hansen
1977). This transition is a winnowing process with change
and selection producing a domesticated strain. Such a
strain must be adapted to the laboratory but may or may
not be well adapted to the original field conditions.

During domestication, we should minimize the important
changes and maximize reproduction. Since those changes
that affect interaction of domesticated and native strains
are important in most experiments and in control pro¬
grams, the species’ biology and behavior must be studied
well (see Boiler 1979 and Bush 1979 for further discus¬
sion of the types of biological studies that can be made).
Given enough information on the biology and behavior of
the species, we can adjust the laboratory environment to
compensate for many of these important genetic changes.
We may have to adjust our rearing procedures or our ex¬
pectations of the mass-rearing colony, but such compro¬
mises should be based on known biological factors, not on
conjecture. Within the limits imposed by budget, facility,
and manpower, the following criteria should be con¬
sidered when appropriate to the species: (1) Recognize
that the effective number of parents will be much lower
than the number of founder individuals, and try to com¬
pensate by starting with large founding populations
(LaChance 1979); (2) compensate for density-dependent
phenomena by using large mating cages, airflow to
remove pheromone accumulations, more egg-laying sites
than are needed, female hiding places, induced flight
behavior, etc. (McDonald 1976); (3) adjust or maintain
rearing densities to produce a proper balance of compe¬
tition but not overcrowding (Sokal and Sullivan 1963,
Peters and Barbosa 1977); (4) set environmental condi-

6

tions for the best, not the worst or average, genotype,
and use fluctuating temperatures and photoperiods in all
phases of the rearing environment; (5) maintain separate
laboratory strains under unique conditions and cross
these systematically to increase Fl variability (Craig
1964, Roberts 1974); (6) measure frequencies of biochem¬
ical and morphological markers in founder populations,
and monitor changes in gene frequencies over time (Bush
and Neck 1976, Novy 1978, Bush 1979); (7) develop mor¬
phological and biochemical genetic markers for popula¬
tion studies and for marking the released strain so that a
genetic-control technique can be evaluated effectively
(Bartlett 1967, 1982; Huettel 1979); and (8) determine the
standards that apply to the intended use of the insects,
and then adapt rearing procedures to maximize those
values in the domesticated strain (Boiler 1979).

Acknowledgment

I wish to thank Hollis M. Flint of the Western Cotton
Research Laboratory, Phoenix, Ariz., for his kindness in
providing the original drawing of figure 1.

References

Bartlett, A. C.

1967. Genetic markers in the boll weevil. J. Hered.
58: 159-163.

1982. Genetic markers: discovery and use in insect
population dynamics studies and control pro¬
grammes. In Sterile Insect Technique and Ra¬
diation in Insect Control, I.A.E.A. Publ. SM-
255, pp. 451-465.

Boiler, E. F.

1972. Behavioral aspects of mass-rearing of insects.
Entomophaga 17; 9-25.

1979. Behavioral aspects of quality in insectary pro¬
duction. In M. A. Hoy and J. J. McKelvey
(eds.), Genetics in Relation to Insect Manage¬
ment, pp. 153-160. Rockefeller Foundation.

Bush, G. L.

1979. Ecological genetics and quality control. In

M. A. Hoy and J. J. McKelvey (eds.), Genetics
in Relation to Insect Management, pp. 145-152.
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Bush, G. L., and Neck, R. W.

1976. Ecological genetics of the screwworm fly, Coch-
liomyia hominivorax (Diptera: Callephoridae)
and its bearing on the quality control of mass-
reared insects. Environ. Entomol. 5: 821-826.
Bush, G. L.; Neck, R. W.; and Kitto, G. B.

1976. Screwworm eradication: inadvertant selection
for noncompetitive ecotypes during mass rear¬
ing. Science 193: 491-493.

Chambers, D. L.

1977. Quality control in mass rearing. Annu. Rev.

Entomol. 22: 289-308.

Craig, G. B., Jr.

1964. Applications of genetic technology to mosquito
rearing. Bull. W.H.O. 31: 469-473.

Crystal, M. M.

1967. Reproductive behavior of laboratory-reared
screwworm flies. J. Med. Entomol. 4: 443-450.

Davidson, G.

1974. Genetic control of insect pests. 158 pp. Aca¬
demic Press, New York.

Dobzhansky, T., and Pavlovsky, O. A.

1957. Indeterminate outcome of certain experiments
on Drosophila populations. Evolution 7: 198-
210.

Falconer, D. S.

1960. Introduction to quantitative genetics. 365 pp.
Ronald Press Co., New York.

Gast, R. T.

1968. Mass rearing of insects: its concepts, methods,
and problems. I.A.E.A. Publ. STI/PUB/185,
pp. 59-67.

Hansen, E. D.

1977. The origin and early evolution of animals. 670
pp. Wesleyan University Press, Middletown,
Conn.

Hoy, M. A.

1976. Genetic improvement of insects: fact or fan¬
tasy. Environ. Entomol. 5: 833-839.

Heuttel, M. D.

1976. Monitoring the quality of laboratory-reared

insects: a biological and behavioral perspective.
Environ. Entomol. 5: 807-814.

1979. Genetic approaches to basic problems in insect
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McKelvey (eds.), Genetics in Relation to Insect
Management, pp. 161-169. Rockfeller
Foundation.

Jaenson, T. G. T.

1978. Mating behavior of Glossina pallidipes Austen
(Diptera, Glossinidae): genetic differences in
copulation time between allopatric populations.
Ent. Exp. Appl. 24: 100-108.

Jansson, A.

1978. Viability of progeny in experimental crosses
between geographically isolated populations of
Arctocorisa carinata (C. Sahlberg) (Heterop-
tera, Corixidae). Ann. Zool. Fennici 15: 77-83.

Knipling, E. F.

1966. Introduction. In Carroll N. Smith (ed.), Insect
Colonization and Mass Production, pp. 2-12.
Academic Press, New York.

LaChance, L. E.

1979. Genetic strategies affecting the success and
economy of the sterile insect release method.
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ics in Relation to Insect Management, pp. 8-18.

7

Rockefeller Foundation.

Lemer, I.

1958. The genetic basis of selection. 298 pp. John
Wiley and Sons, New York.

Lopez-Fanjul, C., and Hill, W. G.

1973a. Genetic differences between populations of
Drosophila melanogaster for a quantitative
trait. I. Laboratory populations. Genet. Res.
22: 51-68.

1973b. Genetic differences between populations of
Drosophila melanogaster for a quantitative
trait. II. Wild and laboratory populations.
Genet. Res. 22: 69-78.

McDonald, I. C.

1976. Ecological genetics and sampling of insect
populations for laboratory colonization. En¬
viron. Entomol. 5: 815-820.

Mackauer, M.

1972. Genetic aspects of insect production. Ento-
mophaga 17: 27-48.

Mayr, E.

1970. Populations, species, and evolution. 493 pp.

Harvard University Press, Cambridge, Mass.

Novy, J. E.

1978. Operation of a screwworm eradication program.
In R. H. Richardson (ed.), The Screwworm
Problem, pp. 19-36. University of Texas Press,
Austin.

Oliver, C. G.

1972. Genetic and phenotypic differentiation and geo¬
graphic distance in four species of Lepidoptera.
Evolution 26: 221-241.

Peters, T. M., and Barbosa, P.

1977. Influence of population density on size, fe¬
cundity, and development rate of insects in
culture. Annu. Rev. Entomol. 22: 431-450.

Pirchner, F.

1969. Population genetics and animal breeding. 274
pp. W. H. Freeman and Co., San Francisco.

Powell, J. R., and Richmond, R. C.

1974. Founder effects and linkage disequilibria in ex¬
perimental populations of Drosophila. Proc.
Nat. Acad. Sci. U.S.A. 71: 1663-1665.

Prakash, S.

1973. Patterns of gene variation in central and mar¬
ginal populations of Drosophila robusta. Genet¬
ics 75: 347-369.

Raulston, J. R.; Graham, H. M.; Lingren, P. D.; and Snow,
J. W.

1976. Mating interaction of native and laboratory-
reared tobacco budworms released in the field.
Environ. Entomol. 5: 195-198.

Richerson, V., and Cameron, E. A.

1974. Differences in pheromone release and sexual
behavior between laboratory-reared and wild
gypsy moth adults. Environ. Entomol. 3:
475-481.

Roberts, W. C.

1974. A standard stock of honeybees. J. Apic. Res.
13: 113-120.

Robinson, A. S.

1977. Genetic control of Hylemya antiqua. II. Can in-
breeding depression be a serious obstacle to the
development of homozygous rearrangement
lines? Entomol. Exp. Appl. 21: 207-216.

Sharp, J. L.

1976. Comparison of flight ability of wild-type and
laboratory-reared Carribbean fruit flies on a
flight mill. J. Ga. Entomol. Soc. 11: 255-258.

Sokal, R. R., and Sullivan, R. L.

1963. Competition between mutant and wild-type

housefly strains at varying densities. Ecology
44: 314-322.

Spurway, H.

1955. The causes of domestication: an attempt to
integrate some ideas of Konrad Lorenz with
evolution theory. J. Genet. 53: 325-362.

Yamazaki, T.

1972. Detection of single gene effect by inbreeding.
Nature (London) New Biol. 240 (97): 53-54.

Young, J. R.; Snow, J. W.; Hamm, J. J.; Perkins, W. D.; and
Haile, D. G.

1975. Increasing the competitiveness of laboratory-
reared com earworm by incorporation of indig¬
enous moths from the area of sterile release.
Ann. Entomol. Soc. Am. 68: 40-42.

8

Artificial Selection

of Desired Characteristics in Insects

By Anita M. Collins1

Introduction

Artificial selection is the alteration over time of pheno¬
typic characteristics of a population of organisms through
the intervention of man. Several major texts deal gener¬
ally with theoretical and actual application of the science
of genetics to artificial selection (see Lush 1945, Mather
1949, Falconer 1960, Li 1968, and Crow and Kimura
1970). Articles on artificial selection for insects include
many on Drosophila spp. (see Robertson and Reeve 1957,
Clayton et al. 1957, Madalena and Robertson 1975, and
Frankham 1977) and on Triholium spp. (see Wilson et al.
1965, Enfield et al. 1969, and Berger and Freeman 1974).
Reviews of the application of artificial selection to insect
populations (Bell 1963, Mackauer 1972, and Hoy 1976)
have all held that it has great potential. Artificial selec¬
tion has been used successfully for some time on two im¬
portant domesticated insects— the silkworm, Bombyx
spp. (Aizawa et al. 1961, Yokoyama 1979), and the honey
bee, Apis mellifera Linnaeus (Rothenbuhler 1958, 1979;
Rothenbuhler et al. 1968; Kerr 1974; Cale and Rothen¬
buhler 1975; and Goncalves and Stort 1978). Attempts
have also been made to improve various insect parasites
for use in pest control (Wilkes 1947; DeBach 1958, 1964;
Hoy 1979; and Roush 1979).

A selection program must begin with basic information
about the organism and the traits to be selected. Some
understanding of the creature’s reproductive strategies is
necessary, including such things as how many males
mate with each female, whether a male can mate more
than once, how many young are produced at one time,
and what interrelationships exist among those offspring.
Theories about artificial selection in animals deal mainly
with mammalian-type systems— a single mating (one male
X one female), few offspring at one time, diploid inheri¬
tance in both parents, etc. The theories must be modified,
then, for insects like the social, multiple-mating, haplo-
diploid honey bee. In any case, one must have a clear
idea of what characters are to be selected and exactly
how they will be measured. Knowledge of how the trait is
inherited— whether it is controlled by one, a few, or many
genes, will profoundly affect how it will be selected. Esti-

1 Research geneticist, Bee Breeding and Bee Stock Research Unit,
Agricultural Research Service, U.S. Department of Agriculture,
Route 3, Box 82-B, Ben Hur Road, Baton Rouge, La. 70808. In
cooperation with the Louisiana Agricultural Experiment Station.

mates of a genetic parameter called heritability will pre¬
dict whether the trait will respond to selection, by how
much, and how fast. Given this basic biological informa¬
tion, one can choose a breeding plan and its selection cri¬
teria intelligently.

Desired Characteristics
Measurement

Many characters have been modified by selection. Mor¬
phological traits such as body weight (Enfield 1972) and
thorax or wing length (Robertson and Reeve 1952) may
be measured easily. Time of pupation (Englert and Bell
1970), temperature adaptation (White et al. 1970), disease
resistance (Aizawa et al. 1961), DDT2 resistance (King
1954, Robertson 1957), and sex ratio (Simmonds 1947), all
physiological traits, may require more complex assess¬
ment. Evaluating behavioral traits such as host prefer¬
ence (Allen 1954), pollen collection (Mackensen and Nye
1966), geotaxis (Erlenmeyer-Kimling et al. 1962), photo¬
taxis (Choo 1975a), dispersal (Ogden 1970), walking (Choo
1975c), and mating behavior (Manning 1968, Eoff 1977)
may also be rather complicated. The special aspects of
behavior genetics have been considered by Hirsch (1967),
Dobzhansky (1972), McCleam (1973), Ehrman and Par¬
sons (1976), and Fuller and Thompson (1978); and Boiler
(1979) discusses the special application of behavior ge¬
netics to insect rearing.

Some traits may be measurable only on certain individ¬
uals of a population or at certain times. For example,
some traits such as egg-laying capacity, may be sex
limited, or they may simply be expressed differently in
each sex (Enfield et al. 1975). Other traits may require
the organism’s death before measurements can be made;
so evaluation must be based on its closest relatives. In
these cases, tests are conducted with progeny or sibs— in¬
dividuals from the same litter or egg hatch of the same
parents— leaving some alive and sacrificing others for
measurement purposes. A familiar example of this form
of study exists in the daily industry, where bulls are
evaluated on the basis of their offspring from many dams
and used for more intensive breeding only if they are
selected as sires. In insects, which usually have distinct

21 , 1 , 1 -Trichloro-2,2-bis(p-chlorophenyl)ethane.

9

life stages or castes, a particular character may have to
be measured during one stage (Tucker 1980). The sting¬
ing behavior in worker honey bees, for instance, is not
expressed by queens and drones. And finally, many be¬
haviors in social insects reflect the activity of a group
rather than of an individual. Honey production is an ac¬
tivity carried out by an entire honey bee colony; so the
colony becomes the experimental organism (Rothenbuhler
1960).

Mode of inheritance

Once a technique for measuring a particular trait has
been devised, understanding how the trait is inherited
may be useful. The two major categories of traits are dis¬
crete, or qualitative, and continuous, or quantitative.

With discrete traits, such as number of hairs and eye and
body color, there may be one or only a few major genes
controlling its expression. In these instances, relatively
simple relationships exist among the several states of the
trait (for example, brown eye color— the wild type— is
dominant over the mutant scarlet). The dominant brown
phenotype will be expressed if there are one or two brown
alleles present. The recessive scarlet phenotype is ex¬
pressed only when the organism is homozygous, carrying
two scarlet alleles. There are also cases when incomplete
dominance occurs, and crosses between discrete pheno¬
types will yield an intermediate phenotypic expression.

An inheritance pattern of the discrete type may vastly
simplify the selection procedure, perhaps even enabling
complete fixation of one expression in one generation.
Falconer (1960) discusses implications of simple modes of
inheritance; more detailed discussions of the genetic
analysis of discrete traits are in general genetics texts
such as Strickberger (1968) or Merrell (1975).

Most characters that interest insect breeders will be
quantitative traits. For characters like DDT resistance
(Pielou and Glasser 1952), body weight (Bartlett et al.
1966), and mating behavior (Manning and Hirsch 1971,
Kraemer and Kessler 1975), many genes are influencing
the phenotypic expression of the trait, and clear-cut dom¬
inance relationships and modes of inheritance are not
seen. Because quantitative traits are more complex and
more common than qualitative traits, my discussion of
artificial selection deals mainly with quantitatively in¬
herited characters.

Effect of environment

The phenotype, or observable properties, of an organism
is what is being measured. The phenotype is the result of
interactions between the genotype, or genetic makeup, of
the organism, and the environment that it lives in. For
selection to occur, the phenotype must have some varia¬
tion that can be attributed to the genotype. If all varia¬

tion of the phenotype is caused by environmental condi¬
tions, then selection cannot be successful. Therefore,
studies need to be conducted with a controlled environ¬
ment that will reduce environmental variation during
measurement of a phenotype. Measurements may have to
be made in precisely controlled environmental chambers
with the insects at a constant temperature and humidity,
on the same food source, measured at the same age, and
so forth. The more closely all environmental conditions
that might affect the character can be controlled, the
more accurate will be the assessment of the genotypic
variation present in the population. The effects of en¬
vironment on heritability and selection response are dis¬
cussed by Falconer (1952) and Rendel and Binet (1974).

The type of character being measured will also influence
the amount of environmental variation affecting the ex¬
pression. Morphological and physiological traits are less
open to environmental influence during development and
expression than are behavioral traits.

Measured phenotypic variance, Vp, can be divided into
several components. The first is the genetic variance, VG,
which itself can be divided into additive genetic variance,
V A, dominance deviation, VD, and interaction deviation,

Vv All other variation is environmental, VE, and is con¬
sidered beyond experimental control. So

VP=VA + VD+V^ -VE.

Of the three types of genetic variance, VD and Vt are
generally considered to be less important than VA. VD
arises from the property of dominance among the alleles
making up a genotype; and Vp usually rather small, is
generally treated as a negligible complication. The
breeding value of the organism, VA, can be measured
relatively easily in several ways and expressed as a ratio
of additive genetic variance to total phenotypic variance.
This ratio is the estimated heritability. The assumption
made here, that environmental deviations and genotypic
values are independent of each other, is not entirely true.
Kulincevic and Rothenbuhler (1975), for example, found
that susceptibility to disease varied with the virulence of
the pathogen. One way this complication can be overcome
is by specifying that the genotype be measured under
specific conditions. Another assumption that is not
always justifiable is that a specific environmental dif¬
ference has the same effect on different genotypes (Bray
et al. 1962, McNary and Bell 1962, Jinks and Connolly
1975). But comparing genotypes under favorable and un¬
favorable conditions may show that a genotype that does
best under one set of conditions may not have the
greatest yield or fitness under another. This variance of
interaction between genotype and environment, usually
regarded as part of the environmental variation, should
not be overlooked in measurement of a characteristic, par-

10

ticularly when the same species is studied in different
habitats (Druger 1962, Nye and Mackensen 1970).

Heritability

One of the most important properties of a quantitative
trait is its heritability, h2, the ratio of additive genetic
variance to the total or phenotypic variance;

h2=VA/VP,

that is, the proportion of total variance attributable to
additive effects, which are the average effects of all genes
affecting a character. The size of h 2 indicates how alike
related organisms are. The most important function of h 2
in the genetic study of quantitative traits is that it can
predict how reliable the phenotypic value is as a guide to
the organism’s actual breeding value. So heritability is a
measurement of the proportion of the phenotypic varia¬
tion that is attributable to genetic causes amenable to
selection.

The value of heritability ranges from 0 (no genotypic in¬
fluence on the variation of the trait) to 1 (all variation of
the trait is genetically produced). Traits that are closely
connected to reproductive fitness generally have low heri-
tabilities. For example, values for litter size, egg produc¬
tion, egg-laying rate, and ovary size range from 0.1 to
0.3. Another reason for a low heritability value is in¬
efficient measurement. If the technique does not accu¬
rately measure the desired trait, environmental variance,
VE, may be increased considerably; the proportion re¬
sulting from additive genetic causes would be reduced;
and heritability would be decreased. Higher H2-values are
expected in characters less important to reproductive
fitness such as coat color, patterns of spotting (A2=0.95
in mice; Strickberger 1968), and spot number (McWhirter
1969). These traits may be controlled by one or just a few
genes. As an example of /^-values, consider some traits of
Drosophila melanogaster Meigen: abdominal bristle
number, 0.5; body size, 0.4; ovary size, 0.3; and egg pro¬
duction, 0.2 (Falconer 1960).

Heritability is a property not only of a specific character
but also of the population and of the environmental cir¬
cumstances influencing measured individuals. Environ¬
mental variance depends on the conditions of culture or
management of the organism— more variable conditions
reduce the heritability, more uniform conditions increase
it. Genetic components are influenced by gene frequencies
in the population, and these may differ between popula¬
tions because of their different histories. Small popula¬
tions maintained for a long time become more genetically
uniform than do large, randomly mating populations, and
they show lower heritabilities.

The simplest way to evaluate heritability would be to
measure a population of mixed genotypes and one of
identical genotypes in several environments. The first
population would provide an estimation of total pheno¬
typic variance;

Vp—VA + VE.

The second would measure only environmental variance,
because all genotypes would be identical. The difference
between these two phenotypic variances would be the ad¬
ditive genetic value. Heritability could then be directly
calculated from the ratio of additive genetic variance to
total phenotypic variance.

The standard approaches to measuring heritability re¬
quire comparing the merits of related individuals and
estimating heritability from the covariance between them
or from a regression or correlation coefficient. Estimates
of heritability from covariance and correlation in do¬
mestic animals must consider a major environmental
source of covariance, the maternal effects. But such in¬
fluences as a common uterine and rearing environment
for animals of the same litter or mother may not be par¬
ticularly important in insects.

A straightforward method for estimating heritability is
to use the regression of offspring on parent. The data,
measurements of parents and the mean values of their
offspring, are used to calculate a regression coefficient, b.
If this is the regression of offspring on one parent, b , it
is a valid measure of Vih2\ if the regression is offspring on
midparent (average of the two parents), bop, it actually
measures h2. Examples of using this method for
estimating heritability in insects are found in Enfield et
al. (1966), Morris and Fulton (1970), and Wong and
Boylan (1970).

Heritability is most often estimated by sib analysis. Each
of several males (sires) is mated to several females (dams),
and some offspring from each female are measured. The
individuals measured form a population of half-sib and
full-sib families. An analysis of variance is calculated to
divide the phenotypic variance into components attrib¬
utable to differences in sires, in dams mated to the same
sires, and among offspring of the same female. The
variance component from sires, dams, and the total must
be calculated from the mean square values (table 1). The
total variance, or phenotypic variance, is calculated
because it is not necessarily equal to the observed
variance as estimated from the total sum of squares,
though the two seldom differ by much. With these
values, estimates of heritability can be made from the
sire component, the dam component, or a combination of
the two (table 2).

11

Table 1. — Formulas for calculating components of phenotypic
variance from analysis of variance mean squares (MS') for a
population of sibs and half-sibs1

Source of variance Variance Calculation2

Between sires a2$[ = MSsire—MSciam/dk

Between dams °2dam = MSdam-MSwlthm /k

Within progenies a\ithin = MS within

Total population a\ ^^re+^dam+^wlthm

‘See also Falconer (1960).

2d=number of dams; fc=number of offspring per dam.

Table 2.— Formulas for calculating
heritabilities from phenotypic var¬
iances determined from analysis-of-
variance mean squares for a popula¬
tion of sibs and half-sibs (table l)1

Heritability

estimate

Calculation

h2 .

sire

sire/** total

b2 dam

dam/** total

^ combined.

2(°2sire + °2daJ°2 total

‘See also Falconer (1960).

Heritability can be estimated from the offspring-parent
relationship in a population with the structure set up for
sib analysis. For many domestic animals, however, such a
population structure has few male parents, so the simple
regression of offspring on one or the other of the parents
is unsuitable. But heritability can be estimated from the
average regression of offspring on dams— regressions are
calculated for each group of dams mated to the same sire,
and they are pooled to give a weighted average.

These methods of estimating heritability have been
developed for use with diploid organisms. If the par¬
ticular insect being evaluated is not diploid in both sexes,
somewhat different forms may be required. For example,
Rinderer (1977) modified sib analysis for the honey bee, a
haplodiploid colonial organism; he also delineated several
problems of estimating heritability in a haplodiploid
social organism.

Artificial Selection
Selection methods

Selection of males and females with the desired charac¬
ters to parent the next generation can be done in several
ways (see Wright 1921, Mather 1941, and Hazel and
Lush 1942). Single individuals can be chosen on the basis
of their own phenotype (for example, the expression of a
particular mutation) to be mated with a specific other in¬
dividual. This is individual selection. In a variation on in¬
dividual selection called mass selection, large numbers of
selected individuals are put together en masse for mat¬
ing, a common occurrence in rearing large numbers of
insects.

Family selection is the choice of individuals based on the
mean phenotype of the family that they come from. This
method requires selection of the entire family for use as
parents and is preferable for characters having low herita¬
bility. But the method is limited because all members of

the family will generally be reared in the same environ¬
ment. One variation of family selection is sib selection,
usually used for traits requiring the death of an orga¬
nism, in which an individual’s phenotypic value is based
on measurements of its siblings. Another is progeny
testing in which offspring are measured. If only the best
individual from each family is chosen for mating, the
method is referred to as within-family selection. This ap¬
proach is desirable if a common environment, such as a
shared uterus, has a major effect on the size of the en¬
vironmental variance.

In all cases of selection by group, the calculations of h 2
and R (response to selection), differ from those used with
individual selection (see Kojima and Kelleher 1963, Wil¬
son et al. 1965, Berger and Freeman 1974, and Katz and
Enfield 1977). For these calculations, an assumption is
made that the generations are kept separate— individuals
are selected from a base population, used to parent off¬
spring (first selected generation), and only the offspring
are chosen to parent the second selected generation. The
generations would not be separate if selected individuals
from the base population and the first generation were
intermated to produce the second generation. For most
insect-rearing procedures, generations will probably be
discrete.

Response to selection

Selection will change the population mean of the selected
character by an amount, R, the response to selection. A
second parameter, selection differential, S, is the measure
of average superiority of those individuals selected as
parents over the total population. R and S (fig. 1) are re¬
lated by the regression coefficient of the offspring on
their parents, bop. So

RIS=bop, or R=bopS.

If there are no significant nongenetic causes of resem-

12

blance, such as maternal effects, and the selected pheno¬
type is not correlated with general fertility and viability
within the selected population, then R and S can be re¬
lated directly to heritability. So

R=h2S.

valuation, the extremes will be quite different from the
average values for the trait, hence a high value of S. With
little variation, the differences can be slight. The estimate
of S for different traits and different populations is ex¬
pressed in terms of c t (square root of Vp) to determine
the quantity i, intensity of selection. So

The relationship of R, S, and h2 is most useful for predic¬
tion. Once parents have been selected for production of
the next generation, S will be known. The value of bop,
the regression coefficient, can be calculated from the
previous generation; or an estimate of heritability can be
made from the parental population. Selection does change
the population, and these two values, bop and h2, can
change with the selection. Theoretically, then, the predic¬
tion is accurate only for a single generation. In practice,
however, the predicted value of regression or heritability
actually holds true over several generations (Falconer
1960).

There are two factors affecting the size of the selection
differential, S. These are the proportion of the population
selected to be parents and the phenotypic standard devia¬
tion of the trait being selected. If a small proportion of
the population is selected for mating, that proportion will
represent only the most extreme members of the popula¬
tion; their mean value, Xpo, will be very different (high
value of S) from the mean of the total base population,
Xp'o (fig. 1). But, if the proportion of parents is larger,
many will have more intermediate values and the paren¬
tal mean, Xp'o, will be less different (low value of S) from
the total mean, Xp'o. The phenotypic standard deviation,
or variation, affects the size of S, the difference between
selected extremes and the base population. With large

Figure 1.— Diagrammatic representation of re¬
sponse to selection, R, and selection differen¬
tial, S. R=Xpt—Xp0, and S=Xp'0— Xp„.

Slap=i, or R =iah2.

Manipulation of heritability, intensity of selection, or
phenotypic variation can improve the rate of response, R.
To increase heritability, one must decrease the environ¬
mental variance by manipulating rearing and measure¬
ment conditions. Decreasing proportion of individuals
selected, and so increasing the intensity of selection, will
have a similar effect. Of course, this decrease is limited
by the size of the population being used for selection,
which will be somewhere between the maximum number
of organisms that can be reared and measured at any one
time and those required to maintain a biologically func¬
tioning population. The selection intensity practiced per
unit of time can also be increased by decreasing the
generation interval or by maximizing the number of off¬
spring produced in each generation. For honey bees, this
increase might be several hundred colonies per year,
while for Drosophila the number of measured units (flies)
may be several orders of magnitude greater. Phenotypic
variation is much less amenable than heritability or inten¬
sity of selection to manipulation because it is limited to
what is biologically available. To manage phenotypic
variation, a breeder should insure that the base popula¬
tion has maximal variation included in its members, or he
should use crossbreeding during the selection program.

While artificial selection is being done, natural selection
will also be affecting the population. Mainly, it will alter
the fertility of the selected parents and the viability of
the offspring (Wilkes 1947, Hiraizumi 1961, Kress et al.
1971). Using weighted values from the parents based on
the different numbers of offspring that each group con¬
tributes to the succeeding generation, one can recalculate
the selection differential. This quantity is the effective
selection differential rather than the expected selection
differential. If there is a large difference between these
two calculated values of S, then this population is under¬
going a great deal of natural selection.

There will be some variability in the population mean
from generation to generation (fig. 2), mainly because of
how the environment affects the expression and measure¬
ment of the phenotype. So, to be most precise, estimates
of R need to be made over several generations. Such esti¬
mates are made by fitting a regression line to a series of
generation means (Robertson and Reeve 1952, Englert
and Bell 1970). The slope of this regression line then
represents the best measure of the average response per

13

CHARACTER SCORE

O - Low Selected Line

5 10 15 20 25

GENERATION OF SELECTION

Figure 2.— Results in a hypothetical bidirectional selection program.

14

generation. If selection is bidirectional— for example, if
one is selecting for high body weight in one population
and low body weight in another— calculation of the
response will be better than a comparison between a
selected population and an unselected control; bidirec¬
tional selection gives a greater divergence of the two
populations being compared.

Heritability can also be estimated as

h2=R/S.

If the value for S used in this calculation is the effective,
or weighted, selection differential, it will eliminate the ef¬
fects of natural selection and estimate heritability only on
the basis of artificial selection. This realized heritability
may not be the best estimate of the actual value of h2,
but it is the most useful figure for comparing the effec¬
tiveness of different selection procedures, especially when
different intensities or methods of selection are in use.
Realized heritability is usually calculated as the slope of
a regression line fitted to a plot of the generation means,
R, by the cumulative value of the selection differential, S
(Meyer and Enfield 1973, Choo 1975a).

Other considerations

in selection

In the same way that generations differ, responses to
selection may differ between different lines or popula¬
tions undergoing selection (Defries and Touchberry 1961).
Genetic parameters such as heritability and the regres¬
sion of offspring on parents may also differ. If the selec¬
tion is bidirectional, the response may be asymmetric
(Englert and Bell 1970). That is, selection may proceed
more rapidly in one direction than in the other (fig. 2).
Falconer (1960) gives several possible causes for this
variation in response.

The genetic structure of populations undergoing selection
will change through generations. One of the effects of
this change is that the size of the response does not re¬
main at its initial level but becomes smaller with suc¬
ceeding generations until the population reaches a
plateau for the phenotype, the selection limit. In figure 2,
for example, the low-selected population has reached a
plateau at generation 20 because the desired alleles have
been fixed (Bell et al. 1955, Brown and Bell 1961,
McEnroe 1967). If further progress is desired, the popula¬
tion must be given greater variation through the in¬
troduction of other alleles. These new alleles can be in¬
troduced by crossing highly selected lines, by introducing
environmental stress, by changing the selection criteria,
by using radiation (Bartlett et al. 1966), or by outcrossing
the selected population.

At the same time that selection for a desired trait is be¬
ing carried out, there may be unintentional selection of
correlated traits (Robertson and Reeve 1957, Hiraizumi
1961, Wong and Boylan 1970, Kress et al. 1971, Choo
1975b). Among the several causes for unintentional selec¬
tion is pleiotropy— one gene influences more than one
characteristic. Or several genes may be closely linked on

the chromosome, and selection for one gene will in¬
advertently carry along its linked associates. Conversely,
beneficial combinations of several genes may have evolved
together and be closely linked; though, if these link¬
ages are broken up during selection procedures, the
product might be less desirable or even harmful, pheno¬
types. Finally, traits may be correlated in some way
because of common parts in those traits. For exam¬
ple, honey bees have a 0.5 correlation between the rate of
hoarding sugar syrup in the comb and their response to
alarm pheromones. So high hoarders are usually fast
responders. This correlation may exist because response
to a chemical stimulus (sugar or alarm pheromone) is
necessary for both types of behavior (A. M. Collins and
H. A. Sylvester, unpublished data).

A major consequence of continued selection, particularly
in small populations, is inbreeding depression, a reduction
in the reproductive capacity and physiological efficiency
of the organisms undergoing selection. Hybrid vigor
(heterosis) occurs when the fitness lost during inbreeding
is restored after two inbred lines are crossed (Cale and
Gowen 1956, Enfield et al. 1966). One selection method
uses both inbreeding depression and hybrid vigor by in-
breeding many lines selectively for several generations
then crossing them to restore their fitness. Generally,
this technique is effective only in closely related popula¬
tions such as those reared in laboratories. Since widely
different wild populations fail to show heterosis when
crossed, each may be adapted to its own environment,
and progeny produced by crossing them are adapted to
neither (Falconer 1960).

In some organisms, breeding can have dramatic effects.

In honey bees, for example, the system for sex determina¬
tion is tied to homozygosity of a specific locus.
Homozygous and, in the case of haploids, hemizygous
bees are male. Heterozygous bees are female. Continued
inbreeding rapidly increases homozygosity. Within a few
generations, the probability that many of the diploid bees
will also be homozygous and develop as males becomes
quite high. As diploid males are generally destroyed by
the workers, the colony rapidly deteriorates. If such a
situation arises in organisms undergoing selection, the
rate of inbreeding during selection must be kept low.

When lines are crossed to make use of hybrid vigor, some
crosses will produce progeny that are more fit them those
produced by others. This variation is due to the combin¬
ing ability of each line. If the ability of a particular line
to combine with several other lines is measured and a
mean calculated, the result is the general combining abili¬
ty. Specific combining ability is the measurement of in¬
creased vigor between two lines. One can choose the ex¬
pression of combining ability by using a program called
reciprocal recurrent selection (Kincaid and Touchberry
1970, McNew and Bell 1970). In this plan, the selection of
parents in the lines is based on the performance of their
progeny from crosses with another line. The best combin¬
ing parents are then mated within their respective lines.

15

The selection index

In practice, the least effective way to select for several
characters is to use tandem selection— to select for one
trait at a time. Instead, an independent culling level can
be established for each trait being selected, and all
animals below this level for any trait are culled. This
technique requires many more animals than are needed
for tandem selection to have enough selected to carry on
a viable population. And selection may be slowed if cull¬
ing levels must be reduced to leave a viable breeding
population. So the best way to select for several
characteristics is to use a selection index, which combines
all the phenotypic measurements into a single value.

Each trait may be weighted according to its relative
economic value, heritability, and correlation with other
traits in the index. The result is one number that deter¬
mines the culling level (Hazel 1943, Falconer 1957, Tallis
1962, Okada and Hardin 1970, and Yamada et al. 1975).
Hoy (1979) gives an excellent flow chart showing the de¬
cisionmaking and experimental steps in developing such
a program. Before attempting to develop an artificial
selection program for a particular organism, one should
review the literature on related organisms to discover if
there may be special problems because of their specific
biology.

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19

Maintenance of Genetic Variability
in Reared Insects

By Dennis J. Joslyn1

Introduction

Without genetic variability, populations of organisms
cannot adapt to changing environments. In recent years,
many studies have been done on the levels and main¬
tenance of heterogeneity in natural insect populations (for
reviews, see Lewontin 1974, Wagner and Selander 1974,
Powell 1975, and Selander 1976). But there have been few
studies on the genetics of laboratory-reared insects, par¬
ticularly those being reared for use in nonchemical pest-
control programs. A few attempts have been made to de¬
scribe some of the genetic events occurring in laboratory-
reared insects (Bush 1975, 1977; Sluss et al. 1978;

Bartlett 1981; and Pashley and Proverbs 1981). But, even
with few studies to verify it, researchers generally sup¬
port the theory that mass-reared insects intended for
release must undergo some genetic changes during colon¬
ization; the artificial environment alone is enough cause
for such changes. To date, how these genetic changes af¬
fect the outcome of pest-control programs has not been
determined.

One effect of colonization can be the reduction in genetic
variability. This reduction could alter the ability of reared
insects to fulfill their role in a pest-control program. So
the adaptability of laboratory insects must be determined
by monitoring changes in genetic content, and adequate
levels of variability must be maintained.

Population geneticists today are debating what mecha¬
nisms cause natural populations to maintain high levels
of genetic variability; the debate is often called the
neutralist-selectionist controversy (Nei 1975, Kimura
1979). Discussions center on whether random processes
(neutralists’ view) or nonrandom events (selectionists’
view) are primarily responsible for maintaining the high
levels of variability in field material. The question is far
from being answered; but apparently both processes are
operating. In laboratory strains, the amount of variability
and the mechanism of maintenance should be determined
for each species, especially in mass-rearing programs. In
conventional biological control, parasites are reared to

'Assistant professor of zoology, Department of Biology, Camden
College of Arts and Sciences, Rutgers University, Camden, N.J.
08102.

suppress the pest species (Mackauer 1976); in genetic con¬
trol, the genetic apparatus of the pest is used to effect its
own reduction. Both approaches aim to curb the repro¬
ductive potential and therefore the size of the target
population by the release of large numbers of artificially
reared insects; and, in both cases, variability permits the
production of insects that will interact effectively with
the wild type. Eventually, the control principle for which
the insect was selected or engineered will spread through¬
out the native population via a series of timed releases.
These releases use planned ratios of laboratory to wild in¬
sects that are based on estimates of the target popula¬
tion’s density.

For any insect-rearing program, then, the purpose of
maintaining genetic variability is to insure the production
of competitive individuals that affect target populations
as planned. So mating competitiveness is especially im¬
portant in genetic-control strategies intended to transfer
a sterility principle into the wild population. Also, pred¬
ators produced for biological control must not lose their
ability to seek out their hosts. Because the laboratory is
an artificial habitat, artificial insects (biotypes or eco¬
types) can be produced inadvertently during colonization,
especially when the rearing environment does not vary
and insects adapt to one set of conditions. Such environ¬
mental uniformity may produce enough release insects;
but, if a less competitive organism results, the purpose of
their rearing is thwarted. For example, in a program
rearing the screwworm, Cochliomyia hominivorax (Coq-
uerel), the flies were unintentionally selected for a variant
of the flight-muscle enzyme, alpha-glycerophosphate
dehydrogenase (Bush et al. 1976, Richardson 1978). These
mutants could not transfer the sterility factor into the
wild population because their diurnal flight activity dif¬
fered from that of wild flies. Reproductive isolation had
developed in the factory -reared flies because of directional
selection.

Understanding genetic variability in laboratory colonies
requires understanding the arrangement of genes; sources
of variation; and the measurement, decay, and mainten¬
ance of heterogeneity. Understanding how genetic
variability decays and is maintained in nature helps us
understand how it decays and can be maintained in the
laboratory. But laboratory insects live in a controlled,
often unvarying environment; and the small size of
laboratory colonies limits the gene pool. Therefore, while
variability, gene arrangement, and sources of variation
may be similar between laboratory and field populations,

20

decay of variability is much more rapid in laboratory col¬
onies. Then, for a successful control program, it is
necessary to design appropriate measures that will offset
such rapid genetic decay.

Arrangement of Genes

External and internal

variability

The evolution of a species depends on changes in gene
frequencies over time. During this process, new genes ap¬
pear through mutation and increase in frequency in a
population, while others decline or are eliminated. These
changes are part of the mechanism of natural selection or
differential reproduction and enable organisms to adapt
to varying environments. Genetic variability is classified
into two categories— external and internal— on the basis
of phenotypic effects. Both help the evolving species to
adapt. External variability includes both dominant and
recessive morphological mutants and an assortment of
behavioral, physiological, and ecological variants. In the
heterozygous condition, recessive mutants, regardless of
the nature of their phenotypic effects, remain hidden;
they are a cryptic variability (Dobzhansky 1970, Lewon-
tin 1974). Genetically regulated physiological variability
is especially important to an insect-rearing program
because of the associated reproductive effects. Lethal,
semilethal, and sterility genes, for example, are built-in
genetic loads that lower reproductive fitness. Rearing
conditional mutants (those expressed only in specific con¬
ditions), such as those responsible for pesticide resist¬
ance, temperature sensitivity, and disease susceptibility,
may severely restrict the diversity of the rearing environ¬
ment. But a restrictive laboratory environment limits the
adaptive potential of the reared insect and may reduce its
effectiveness at release.

Internal variability comprises cellular and molecular
genetic differences; studying it usually requires special
methods of analysis. Currently, the forms of internal
variability being most widely applied in population
analysis are chromosomal and protein variation (Selander
1976, John 1981).

Many insect vector species are genetically polymorphic
for both external and internal traits. In their review of
the genetics of medically important arthropods, Spielman
and Kitzmiller (1967) pointed out the extent of external
variability in both field and laboratory populations.
Although they considered the value of internal chromo¬
somal variation to be important for analysis of population
genetics, Spielman and Kitzmiller also cited pioneer
studies on external variability in laboratory strains of
such pests as C. hominivorax, by La Chance and Hopkins
(1962); house fly, Musca dome stic a Linnaeus, by Sullivan

and Hiroyoshi (1960); German cockroach, Blatella ger-
manica (Linnaeus), by Ross and Cochran (1965); scrub
typhus chigger mite, Trombicula akamushi Brumpt, by
Goksu et al. (1960); and yellowfever mosquito, Aedes
aegypti (Linnaeus), by Craig et al. (1961). For Ae. aegyp-
ti, VandeHey (1964) relied on the inbreeding procedures
(Dobzhansky 1970) long used for studying variation in
Drosophila species. The procedures reveal the hidden
variability contained in heterozygous individuals.

Another genetic technique adapted to analysis of insect
pest populations from Drosophila studies has involved
the use of giant chromosomes. In salivary glands of
Anopheles mosquito larvae, polytene chromosomes have
been used extensively to detect inversion polymorphisms
(Kitzmiller et al. 1967; Davidson and Hunt 1973; Coluzzi
and Kitzmiller 1975; Kitzmiller 1976, 1977; Coluzzi et al.
1979). Unfortunately, most pest species do not have these
giant chromosomes. Recent developments in chromosome¬
banding techniques (Newton et al. 1974, Steiniger and
Mukherjee 1975, Motara and Rai 1977, Mezzanotte et al.
1979) use mitotic and meiotic chromosomes to reveal het¬
erochromatin variability in populations. These techniques,
including C, G, and Q banding, should provide an alter¬
nate source of information about the organization of chro¬
mosome variability in insect pests.

Some of the methods now being used to characterize in¬
ternal variability in populations stem from the field of
molecular biology. One method, gel electrophoresis, has
been particularly useful. By separating charged protein
molecules in an electric field, it has permitted the iden¬
tification and quantification of gene-enzyme products
known as allozymes. One advantage of the technique is
that gene products can be examined directly in an indi¬
vidual, and variability can be easily determined. Gene
frequencies can be found rapidly for a large number of
individuals, and shifts in allozyme content in a popula¬
tion can be detected for many environmental influences.

Although the value of genetics to insect rearing has been
recognized (Craig 1964), only recently has electrophoresis
been used for genetic analyses of mass-reared colonies or
longstanding laboratory strains (see, for example, Bush et
al. 1976, Munstermann 1979, Bartlett 1981, and Pashley
and Proverbs 1981). Analyses of natural populations have
shown that considerable variability exists at the allozyme
level (Powell 1975, Selander 1976). Insects being propa¬
gated for release must contain comparable levels of varia¬
tion and should genetically resemble as nearly as possible
the population to be controlled. When levels of variability
have declined, they should be restored before release. Or
loss of variability can be prevented when the colony is
established or when it is replenished at periodic intervals
afterwards.

21

Genetic load

Not all variability is beneficial. Inbreeding, which is un¬
avoidable in laboratory colonies, may elicit recessive
alleles that reduce the reproductive fitness of the popu¬
lation. So these genes are a genetic load; they may have
lethal or sterile effects. In a genetic-control program,
such genes could be very useful as a source of reproduc¬
tive incompatibility between laboratory and field stocks;
in fact, they are important as control mechanisms (David¬
son 1974, Pal and Whitten 1974). But, in general rearing
programs, the goal is to provide enough healthy insects
that can seek out wild insects. This goal requires that the
natural genetic load of the species be low; in a genetic-
control program, additional load or reduction in fitness in
the form of sterility is induced at the time of release.

Countering genetic load is helpful to a laboratory colony.
Much of the variability of heterozygous genotypes is po¬
tentially adaptive in appropriate circumstances. Such
variability allows insects to adapt to changing environ¬
ments. It is also highly desirable in a control program re¬
quiring that founding stock adapt to the laboratory
quickly yet remain variable enough to be able to interact
effectively with the target.

Sources of Genetic Variability

Mutation and adaptability

The ultimate source of all genetic variability is mutation,
or hereditary change, in the genetic material itself. Wheth¬
er or not a given mutant will be harmful depends on the
genetic composition at other loci and on its surroundings.
Mutations are random events that provide the variability
for selection to act on so organisms can adapt to varying
environments. Some mutations are not subject to selec¬
tion. These are distinguishable from both the harmful
(maladaptive) and beneficial (adaptive) types and are con¬
sidered to be adaptively neutral. In Drosophila pseudoob-
scura Frolova, for example, Yamazaki (1971) found that
laboratory flies carrying two different alleles for the es-
terase-5 locus showed no differences in viability, develop¬
ment, or fecundity. Furthermore, that the starting fre¬
quencies of these genes remained the same for 2 years in
experimental populations suggests that they were adapt¬
ively equivalent.

Since mutant genes do not exist alone, genetic back¬
ground influences their fate. In comparing the adaptive
value of radiation-induced mutants in Drosophila melan-
ogaster (Meigen) in a hymozygous background to then-
adaptive value in a heterozygous background, Wallace
(1958, 1963) showed that a new mutation in a hetero¬
zygous condition was better for a fly if the rest of its
genome was homozygous not heterozygous. Futuyma

(1979) suggested that such mutations could lead to en¬
hanced adaptation either in populations that are highly
inbred or in those in new environments. Both circum¬
stances occur in insect-rearing programs.

Survival of mutants

in insect colonies

Conditions that influence the spread of a newly arisen
mutant in laboratory colonies were generally considered
for diploid organisms by Kimura (1955, 1962). He found
that, as laboratory colonies are finite populations, var¬
iability in reared insects has to depend, in part, on muta¬
tions being able to survive and increase in frequency.
After the mutation rate, perhaps the greatest influence
on the rate of a mutation’s diffusion is population size.

In the laboratory, population sizes fluctuate. During an
expansion, a given mutation should increase in frequen¬
cy. A beneficial mutant would increase rapidly under
these conditions and eventually attain a fixed frequency;
a neutral mutation would ultimately become extinct. But
contractions of colonies should diminish the spread of
new variants. Spiess (1977) recounted the probabilities
for both the fixation and extinction of neutral alleles aris¬
ing through mutations; for q, a single neutral mutant
(one with no selection value) in a population having N in¬
dividuals, the ultimate probability for fixation is qo or
ll(2N). The initial frequency of a neutral mutation is also
1/(2A0. Since the frequency of the alternate allele, p, is po
or 1 — qo, then the probability of extinction for mutation q
is 1 — [1/(2A0], since po+q0=l. Usually, neutral alleles re¬
quire a long time to become fixed, about 4 Ne generations
(Hartl 1980), where Ne is the effective population size or
number of individuals actually contributing to the next
generation’s gene pool. In mass-rearing programs where
the colony must be divided to facilitate the rearing of
enough insects, many subpopulations may approach a
size of 10,000 or more individuals. A neutral allele arising
in a subpopulation of 10,000 members would have a very
low probability of becoming established (1 chance out of
20,000) and would require at least 30,000-40,000 genera¬
tions to become fixed.

Another factor influencing the rate of a mutant’s spread
in a colony is the generation time of the species. Shorter
generation times result in higher numbers of mating
pairs available to a colony during any given interval. For
selectively adaptive mutants in particular, higher num¬
bers of immediate descendants are more likely to receive
a mutation if the parent generation has more mating
pairs. Neutral and harmful mutations, however, would be
eliminated eventually, regardless of population size or
reproductive rate.

22

Genetic recombination

Most rearing programs involve sexually dimorphic spe¬
cies. Because of this separation of sexes into different in¬
dividuals, reproduction requires the union of haploid
gametes from each sex to form a diploid zygote during
fertilization. This fundamental aspect of sexual repro¬
duction affords another source of variability through
genetic recombination. During meiosis, homologous chro¬
mosomes may exchange segments during pairing (synap¬
sis) and crossing over. This reassociation of alleles am¬
plifies genetic variability, producing new phenotypes. For
example, in the experiments on D. pseudoobscura done
by Dobzhansky and his coworkers on how recombination
affects variation in viability, recombining advantageous
alleles at different loci produced different viabilities in
changing environments (Lewontin 1974). Though genetic
recombination results in new gene assortments, not all
arrangements may be beneficial. Often, a population will
have blocks of coadapted genes (adapted to function to¬
gether). If these blocks are disturbed by recombination,
the fitness of the organism will change.

Measuring Heterogeneity

Heterozygosity and polymorphism

Heterozygosity and polymorphism are measurements
that provide estimates of the amount of variation in a
population. In classical genetics, heterozygosity occurs
where one or more loci have dissimilar alleles. For homo¬
zygosity, alleles at a given locus are the same. But unex¬
pectedly high heterogeneity has been found in natural
populations, mainly through electrophoretic analysis of
allozymes. So heterozygosity has now become an impor¬
tant measure of population variation. Two types of heter¬
ozygosity-individual (/z) and population (H)— are now
recognizable, and may provide useful information about
the genetic profile of any population. The quantity h is
defined as 1— [2(x(.2)], where xt is the frequency of the ith
allele in a population (Selander 1976). The value of h is as
a measurement of the average amount of variability that
exists at a locus. The average h across all loci examined
(AO is the value H defined as |1 — [2(*,2)]} IN. H is the
most frequently used measure of heterozygosity and
denotes an average overall level of variability in a
population.

Polymorphism has been defined by Selander (1976) as the
occurrence of two or more alleles at a locus with the least
frequent not maintained by recurrent mutation alone.

The frequency of the least common allele is usually set
between 0.01 and 0.05. In practice, most levels of genetic
polymorphism are determined from electrophoretic data.
Allozyme separations allow calculation of gene frequen¬
cies directly from gels. The most meaningful measure of

polymorphism is the percentage of loci that are poly¬
morphic IP). In natural populations of insects, this figure
commonly runs as high as 50% or more. But H values for
Drosophila species, for example, range from 10% to 20%.

Other measurements of variability that are commonly
used have also developed from electrophoretic studies.
The average number of alleles per locus provides one such
estimate. In two studies of esterase isozymes in mos¬
quitoes, for example, considerable variation has been
found in natural populations. Saul et al. (1977) found as
many as 14 codominant alleles at the esterase-6 locus in
Ae. aegypti. In the mottled wing Anopheles, Anopheles
punctipennis (Say), the esterase- A and esterase-B loci
(Narang and Kitzmiller 1971) have seven codominant
alleles each. As more loci are studied in a population, the
total number of electrophoretically distinguishable alleles
can be determined and used as an estimate of overall
heterogeneity. In both field and laboratory populations,
as many (at least 25) different gene-enzyme systems
should be assayed as possible to accurately estimate the
amount of variation. Statistically reliable sample sizes
are critical also. Lewontin (1974) suggests that at least
50 individuals be examined per locus for a reliable profile.
This number is based on 2 parental genomes per in¬
dividual, or 100 total genomes. Although the electro¬
phoretic separation of gene products is a sensitive tool
for examining population structure, it does not detect all
changes at the gene level (Nei 1975). So the true amount
of variation is underestimated; this fact should be care¬
fully considered, especially in monitoring populations as
they become adapted to the laboratory.

Decay of Variability

Random events — genetic drift

and founder effect

If unopposed by directed events like selection, random
processes may reduce variability by eliminating some al¬
leles. Loss in diversity of genetic material can be caused
by both random and directed processes. Decay actually
begins at the time wild material is selected for propaga¬
tion in the laboratory (founder effect). “Bottleneck
effect” describes differences in genotypic variability that
occur after wild insects have been incorporated into a
rearing program— with a decline in variability comes an
associated loss of adaptive potential. But much of the
heterozygosity present in wild insects may be nonadapt-
ive. So, even if gene frequencies are squeezed at the time
of colony establishment, the bottleneck effect may not be
severe.

Because of the founder effect, which is a form of sam¬
pling error, only part of the wild gene pool is actually
incorporated into the laboratory stock. The founder effect

23

is an extreme example of genetic drift (Wright 1977),
which is the change in gene frequencies in a population
due to random events. Loss of genes during establish¬
ment of the laboratory stock restricts the adaptability of
the colonized material. For many species, counteracting
this source of decay may require that new genes be add¬
ed early in a mass-rearing program. At least, a profile of
the wild material from throughout its range should be
compared to a profile of the founder stock at the time of
colony establishment. Either external or internal markers
would be useful for this comparison.

Genetic drift is the most important of the random pro¬
cesses influencing gene frequencies in a laboratory colony
of insects. Even mutation rates of 10-5-10~6 do not
change genetic content of a population as significantly
over the short term. In larger populations, the decay due
to drift is less extreme than in smaller ones.

As population size varies after colony establishment,
more genetic decay occurs, especially if the number of in¬
sects declines. Loss due to disease or arbitrary culling,
for example, will reduce the effective population size (N).
Also, to make rearing on a large scale more manageable,
colonies are usually broken up into subpopulations that
may actually undergo varying degrees of drift. One cage
of adults, for example, may have a distortion in sex ratio,
with females far outnumbering males. In such a case, the
AT-value would diminish because males would be insem¬
inating more females.

Directional events

Inbreeding.— Inbreeding generally causes genetic decay,
as heterozygous genotypes are lost and homozygosity in¬
creases. During inbreeding, many of the recessive alleles
that were not being expressed in the heterozygous condi¬
tion can be uncovered. If harmful, these recessives in the
homozygous condition may cause inbreeding depression
or reduced fitness. The simplest case to consider is a
diploid population where a particular locus has two al¬
leles— a dominant wild type, A, and a recessive mutant,
a. Matings between two heterozygous individuals, Aa,
will result in 50% of the progeny being heterozygous.
With random mating among progeny genotypes and an
infinite population size, the proportion of heterozygous
individuals in the next generation would again be 50%.
But no population, laboratory or natural, breeds com¬
pletely at random or has infinite size. So AA genotypes
inbreed, as do aa genotypes. This inbreeding causes
decay because the number of heterozygous individuals
declines. Through hybrid vigor, heterozygous individuals
may be better adapted than either homozygous genotype.
Neither overall gene frequencies nor amounts of variation
change directly because of inbreeding, but the propor¬
tions of genotypes do.

When closely related individuals mate among themselves,
homozygosity increases across all loci. This is the pri¬
mary effect of inbreeding. If phenotypes controlled by
one gene mate assortively, then that locus only (and
sometimes associated loci) will undergo homozygosis.
Such single-gene effects may not seriously influence the
amount of heterogeneity. But, in a laboratory colony,
they could further the process of subdivision in one sub¬
population.

Not all colonies undergo inbreeding depression. For ex¬
ample, I made allozyme comparisons among four
wild populations of the mosquito Anopheles albimanus
(Wiedemann) and a stock that had been maintained at
the U.S. Agricultural Research Service’s Insects Affect¬
ing Man and Animals Research Laboratory in Gaines¬
ville, Fla. I found that the colonized strain lost little of
its variability, even after 40 generations (table 1). In
these studies, I used H and P values for several loci
whose mode of inheritance had been determined. Similar
comparisons by Bartlett (1980) of the pink bollworm, Pec-
tinophora gossypiella (Saunders), also revealed little
change in the genetic content of a laboratory strain. An¬
other electrophoretic study, of 12 loci, was conducted by
Pashley and Proverbs (1981) on the codling moth, Las-
peyresia pomonella (Linnaeus). Although they noted a
slight loss, their findings showed that heterozygosity
values did not change significantly over 25 generations.
The adaptive value of the structural loci examined elec-
trophoretically in these studies was not known. But none
of the three colonies declined in variability. And the sim¬
ilarity of their genetic profiles suggests that genetic com¬
patibility can be maintained between longstanding
laboratory stocks and wild insects.

Wahlumd’s effect. — A laboratory colony that is divided
into several isolates, as is often done in mass-rearing pro¬
grams, may decline in genetic diversity because it loses
heterozygous genotypes. This decline, known as Wah-
lund’s effect, or the stratification principle, occurs if gene
frequencies differ among the isolates. The result is an
average simulated inbreeding effect across all isolates
despite random mating in any one subpopulation. Homo¬
zygous genotypes increase and heterozygous ones de¬
crease as a function of the variance in alleles between
isolates. This variance is generally defined as the differ¬
ence between the observed frequency of the average
homozygote, aa, among isolates and the expected homo¬
zygote frequency obtained by squaring the average of the
frequency of the recessive allele of all subpopulations.

In insect colonization, Wahlund’s effect is important as a
potential source of genetic decay. If the mating pattern
of the insect is known, subpopulation structuring and its
associated simulated inbreeding can be distinguished
from true inbreeding as a contributor to decay, and ap-

24

Table 1. — Gene frequencies in populations of Anopheles albimanus (Wiedemann)

Frequencies in—

Natural populations in El Salvador

- A laboratory1 population—

1 2 3 4 SANTA TECLA

Locus

[H= 0.21;
P= 0.59)

(if =0.28;
P= 0.67)

(ff= 0.25;
P=0.61)

(ff=0.14;

P=0.51)

(ff=0.15;

P=0.53)

PGM,100

0.86

0.81

0.84

0.72

0.87

PGM, 095

.14

.19

.16

.28

.13

PGM2100

.99

.96

.99

.91

.99

pgm2102

.01

.04

.01

.09

.01

IDH1 100

.92

.86

.89

.90

.96

IDH j091

.08

.14

.11

.10

.04

aldox2100

.98

.99

.99

.99

.99

ALDOX2094

.02

.01

.01

.01

.01

HK^-00

.74

.82

.77

.83

.88

HK1 0-98

.26

.18

.23

.17

.12

HK4100

.83

.83

.88

.91

.93

HK .0-96

.17

.17

.12

.09

.07

‘Laboratory colony maintained at the U.S. Agricultural Research Service’s Insects Affecting Man and
Animals Research Laboratory, Gainesville, Fla.

H= Population heterozygosity. P= Percentage of polymorphic loci.

propriate measures can then be taken to offset the
decline. Hartl (1980) took another view of Wahlund’s ef¬
fect. He suggested that it can be, on a limited scale, one
means of negating subpopulation effects. Hartl refers to
the “isolate-breaking” character of Wahlund’s effect,
where isolated subpopulations behave genetically as
though fused together so that heterozygosity levels in¬
crease. In natural populations, such fusion, which leads
to isolate breaking, is done through migration; however,
in the laboratory, this could be simulated through con¬
trolled mingling of isolates before release.

Directional selection. — Selection operates continuously
during the establishment and maintenance of a colony.
The all-too-familiar problems that accompany the early
generations are exasperating evidence of the action of
directional selection, one form of which is artificial se¬
lection. Usually, the traits being selected are quantitative
or continuous; so several genes may be contributing to
the final phenotype. These polygenic traits that a new
colony must contend with include fecundity; develop¬
mental rates; longevity; sex ratio; dietary range; photo¬
period; and preferences in temperature, humidity, and
host. If the uniform environment of the rearing facility is
to accommodate all these environmentally sensitive

genes, then directional selection must differentiate each
appropriate genotype to mold the insect to the rearing
conditions. In the early stages of establishment, direc¬
tional selection filters out insects that cannot adjust to
the extensive change This action contributes to genetic
decay and accounts for the high mortality of newly intro¬
duced material. It may also be responsible, in part, for
some of the mysterious reductions in size that afflict
mass-reared colonies from time to time. After colony es¬
tablishment, the maintenance procedure automatically
maximizes production by selecting as parents the most
fit individuals. Here, directional selection is artificial
selection in the truest sense, because any phenotype may
be refined to any degree by the rearing team. Such a con¬
certed selection regime, involving quantitative traits, is
called truncation selection (Wright 1977). In the mass-
reared colony, such traits as fecundity, dietary prefer¬
ence, and longevity should be the most susceptible to
truncation selection. In any population, the variation of
any one of these three traits follows a normal distribu¬
tion curve. Depending on the needs of the control pro¬
gram, a cutoff, or truncation, point is chosen for a phenotype.
This truncation point determines the precise portion of
each generation’s progeny that will be parents for the
next generation. Individuals with phenotypes above the

25

truncation point are saved while all others are discarded.
The truncation point lies closer to the mean phenotype of
the most fit individuals for the character being selected
than it does to the mean phenotype of the whole
population.

Through directional selection, genotypes adapt to un¬
varying laboratory conditions and so lose their ability to
survive in varying environments. While this loss helps ef¬
ficient production, the insect produced is highly inflexible
and could not be expected to compete with its wild
counterparts. To neutralize the effects of truncation
selection, environmental conditions should be varied in
the insect-rearing facility.

Methods

of Maintaining Heterogeneity
in a Laboratory Colony

Modes of selection

The means that sustain genetic variability in natural
populations can be used as models for ways to maintain
it in the laboratory. Sources of decay in the laboratory
can be offset by several different types of selection and
by other phenomena. One type of selection, disruptive or
centrifugal, can be multidirectional either across individ¬
uals in one local population (deme) or across demes in a
species (Wright 1977). The net effect is a mosaic distribu¬
tion of allelic frequencies in a spatially restricted breeding
group. Density-dependent (Clarke 1972) and frequency-
dependent (Gromko 1977) selections promote changes in
the frequencies of opposing alleles. Given alternate condi¬
tions, the number of such alleles will either increase or de¬
crease so that they always have inverse frequencies. Group
selection favors the survival of the population over that
of the individual (Hartl 1980). In gametic selection, an
allele that would be lost in a gamete may be maintained
in a zygote. Overdominance, or heterozygote superiority,
is another means of maintaining alleles in a population
(Dobzhansky 1970); and, in the hitchhiking effect, a neu¬
tral allele may fluctuate because it is physically associ¬
ated with a selected locus (Thompson 1977).

Precolonization methods —

pooled multiple-founder colonies

Precolonization and postcolonization events affect the es¬
tablishment and maintenance of variability. Partial es¬
tablishment of heterogeneity, for example, can occur
when insects are field-collected for propagation. In na¬
ture, species consist of genetically unified, yet heterogene¬
ous, populations that are adapted to the local environment.
If insects intended for establishing a laboratory strain
come from one population, the adaptability of the colony
is limited; and release insects may not interact effectively

or uniformly throughout the range of the target pest.
Selection and pooling of founder insects from throughout
the range of the species can provide a much wider repre¬
sentation of the gene pool. The wider representation will
insure that laboratory material has greater fitness. Ini¬
tially, insects from a variety of geographical areas should
be mingled in the laboratory. The insects must, of course,
have reproductive compatibility. Although reproductive
isolation is the basis of speciation (Mayr 1963), its occur¬
rence in C. hominivorax, a major pest species, added a
surprising consideration to control programs. Makela and
Richardson (1978) reported that hidden reproductive iso¬
lates were present in the range of C. hominivorax; these
isolates contributed to mating incompatibility between
the release and field populations. The use of pooled mul¬
tiple-founder stocks or colonies to establish a laboratory
strain would preclude this source of incompatibility.

Postcolonization methods

Variable laboratory environment.— -For a laboratory col¬
ony of insects, even minor changes in rearing conditions
can affect variability levels. A static environment leads
to a static genotype and ultimately to less fit insects. En¬
vironments that are varied over space (Hedrick et al.
1976) and varied over time can contribute enormously to
the flexibility of an ongoing colony; the adaptive chal¬
lenges will be continual. Certainly, the dynamic environ¬
ment of the target populations is more closely simulated
by such a variable laboratory environment. The concept
is simple; putting it into practice in a large-scale rearing
program is not. One approach would be to pool insects
from parallel but environmentally different subcolonies
just before release. The number of different subcolonies
required would depend on the biology of the insect and
the type of control program, but procedures could be
varied in the same rearing facility. Some variables to con¬
sider with this approach are population densities and sex
ratios; temperature, humidity, and dietary preferences;
and container sizes. Heterogeneity could also be main¬
tained by varying the regime of each subcolony by, for
example, rotating parallel subcolonies through the en¬
vironments that were available in the facility. Such a pro¬
gram is feasible in most of today’s mass-rearing facilities.

Gene infusion.— Gene transfer between wild populations
usually occurs through migration. In reared insects, the
analogous process is gene infusion. New genes can be ob¬
tained either from natural populations or from parallel
subcolonies. Alleles introduced in this way have the same
limitations as a new mutant, particularly if a rare gene is
involved. During the life of a colony, the gene pool should
be rejuvenated occasionally with wild insects. Introducing
wild stock to the established colony restores some of the
heterogeneity that becomes lost because of drift or in-
breeding. The infused genes also help to maintain genetic

26

similarity between field and laboratory material; and they
augment mutation as a source of new, or unique, alleles
in the colony. Finally, through selective monitoring of
both field and laboratory populations (with electrophore¬
sis, for example), specific genes can be selected from the
donors to help genetically tailor the recipients. Because
the laboratory colony is a subpopulation of the species it
belongs to, gene infusion cements its genetic relationship
to the wild populations. So gene infusion offsets the
isolate-forming tendency of Wahlund’s effect.

Effective population size.-— Many reductions in colony
size that are caused by directional selection can be traced
to a specific environmental factor. Often the agent is a
subtle one, such as a change in a manufacturer’s dietary
ingredient or inadvertent use of cage materials that have
been treated with a toxin. These reductions decrease the
effective population size (IV) of the colony and must be
compensated for by adding outside individuals. To main¬
tain sufficient heterogeneity, a colony should not decline
below the number of founder insects. In natural popula¬
tions, several factors influence the Ne value. In Droso¬
phila species, such agents as disease, predation, competi¬
tion, or bad weather may require an effective population
size of 1,000 or more during warm weather, but only
about 100 when densities decline in winter (Begon 1977).
Laboratory colonies do not experience predation or hos¬
tile climatic conditions but are subject to disease and
competition. These can reduce the TV of the laboratory
colony to about 500. For mass-rearing programs, this
figure refers to the number of adults in a subpopulation
cage. The maintenance of a high IV reduces the effects of
random drift and inbreeding and helps to offset decay.

Acknowledgments

I collected allozyme data on Anopheles albimanus while I
was a research associate with the Insects Affecting Man
and Animals Research Laboratory, Agricultural Research
Service, U.S. Department of Agriculture, Gainesville, Fla.
32604. I wish to thank the New Jersey State Mosquito
Control Commission and the Rutgers University Re¬
search Council for support of this project. I gratefully
acknowledge Anne Celia for typing the manuscript.

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28

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603.

29

Section 2

Diets and Containerization
for Insect Hearing

The evolution of artificial diets, assurance of diet-ingredient
quality, and new container designs have had a major
impact on the development of basic and applied ento¬
mology. The rearing of insects on artificial diets has been
confined to this century, especially to the last two to
three decades. (The published research involving
laboratory-reared insects amounts to about 50% of recent
entomological publications.) Artificial diets relieve re¬
searchers from the trouble and expense of maintaining
greenhouse facilities and host plants for rearing
phytophagous insects. Continual advancements in such
diets and improvements in containerization have made it
possible for many additional species of insects to be
reared. Also, new in-field strategies for controlling pest
insects— the sterile-male technique, augmentative release
of parasites and predators reared in vivo and in vitro, and
application of microbial pathogens produced directly or
indirectly on artificial media— have placed new emphasis
and dependence on insect rearing.

To have a consistent, economical production of vigorous,
competitive insects (phytophagous or entomophagous), a
greater understanding about feeding is needed for each
phase of the development cycle in relation to behavior,
preference, nutritional requirements, etc. The diet should
insure proper insect nutrition at the lowest possible cost
and yet be practical to use. So, further research is re¬
quired for improving existing diets and formulating new
diets for those species that cannot be reared at the pres¬
ent time.

Since mass production has become more common, future
advances will depend on several things, including stand¬
ardizing diets and developing quality-control standards

for the diets and their individual components. Manj^ com¬
mercial dietary ingredients are presently subjected to
standardized analytical tests such as those found in the
Official Methods of Analysis of the Association of Of¬
ficial Analytical Chemists. But, little or no technical data
are provided for some commercially available products
used in artificial diets. In the final analysis, bioassays are
probably more important than chemical assays in deter¬
mining an ingredient’s nutritional value. A bioassay an¬
swers the question “what is the ultimate effect of the
diet on the reared insect?” The size and importance of
the rearing operation should determine the resources that
can be used for dietary assays. The handling and storage
of incoming ingredients are also important to overall diet
quality.

No discussion of artificial diets for insects would be com¬
plete without giving consideration to rearing containers
and their closures. Many different kinds of containers
have been used that were adapted from currently mar¬
keted items. As artificial diets were improved and more
were used, certain containers became generally accepted
because they provided favorable microenvironments for
the diets and insects. Sizes and shapes of containers vary
according to the type of diet and the behavior of the in¬
sect. Some desirable characteristics shared by most insect¬
rearing containers and closures include the prevention of
microbial contaminants and pathogens, allowance for
proper gas exchange, prevention of escape, moisture regu¬
lation, economics, visibility and accessibility, convenience
of handling and harvesting, and ease of cleaning and dis¬
infection. Additional research in containerization might
well center on design improvements for the particular
species and stress the use of reusable containers.

F. D. Brewer, research entomologist;

W. A. Dickerson, research entomologist;
R. L. Burton, research entomologist;
and

R. A. Bell, research leader;

Agricultural Research Service

31

Insect Diets

Historical Developments, Recent
Advances, and Future Prospects

By Pritam Singh1

Introduction

The continuous availability of large numbers of uniform
laboratory-reared insects of acceptable quality has been
an important contribution to the development of modem
experimental and economic entomology. Successful rear¬
ing of these insects has depended on sound knowledge of
insect biology, behavior, habitat, and nutrition. An under¬
standing of the mating habits, preoviposition and oviposi-
tion periods, fecundity, longevity, sex ratio, environmental
requirements, and food and feeding preferences of the in¬
sect is necessary in developing rearing techniques. So in¬
sect rearing is a complex field closely related to other
disciplines, especially dietetics and nutrition. Here, “die¬
tetics” is the study of the kind and quantity of food that
will be eaten. “Nutrition” refers to the specific, chemi¬
cally defined components that the insect must have to
grow, reproduce, and perform as it should. This paper re¬
views the development of insect dietetics and discoveries
about insect nutrition. It also discusses recent advances
in insect diets and future needs and possibilities. Finally,
it lists the most important references on various aspects
of insect diets.

Historical Developments
in Insect Diets

Milestones in the development

of insect diets

General insect rearing.— The pioneering work on artificial
diets for insect rearing was done by Russian and French
scientists in the early 1900’s. Bogdanow (1908) was the
first to rear an insect axenically from egg to adult when
he reared the blow fly, Calliphora vomitoria Linnaeus, on
a diet compounded from peptone, meat extract, starch,
and minerals. Later, Loeb (1915) succeeded in rearing
Drosophila spp. for five generations on a diet of grape
sugar, cane sugar, ammonium tartrate, citric acid, dipo¬
tassium hydrogen phosphate, magnesium sulfate, and
water. Guyenot (1917) was the first to rear an insect (Dro-

‘Leader, Insect Rearing and Nutrition, Entomology Division,
Department of Scientific and Industrial Research, Private Bag,
Auckland, New Zealand.

sophila ampelophila Loew) on a completely artificial diet.
And the first multicellular organism to be reared axe¬
nically from egg to adult on a chemically defined diet was
Drosophila melanogaster Meigen, which Schultz et al.
(1946) reared on pure amino acids, minerals, vitamins,
etc. Hawkes (1920) was partly successful in feeding the
larvae of the twospotted lady beetle, Adalia hipunctata
(Linnaeus), a coccinellid, on cooked or raw chicken eggs
and powdered dates. And Szumkowski (1952) was the
first to rear the predatory lady beetle, Coleomegilla macu-
lata DeGeer, on mammalian liver enriched with vitamin
C. Later, Szumkowski (1961) developed a mixture of fresh
yeast with glucose or sucrose solution that was better
and thus replaced the liver diet. Atallah and Newsom
(1966) improved on Szumkowski’s diet by formulating an
artificial diet that fed eight successive generations of C.
maculata and allowed the females to oviposit into diet en¬
capsulated in a sealed Parafilm tube where the eggs
hatched and the larvae developed to maturity.

Zabinski (1926, 1928) reared the oriental cockroach, Blat-
ta ( =Periplaneta ) orientalis Linnaeus, and the German
cockroach, B'latella germanica (Linnaeus), on 18 parts of
ovalbumin, 56 parts of starch, 20 parts of saccharose, 2.3
parts of agar, and 3.7 parts of a salt mixture. Later,
House (1949) formulated a chemically defined diet for B.
germanica. Several stored-product pests were reared on a
casein-based diet by Fraenkel (1943) and his associates in
the early 1940’s. These achievements have been reported
in earlier reviews, first by Uvarov (1928) who listed 600
titles, then by Trager (1941, 1947). Singh (1955) was the
first to report a chemically defined diet for the yellow-
fever mosquito, Aedes aegypti (Linnaeus); this diet has
formed the basis for rearing other species of mosquitoes
for nutritional studies. Rearing of hemipterous insects on
diet was not achieved until the mid-1950’s when Scheel et
al. (1957) succeeded with the large milkweed bug, Onco-
peltus fasciatus (Dallas), and the onespotted stink bug,
Euschistus variolarius (Palisot de Beauvois). Aphids ap¬
parently were more difficult than other insects to culture
and were first reared by two independent groups: In the
United States, Mittler and Dadd (1962) reared the green
peach aphid, Myzus persicae (Sulzer); in Canada, Auclair
and Cartier (1963) reared the pea aphid, Acyrthosiphon
pisum (Harris). Since then, over 30 species of aphids have
been successfully reared on chemically defined diets,
some for several generations (Kunkel 1977).

32

Rearing phytophagous insects. — Bottger (1942), working
in South Africa, was the first to report the use of an
agar-based diet for rearing of a phytophagous insect, the
European com borer, Ostrinia nubilalis (Hiibner). The diet
was compounded from casein, sugar, fats, salts, vitamins,
cellulose, agar, and water. This work was followed by
that of Beck et al. (1949) at the University of Wisconsin,
whose diet for O. nubilalis was formulated from highly
purified natural products containing an unidentified
growth factor that was later identified as ascorbic acid by
Chippendale and Beck (1964). In Japan, by including in
the diet an extract of the host plant used in the O. nu¬
bilalis diet developed by Beck et al. (1949), Ishii (1952)
successfully reared the Asiatic rice borer, Chilo sup-
pressalis (Walker), and Matsumoto (1954) successfully
reared the oriental fruit moth, Grapholitha molesta
(Busck). Vanderzant and Reiser (1956) used an aseptic
technique to rear the pink bollworm, Pectinophora
gossypiella (Saunders), on a diet devoid of plant extracts.
Since then, several phytophagous insects have been
reared on diets compounded from pure chemicals and
other nutritive substances completely unfamiliar to
insects.

One of the most important advances in rearing lep-
idopterous and other phytophagous insects was the use
of wheat germ as a primary nutrient source in for¬
mulating insect diets. This work was done at College Sta¬
tion, Tex., by Adkisson et al. (1960) for the pink boll-
worm. Berger (1963) further developed the use of this
wheat-germ-based diet for the large-scale rearing of
Heliothis spp. These two diets, with modifications, have
been the basis for rearing many other insects (for exam¬
ple, see Ignoffo 1963). Since then, wheat germ has been
used in several hundred insect diets (see listings in House
et al. 1971 and Singh 1974b, 1977a, 1977b). Shorey and
Hale (1965) used beans as the primary protein source in a
diet to rear nine species of noctuid larvae. Several va¬
rieties of beans have since been used for similar pur¬
poses in the formulation of lepidopterous diets.

Rearing entomophagous insects. — Yazgan and House
(1970) achieved a breakthrough in rearing an ichneumonid
endoparasitoid, Itoplectis conquistor (Say), on a chem¬
ically defined diet in aseptic conditions; and Yazgan
(1972) obtained 62% fecund adults from neonate larvae
reared on this diet. But House (1978) was the first to
achieve complete success by using an encapsulated
medium for rearing this species from egg to adult. Also,
Hoffman and Ignoffo (1974), using a semisynthetic
medium, reared an endoparasitoid wasp, Pteromalus
puparum (Linnaeus), with 44% adult emergence from
neonate larvae. Thompson (1975) succeeded in rearing an
ichneumonid ectoparasitoid, Exeristes roborator
(Fabricius) on artificial diet, with 80% survival. Hoffman
et al. (1975), using a synthetic diet, were the first to rear

an egg parasitoid, Trichogramma pretiosum (Riley), from
egg to adult in vitro. Several studies have concentrated
on developing an artificial diet for rearing the common
green lacewing, Chrysopa camea Stephens. Hagen and
Tassan (1965, 1970), for example, succeeded with an adult
diet containing enzymatic protein hydrolyzate and
Wheast, while Ridgway et al. (1970), also using artificial
diet, described a mass-rearing technique for C. carnea.
Vanderzant (1969b) reported the use of a semidefined diet
for rearing C. camea larvae, and Vanderzant (1973)
discussed improvements in the diet. Martin et al. (1978)
mechanically encapsulated Vanderzant 's diet in paraffin
wax, candelilla wax, polyethylene, and polybutene for
purposes of mass rearing C. carnea; the capsule is easily
consumed by second-and third-stage larvae, but first-
stage larvae have difficulty penetrating it. Mass-
production techniques using artificial diet for rearing C.
carnea and other entomophagous arthropods have been
reviewed by Morrison and King (1977). Currently, the
technology for rearing parasites and predators on ar¬
tificial diets is far less developed than the technology for
rearing Lepidoptera, Coleoptera, and Diptera.

Factory or mass rearing on artificial diets. — Several in¬
sects have been mass-reared successfully on different ar¬
tificial diets and used for various insect-control programs.
Some examples are: European com borer for virus and
plant-resistance studies; tobacco hornworm, Manduca
sexta (Linnaeus), for hormone studies; nutsedge moth,
Bactra verutana Zeller, for control of purple nutsedge;
saltmarsh caterpillar, Estigmene acrea (Drary), for virus
production; gypsy moth, Lymantria dispar (Linnaeus), for
virus production; and several species for programs using
sterile-male release, including Culex fatigans Wiedemann;
horn fly, Haematobia irritans (Linnaeus), and stable fly,
Stomoxys calcitrans (Linnaeus). Recent developments in
the mass production of tsetse fly, Glossina morsitans
Westwood, feeding on citrated blood via membranes have
made large-scale colonization possible (Mews et al. 1977).
The commercial production of crickets, blow fly larvae,
and Tenebrio larvae as baits for the fishing industry is
well established in Europe and the United States.

The rearing of the screwworm, Cochliomyia huminivorax
(Coquerel), by Melvin and Bushland (1936) on a mixture
of milk, blood, lean beef, and formaldehyde is another
milestone in insect rearing, as these were the first blood¬
feeding insects to be reared on an artificial diet. In 1978,
10 billion flies were produced for release in Mexico and
southern Texas as part of a sterile-male release program
(Bush 1978). The success of this effort has led to other,
similar programs also based on mass-rearing techniques
developed earlier. For example, in the mid-1960’s the boll
weevil, Anthonomus grandis grandis Boheman, was
mass-produced on artificial diet by Gast and Davich
(1966), and the R. T. Gast Rearing Laboratory at

33

Mississippi State, Miss., has since produced as many as
3-6 million boll weevil adults per week (Griffin et al.
1979). Martin (1966) and Mangum et al. (1969) described
a mass-rearing procedure using artificial diet for the pink
bollworm that enables production of more than 1 million
moths per day in the U.S. Animal and Plant Health In¬
spection Service’s rearing facility at Phoenix, Ariz.
Hamilton and Hathaway (1966) developed a mass-
production method using artificial diet for the codling
moth, Laspeyresia pomonella (Linnaeus), that is presently
used at the U.S. Agriculture Research Service’s (ARS)
laboratory in Yakima, Wash. This facility can now pro¬
duce 3 million moths per year; using similar procedures,
the Canadian Department of Agriculture’s facility in
Summerland, British Columbia, can produce 2 million
moths per month (La Chance 1974). A technique that in¬
cludes a prototype machine for dispensing artificial diet
for the com earworm, Heliothis zea (Boddie), was
developed at the ARS Southern Grain Insects Research
Laboratory in Tifton, Ga., where up to 1 million eggs/day
could be produced (Burton 1969). The increase in produc¬
tion of the com earworm from a few hundred per day to
about 120,000/day was achieved mostly through this and
similar mechanization; from March 1972 to February
1974, 6 million pupae were produced (Sparks and Harrell
1976). Researchers at the ARS Cotton Insect Research
Laboratory in Brownsville, Tex., developed a technique
and a low-cost soyflour and wheat germ diet for mass
rearing the tobacco budworm, Heliothis virescens
(Fabricius); production was stabilized at about 70,000
pupae/day, and pupae were shipped to St. Croix, U.S.
Virgin Islands, for moth emergence and release as part of
a test of sterile-male release (Raulston and Lingren 1972).
The mass-production method described by Henneberry
and Kishaba (1966) for rearing the cabbage looper,
Trichoplusia ni (Htibner), on artificial diet was used to
produce 10,000 adults/week at an average cost of
$0. 36/pupa. Recently, Chauthani et al. (1971), Poitout and
Bues (1972), and Vail et al. (1973) gave improved formula¬
tions for diets to rear cabbage loopers for production of
Bacillus thuringiensis Berliner and viruses.

The house fly, Musca domestica Linnaeus, and the little
house fly, Fannia canicularis (Linnaeus), were first suc¬
cessfully colonized by Lodge (1918) in England. He used
a mixture of casein, bread, water, and banana surrounded
by a layer of dry rubbish where the maggots could pu¬
pate. In the United States, Glaser (1927) continuously
reared the house fly on horse manure plus yeast. This
diet, with slight modifications, was the basis of most
house fly rearing until Richardson (1932) introduced
CSMA (for Chemical Specialties Manufacturers Associa¬
tion), a diet that provides year-round rearing on an ef¬
ficient medium. It is now possible to mass-produce
millions of house flies on diet, as is currently being done
at the ARS Insects Affecting Man and Animals Research

Laboratory in Gainesville, Fla., where 6 million flies are
produced each week for the production of 4-5 million
Spalangia endius parasites. The house fly has also been
mass-produced on diet for sterile-insect programs in Italy
and the Bahama Islands.

Recent Advances in Insect Diets

In recent years, much has been learned about insect
dietetics and nutrition and about ways to protect insect
diets from chemical and biological, especially micro¬
biological, contamination. The information that follows
summarizes these recent advances.

Dietetics

Over the past 20 years or so, much effort has been ex¬
pended combining the 40 to 50 nutrients common to
most foodstuffs into acceptable food for insects. Re¬
searchers have learned that, to encourage consistency in
compounding, the raw materials should be generally
available, economical, uniform in nutrient density, and of
unvarying quality. Adequate chemical analyses must be
conducted to aid the selection and formulation of diet
ingredients and to insure freedom from degradation, in¬
festation, contamination, or other harmful changes.
Texture, shape, particle size, and other physical char¬
acteristics of the diet are also important, depending on
the feeding habits of the insect.

Nutrition

The choice of a specific food by a species is often deter¬
mined by nonnutritional factors such as physical proper¬
ties and phagostimulants. So proteins, carbohydrates,
lipids, and vitamins must all be present in adequate sup¬
ply since not having enough of one nutrient can cause the
insect to use the food more slowly and less efficiently
than it should. An unsatisfactory nutrient balance may
lead to nutritional diseases affecting growth, develop¬
ment, reproduction, and other life processes (Friend 1959,
House 1959, 1961b, 1963, 1965). So House (1966) pro¬
posed three principles applicable to insect nutrition: (1)
The rule of sameness— all insects need the same nutrition¬
al quality whatever their feeding habits and systematic
position of classification, (2) the principle of nutrient pro¬
portionality-normal nutrition requires that nutrient pro¬
portions are metabolically suitable (there may be several
equally good nutrient balances), and (3) the principle of
cooperating supplements— supplementary sources of nu¬
trients may be important to most insects.

Sources of protein and energy are the largest parts of a
diet. The protein content of the diet should ideally con¬
tain all 10 essential amino acids in the correct propor¬
tions and be readily digestible so that they are available

34

to the insect. The amount of protein required in a diet is
influenced by its nutritional quality, which is determined
not only by its amino acid composition, but also by how
efficiently the digested food is used (Vanderzant 1973).
Nutritional quality may be seriously impaired by treat¬
ment during processing, particularly by overheating,
which can destroy amino acids and cause indigestible or
otherwise unavailable complexes to form. One of the two
most vulnerable and easily measured constituents is ly¬
sine, and it is generally used as an indicator of protein
quality. The most common protein sources used in insect
diets are casein, egg albumen, lactalbumin, and soy
protein.

When an insect is allowed unlimited access to food, it will
eat at least enough to satisfy its energy requirements.
Carbohydrates (starches and sugars) are the major energy
source in most diets. Sugars also serve as phago-
stimulants. Fats give more than twice the energy of
sugars or starches. Proteins also supply energy, but are
an expensive source. So insects are likely to eat less of a
high-energy diet than they would of a low-energy one.
Then the intake of the other ingredients will be propor¬
tionately less, and their concentration must be increased
so that the amounts ingested do not fall below what is
needed.

The dietary requirements for lipids, fatty acids, and
sterols in insects have been thoroughly documented (see
Clayton 1964; Fast 1964, 1966; and Gilbert 1967). They
provide energy; they are also essential to the develop¬
ment of wing buds— deficiencies, especially of linoleic and
linolenic acids, cause deformed wings in adult Lep-
idoptera (Chippendale et al. 1965). Unlike mammals, in¬
sects cannot synthesize the steroid ring. So cholesterol,
the most common sterol in insects, must either be ob¬
tained from dietary sources or be added in pure form.
Cholesterol is a precursor of ecdysone, the molting hor¬
mone. Insect sterol nutrition and metabolism are re¬
viewed by Clayton (1964) and Robbins et al. (1971).

Vitamin C (ascorbic acid) is in the diet for most
phytophagous species, though stored-product insects,
flies, and cockroaches can grow without it. Fat soluble
vitamins such as A, D, E, and K are often unnecessary.
But vitamin A is required by some insects for normal
vision, and vitamin E has been associated with reproduc¬
tion. The vitamin B complex— including thiamin, ribo¬
flavin, niacin, pyridoxine, pantothenic acid, folic acid, and
biotin is required, though cobalamin (B12) is not clearly
necessary to insects. Mineral requirements are more dif¬
ficult to assess because excluding particular inorganic
ions from synthetic ingredients grossly alters the balance
of those remaining. Nevertheless, sodium, potassium,
magnesium, chloride phosphates, and minor elements in¬

cluding iron, copper, zinc, and manganese are necessary
for optimal growth.

A major difficulty in compounding diets for chewing in¬
sects is to provide solidity with a water content of 80%
or more. Agar is most commonly used because it forms a
rigid gel at low concentrations of 2%-3% in most diets.
But agar is expensive, and replacements must be sought
for use in mass rearing. Agar may be reduced by increas¬
ing the fiber content. Fiber contributes little to overall
nutrition but is essential for binding the nutrients, pro¬
viding bulk, and giving proper dietary shape and texture.

Microbial contamination

Generally, micro-organisms in artificial diets cause
spoilage (Clark et al. 1961, Ludemann et al. 1979), alter
the biological performance of the insect (Singh and House
1970a, 1970b, 1970c; Singh and Bucher 1971), and may
harm symbionts in the gut (Buchner 1953; Fraenkel
1959a, 1959b; Brooks 1960, 1964; N.C. Pant 1973b). The
microbial contaminants most often encountered in ar¬
tificial diets are Aspergillus yeasts, Rhizopus bacteria,
and Penicillium molds. Several species of these organisms
may often be found on one diet. The antimicrobials com¬
monly used to combat these organisms include formalde¬
hyde, methyl p-hydroxybenzoate, sodium benzoate,
potassium sorbate, sorbic acid, streptomycin, penicillin,
and Aureomycin (chlortetracycline).

In reviewing what is known about how antimicrobials af¬
fect insects, Singh and House (1970b) examined what is
called the safe level. A compound’s safe level is the con¬
centration that does not reduce the yield of pupae and
adults or increase the time for larval development by
more than 25% of normal. Above the safe level, anti¬
microbials are harmful in proportion to the concentration
used (Singh and House 1970b, Singh and Bucher 1971);
insect size may be reduced, larval life prolonged, and mor¬
tality in larval and pupal stages increased (Singh and
House 1970a, 1970c). The ideal antimicrobial food ad¬
ditive should suppress a wide variety of micro-organisms
at a concentration safe for the insect.

If possible, diets should be sterile, and this can be
achieved by autoclaving at 15 lb/in2 of pressure for 15-20
minutes or by other means such as irradiation, chemical,
flash, or gas sterilization. Microbial growth can also be
prevented in diets by including mold inhibitors and an¬
tibiotics or by adjusting the pH. Such procedures are
satisfactory if the environment is clean, equipment is
sterilized, dietary ingredients are not initially contami¬
nated, and eggs are washed in detergent and sterilizing
solutions. Rearing laboratories and diets should be moni¬
tored regularly for microbial contaminants, and strict
sanitation and hygiene standards should be maintained.

35

Future Prospects
for Insect Diets

The developments in artificial diets discussed above have
made both small- and large-scale insect rearing possible.
But many problems remain, and researchers are concen¬
trating on resolving these. More must be learned about
how diet ingredients affect insect quality. More must be
learned about the dietary needs of host insects in pro¬
duction of parasites and pathogens. Optimum diets must
be developed for species that are not yet reared on artifi¬
cial diets. And ways to standardize diets must be explored.

Quality control

The performance of laboratory-reared insects is affected
by many factors that must be controlled for production
of a uniform, high-quality insect (Boiler 1972; Mackauer
1972, 1976; Hoy 1976; Huettel 1976, 1977; Boiler and
Chambers 1977a, 1977b). Chambers (1977) listed the crit¬
ical performance traits as vigor, irritability, activity,
sound production, response thresholds, reproductive po¬
tential and drive, biotic potential, and others. Changes
may occur in metabolic functions such as C02 output and
nutritional needs; in tolerance to temperature, irradiation,
or other physical factors; in fertility, fecundity, longevity,
or population-stress tolerance; and in biological conform¬
ities such as rhythms, mating behavior, host specificity,
other chemical and physical responses, pheromone
production, and mate recognition. Much research still
must be done on how diet ingredients affect insect quali¬
ty, so that quality-control procedures in mass-production
programs can be suitably adjusted. Some preliminary
work has already been done on how diet affects phero¬
mone production, enzymes, and vision.

Dietary effects on pheromone production. — Pheromone
precursors may be needed in the diet of insects mass-
reared for use in sex-pheromone traps used to monitor in¬
sect populations. Diets that do not contain natural host
material may cause the reared insect to produce insuf¬
ficient pheromones. For example, laboratory -reared boll
weevil males are as attractive as natives if they have ac¬
cess to cotton squares and flowerbuds as food; but phero¬
mone production is reduced by 50% at 1 hour after such
food is removed and by 90% at 24 hours (Hardee 1971).
Similarly, the pheromone production of bark beetles is en¬
hanced if they are fed glucose-supplemented diets (Pit¬
man et al. 1966).

Field-collected larvae of the brownheaded leafroller, Cte-
nopseustis obliquana (Walker), can be satisfactorily
reared to the adult stage on a general-purpose artificial
diet (Singh 1974a). But the species could not be con¬
tinuously reared on artificial diet in the laboratory
because males and females would not mate. Replacement

of 20% of the diet with lyophilized plant powder, from
Acmena smithii (Poivet), increases pheromone production
and therefore mating (R. A. Galbreath, unpublished data).
On the other hand, the light brown apple moth, Epiphyas
postvittana (Walker), has been continuously reared for
over 80 generations on the same diet devoid of any plant
material. Likewise, Miller et al. (1976) reported that there
is no difference in pheromone production between moths
of the female oak leafroller, Archips semiferanus (Walker),
reared on artificial diet and those reared on three species
of oak ( Quercus spp.). Also, males fed these diets respond
identically in laboratory bioassays and in field tests.
These findings conflict with the hypothesis that the com¬
position of moth sex pheromones varies with slight
changes in diet (Hendry et al. 1975, Hendry 1976). This
hypothesis has been further refuted by Hindenlang and
Wichmann (1977) who were among its original proponents.
Clearly, more study is needed of the relationship between
specific diet ingredients and pheromone production.

Dietary effects on enzymes. — Another concern for research¬
ers is how artificial diets influence insect enzymes. For
example, Ahmad and Forgash (1978) reared gypsy moth
larvae on a wheat-germ-based artificial diet and on oak
leaves and reported that growth, development, and
maturation are comparable for the two diets but that ac¬
tivity in the mixed-function oxidase enzyme is higher in
larvae reared on the artificial diet. In insects, mixed-
function oxidases often affect the duration and intensity
of insecticide action and thus influence insecticide meta¬
bolism, detoxification, and resistance.

Similarly, Bush (1978) discovered significant differences
in the genetic makeup of wild screwworm flies, particular¬
ly in certain genes controlling enzymes involved in flight
activity. One very important enzyme, a-GDH (a-glycerol
phosphate dehydrogenase) exists in two different forms.
The factory-reared flies have contained mostly a-GDH!.
And G. B. Kitto (unpublished data) has shown that the
two forms of a-GDH have optimal activity at different
temperatures; a-GDH2 is less active than a-GDH, in the
temperature range experienced in nature. The high con¬
stant temperature used to speed development in the fac¬
tory has apparently been exerting a strong selective force
favoring a-GDH2 over a-GDH,. This finding also sug¬
gests that the competitive ability of the fly in nature
might decrease as the frequency of a-GDH2 increases, be¬
cause factory flies would have to cope with a wide tem¬
perature range in nature; those individuals lacking the
a-GDH, enzyme simply would not be able to fly as well
as wild individuals.

Dietary effects on vision. — Not much is known about how
diet affects insect vision. Tryptophane, an important
amino acid, is metabolized to ommochromes, a group of
vital pigments associated with screening in the insect

36

eye. Kayser (1979) has demonstrated that the trypto¬
phane requirement in P. brassicae is very precise; lower
amounts result in fewer metabolites essential for normal
vision. Vitamin A is another nutrient associated with vi¬
sion in many insects and is therefore another important
constituent of the diet. Agee (1979) reported on an instru¬
ment that measures the visual sensitivity of insects and
could be used to monitor these differences. So, more
research is needed on the relationship of dietary ingre¬
dients and insect vision and on means of monitoring
changes in this essential attribute.

Host insects in production

of parasites and pathogens

Parasite production.— In parasite production, the diet de¬
termines how well the host’s nutritional needs are met,
which in turn may influence the quality of parasites pro¬
duced on host insects. For example, when the Asiatic rice
borer was reared on artificial diet, its parasite, Apanteles
chilonus Munakata, was adversely affected (Kajita 1973).
Likewise, Etienne (1973, 1974) reported that the tachinid
Lixophaga diatraeae (Townsend) cannot be continuously
reared on larvae of an unnatural host— the greater wax
moth, Galleria mellonella (Linnaeus)— that has been fed
beeswax and pollen unless the host diet is supplemented
with vitamin E or wheat germ. In this case, problems in
rearing L. diatraeae on greater wax moth larvae were
eliminated by fortifying the diet with a high-protein
cereal and the addition of 120 g of wheat germ per
kilogram of diet (Morrison and King 1977, King et al.
1979). Morrison and King (1977) concluded from this
result that “suitability cannot be determined merely by
screening hosts, but host nutrition and other factors
must also be considered, and compromises between ento-
mophagous arthropod quality and quantity may have to
be made because of cost and the numbers required.” Cur¬
rent and future research should solve similar problems
for other hosts and parasites.

Some progress has been made in rearing entomophagous
species (Hoffman et al. 1975) on medium without host in¬
volvement; but this type of rearing will be more common
and probably more economical in the future. This will be
a major advance in parasite production for suppression
programs.

Pathogen production. — Rearing larvae on artificial diet to
produce virus has many advantages over the conven¬
tional leaf-feeding method. For example, the diet can be
sterilized to avoid contamination by other microbes, the
need for growing plant material is eliminated, and
measured doses of the virus can be given more conven¬
iently to the larvae. Tanada (1965) and Helms and Raun
(1971) have found that nutrition is important in the sus¬
ceptibility of insects to virus. Shvetsova (1950) reported

that greater wax moth larvae are most susceptible to nu¬
clear polyhedrosis virus when fed wax enriched with
nitrogen and carbohydrate. Pimentel and Shapiro (1962)
had similar results with a high-nitrogen diet but found
that nuclear polyhedrosis virus does not increase when
greater wax moth larvae are fed standard diet or extra
carbohydrates. The incidence of virus infection can also
be increased by a reduction in certain diet ingredients
(David et al. 1972). For example, when sucrose is omitted
or the content of casein reduced from a semisynthetic
diet for Pieris brassicae (Linnaeus), the incidence of
granulosis virus disease increases (David and Taylor
1977). Similarly, Shapiro et al. (1978), in examining how
diet influences production of nuclear polyhedrosis virus
from gypsy moth, found that the total virus yield varies.
Virus production is more economical from moths reared
on a diet with a high concentration of wheat germ. An in¬
crease in vitamin concentration improves virus yield
slightly, but few differences are caused by change in pH
from 4 to 7.

Clearly, pathogen production is influenced by the nutri¬
tional status of the host, which in turn depends on diet.
As with parasite production, then, formulations of diets
specifically for pathogen production should receive con¬
siderable attention from researchers in the future.

Development of optimum diets

for new species

Analysis of the reference listings in Singh (1977b) shows
that fewer than 1,000 insect species have been reared on
artificial diets. Only 160 pre-1950 references were found,
but there were more than 2,300 references up to 1978,
and the list continues to grow. Most species reared on ar¬
tificial diet are from the orders Lepidoptera, Coleoptera,
and Diptera. Though there are 45 species listed under the
order Hymenoptera, 32 of these are ants, and 13 of the
ant species have been reared for only part of their life cy¬
cle. A review of Singh (1972, 1977a, 1977b) shows that
only about two dozen or so species have been successfully
reared for several generations. Also, diets for predators,
parasites, and blood-feeding insects have received much
less attention than diets for plant pests. Though tre¬
mendous advances have been made in colonizing some
species, much work still remains to be done in developing
the best artificial diets and rearing techniques for new
species as well as those already colonized.

Standardization of insect

diets and rearing methods

It is time to standardize diets, at least for commonly used
test insects. Standardization of insect diets is important
because the biological (for example, life cycle and fecundi¬
ty) and chemical (for example, pheromone, hormone,

37

biochemical, and enzyme) characteristics of the reared in¬
sect depend partly on nutrition. So quality standards must
be applied to the formulation of artificial diets, including
the origin and analyses of commercial ingredients. Diets
should be prepared by standard methods and the following
variables recorded: percentage of protein, carbohydrate,
lipid, minerals, and vitamins; osmolarity; pH; texture;
moisture content; microflora activity; use of mold in¬
hibitors; and other secondary substances.

Literature Review

In Singh (1977b), I reviewed the literature on artificial
diets for insects, mites, and spiders from 1900 to 1976.
Diets were listed for more than 750 species collated from
nearly 2,000 references. (See also Singh 1972.) Insect diets
have been cataloged in recipe books by House (1967),
House et al. (1971), Singh (1974b, 1977a, 1977b) and
Gomez (1978a). (See also Gomez’s 1978b bibliography of
artificial diets for Lepidoptera.) Wyniger (1974) listed some
artificial diets. Various matters related to diet have been
reviewed by Yushima (1962), Vanderzant (1966, 1969a,
1974), Boness (1968, 1969, 1970), McKinley (1971), and
Gardiner (1978). Five general insect-rearing books (Ishii
1959; Needham 1959; Smith 1966, 1967; and Joint
FAO/IAEA Division of Atomic Energy in Food and Agri¬
culture 1968) have useful discussions of artificial diets.

Insect nutrition has been exhaustively reviewed by House
(1961a, 1962, 1965, 1972, 1974), Patton (1963), David
(1967), Guennelon (1967), Gordon (1968), Chapman (1969),
Dadd (1970, 1973), Hodgson and Rock (1971), N. C. Pant
(1973a), and Levinson (1976). Specialized reviews are avail¬
able on honey bees, Apis mellifera (Linnaeus), by Haydak
(1970); on the silkworm, Bombyx mori (Linnaeus), by Ito
(1967, 1972, 1979); on Drosophila by Sang (1978); on
Diptera by Friend (1968); on phytophagous insects by
Friend (1959), McGinnis and Hastings (1964), Beck and
Chippendale (1968), and J. C. Pant (1973); on locusts and
grasshoppers by Dadd (1963); on aphids by Auclair (1963),
Yushima (1968), Massonie (1971), and Kunkel (1977); on
muscoid flies by Spiller (1964); on plant-sucking insects by
Auclair (1969); on natural enemies by House (1977); and on
stored-product pests by Punj and Girish (1968) and Misra
(1973). Altman and Dittmer (1968) and Dadd (1977) give
comprehensive tabular summaries of quality requirements
for insect nutrition. And Rodriguez (1972) presents several
papers on the significance and implications of insect nutri¬
tion in ecology and pest management.

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1960. A wheatgerm medium for rearing the pink

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Agee, R. H.

1979. Eye clinic for insects. Agric. Res. 18(2): 12-13.

Ahmad, S., and Forgash, A. J.

1978. Gypsy moth mixed-function oxidases: gut en¬
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71: 499-452.

Altman, P. L., and Dittmer, D. S. (eds.).

1968. Metabolism. 737 pp. Federation of American
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Atallah, Y. H., and Newsom, L. D.

1966. Ecological and nutritional studies on Coleo-
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Auclair, J. L.

1963. Aphid feeding and nutritiqn. Annu. Rev. En¬
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Berger, R. S.

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Buchner, P.

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tomol. 102: 1304-1306.

Yushima, T.

1962. Rearing of insects on artificial diets— present
and future. Nogyo Gijutsu 17: 172-175,
212-215, 269-273, 314-317, 369-372,

419-422, 526-529, 581-586. (In Japanese.)

1968. Rearing methods of aphids on artificial diets.
Shokubutsu Boeki 22: 306-310. (In Japanese.)

Zabinski, J.

1926. Observations sur l’elevage des cafards nourris
avec des aliments artificiels. C.R. Soc. Biol.

95: 545-548. (In French.)

1928. Elevage des blattides soumis a une alimenta¬
tion artificielle. C.R. Soc. Biol. 98: 73-77. (In
French.)

44

Ingredients for Insect Diets

Quality Assurance, Sources,
and Storage and Handling

By F. D. Brewer1 and Oliver Lindig2

Introduction

Analysis of an insect’s natural food source often suffices
as a basis for formulating an artificial diet for the insect.
But an acceptable artificial diet must have more than the
right ingredients. The ingredients must be of the quality
needed to rear the insect for a particular purpose. Of
course, even if the diet is imperfect, it may be satisfac¬
tory, and the insects may adapt to it quickly (Davis 1972)
whatever the sources of the dietary components (proteins,
fats, etc.). This paper discusses ways to assure ingredient
quality, sources of dietary components, and general con¬
siderations for storage and handling of diet ingredients to
maintain quality.

Assuring Quality
of Diet Ingredients

Most processors and suppliers of diet ingredients will fur¬
nish, on request, fairly detailed technical data on their
products, assurances of product quality, and manufac¬
turers’ recommendations on shelf life and the best
storage conditions. But many diet components* such as
laboratory-processed plant materials or feed-grade ingre¬
dients, have limited or no technical data available. So the
user is often forced to “take it or leave it.”

Many analytical tests, standardized by the Association of
Official Analytical Chemists (A.O.A.C.) in their Official
Methods of Analysis (1975), are available to test the
quality of dietary ingredients. But type, frequency, and
amount of testing are generally dictated by what the in¬
sect is being reared for, type of program, number and
qualifications of employees, and availability of space and
equipment. If, for example, the diet is chemically defined
and the insect reared is used in metabolic or nutrient-
deletion studies, the dietary ingredients will have to be

‘Research entomologist, Southern Field Crop Insect Manage¬
ment Laboratory, Agricultural Research Service, U.S. De¬
partment of Agriculture, P.O. Box 225, Stoneville, Miss. 38776.

“Research entomologist, Boll Weevil Research Laboratory, Agri¬
cultural Research Service, U.S. Department of Agriculture, P.O.
Box 5367, Mississippi State, Miss. 38762.

tested extensively. But, if the insect is mass-reared for
pathogen production on a crude diet containing plant
material or minimally processed feed-grade material,
fewer tests will be needed. Chemical analyses might in¬
dicate an adequate supply of an essential nutrient, such
as an amino acid, but the processing may have caused
nutrient loss or made some nutrients unavailable to the
insect. In these instances, bioassays can be used after all
processing and possible ingredient interactions have oc¬
curred. Finally, physical characteristics of the prepared
media, such as pH, viscosity, color, and texture, can be
checked.

Sudden changes, however minor, in ingredient quality
may affect insect colonies dramatically. So, if the rearing
program is sufficiently large or critical, all incoming
shipments of ingredients should be tested. This testing
could be especially important if suppliers or manufac¬
turers are changed. In the future, specifications for
nutrient content and product purity may be developed for
insect diets. But, not having such standards, rearing pro¬
grams must do careful testing.

Most commercial suppliers of food products, especially
those used for human consumption, are regulated by the
Federal Food, Drug, and Cosmetic Act administered by
the U.S. Food and Drug Administration. The suppliers, in
turn, maintain strict quality control in accordance with
the A.O.A.C. These detailed and extensive analyses in¬
clude protein content (NX 6. 25); amino acid composition;
and mineral, vitamin, heavy metal, and microbiological
profiles. Physical properties of the materials are also in¬
cluded in the technical data from the processors. Of
these, size or fineness of the material is the major item.
Other information includes weight per given volume
(usually in cubic feet) or density, free-flowing quality of
dry powder (sliding or bridging characteristics), type of
container, and weight of standard packaging. The pH of
single ingredients may also be critical to the acceptability
of the final insect diet. For example, two methods of pro¬
tein isolation (precipitation) are generally used in pro¬
cessing soy protein. One is acidic, producing a protein
with a pH of about 5 (1 : 10 aqueous dispersion) and the
other basic, with the protein having a pH of about 7.
Depending on the quantity used, pH could significantly
affect the activity of microbial inhibitors such as methyl-
paraben (methyl p-hydroxybenzoate) or sorbic acid. Indi-

45

vidual ingredients, especially alfalfa meal and wheat
germ, should be monitored for chemical contamination
from pesticides. Also, the ingredients should be checked
for insect pathogens that could introduce disease into the
colony. Further chemical analysis may not be needed ex¬
cept for the most stringent metabolic or nutrient-deletion
studies.

Bioassay would probably be the chosen method of quality
assurance for most insect-rearing operations. Bioassay is
preferable to chemical analysis mainly because it meas-
sures how the ingredient affects the product (the insect)
rather then simply defining the ingredient. And sophisti¬
cated equipment, highly trained technicians, and sup¬
portive laboratory space generally are not required for
bioassays as they are for chemical analysis. Bioassay is
also a measure of the insect’s ability to digest and assim¬
ilate the nutrients. For example, Davis (1972) reported
different growth rates in yellow mealworm, Tenebrio
molitor Linnaeus, reared on casein obtained from four dif¬
ferent suppliers. In such cases, both biological and chemi¬
cal analyses would be required to determine if the amino
acid composition was similar for like ingredients or
whether methods of processing made one or more of the
amino acids inadequate or inaccessible to the insects.

Common Sources
of Diet Components

Proteins

Casein (minimum of 14.5% total dry nitrogen) is a com¬
mon protein source in artificial diets for insects because
it is highly purified and readily available. (See appendix
for a comprehensive list of diet ingredients and some sup¬
pliers.) For example, according to ICN Nutritional Bio¬
chemicals’ catalog for 1979/80, their entire manufacturing
process is rigidly controlled, and the final product is sub¬
jected to biological and chemical assays. Maximums for
total moisture, ash, and ether-extractable fat are met, as
are specifications for total nitrogen (protein), amino acid
profile (A.O.A.C. 1975), concentration of hydrogen ions
(pH), and amount of lactic acid. The micrograms of
B-vitamins per gram of casein are assayed and given on
every batch or lot of Vitamin-Free Casein. Microbial
limits are also established per gram of material (for exam¬
ple, standard plant count <5,000 colonies, coliform <50
colonies, yeast and mold < 250 colonies, and Salmonella
spp. and Escherichia coli negative). Apparently,
bioassays are the most sensitive means of determining
minor variations in the amino acid concentration of ca¬
sein (Davis 1972) and other primary protein sources such
as vegetable proteins and oilseeds that are deficient in
one or more of the essential amino acids (Agricultural
Research 1979).

Within the last decade, toasted, defatted soyflour
(minimum of 50% protein) and cottonseed materials have
been substituted for casein in Heliothis spp. diet. Purified
mixtures of essential amino acids, plant extracts, wheat
germ, animal lean meat and organs, egg albumen, and
microbial protein concentrates have also been used to
supply protein to the diet. According to the Archer
Daniels Midland Co. (personal communication), heat
treatment is used at various stages in the making of their
soyflour toasted products. During this production, a pro¬
tein dispersibility index is measured to insure the correct
range or protein level (minimum of 50%) of the product
before it is stored. Other tests done during these produc¬
tion steps may include analyses of moisture content, oil
content, crude-fiber composition, urease activity, and par¬
ticle size. Urease activity is a good indicator of trypsin-
inhibitor level— the higher the level of urease, the greater
the trypsin inhibition. In addition to routine chemical
analysis, the soyflour also receives a microbiological
assay similar to the one for casein. In diets based on
wheat germ, both chemical and biological assays may be
used on the wheat germ, which contributes protein and
crude fiber to the diet.

Carbohydrates

Dextrose, sucrose, and fructose are added to the diet as
major carbohydrate sources. Others may include honey,
starch, cellulose, cereal grains, etc. Appropriate A.O.A.C.
tests for types and amounts of sugar (both reducing and
nonreducing) and starch materials may be run directly on
these materials and on the wheat germ, since it also con¬
tributes carbohydrates.

Fats

Wheat germ, lecithin, cephalin, cholesterol and other
sterols, grain oils, lard, and glycerol are probably the
most common sources of fats in artificial diets. Some use
purified essential fatty acids such as linoleic acid and
linolenic acid. Wheat germ oil furnishes dietary lipids.
Measuring the peroxide value of the extracted oil is an
excellent method of determining its rancidity. According
to Bio-Serv (personal communication), two widely used
methods for determining the peroxide value (<70 milli-
equivalents/kg) are the active-oxygen method (Eastman
Food Laboratory 1973) and a modification of the Wheeler
(1932) method. Other grain oils, such as corn oil, are
checked similarly.

The toasted wheat germ is also microbiologically assayed
like the casein and soyflour (Bio-Serv, personal communi¬
cation). Specifications for raw wheat germ and other
natural ingredients, such as corncob grits, soybean meal,
etc., are not as rigid as they are for casein and soyflour.

So the microbial load may fluctuate from 50,000 to 1

46

million spores /g, with a high incidence of coliform and
fungal contaminants. As these high microbial loads are
intrinsic to wheat germ, lots of raw wheat germ are gen¬
erally accepted for industrial processing once they have
passed rancidity tolerances.

Supplements

Salt or mineral mixtures, such as Wesson salts, are com¬
monly added to the diet to supply the minerals needed
for growth and development. These necessary ingredients
are also found in wheat germ, ash of whole meat, fresh
plant materials, and even water. Salt or mineral sources
are analyzed quantitatively by appropriate A.O.A.C.
tests, and some qualitative scanning for heavy metals
and chlorinated hydrocarbons is also done.

Wheat germ, brewers’ yeast, custom-formulated pre¬
mixes, and mixtures of the B-vitamin complex are usually
added as supplements. Chemicals such as ascorbic acid,
choline chloride, and a-tocopherol are often added sep¬
arately. Potency tests are conducted to establish pharma¬
ceutical grade according to established procedures
(Hoffman-LaRoche, personal communication). Techniques
such as high-pressure liquid chromatography, which is
quick and sensitive for such determinations, are de¬
scribed in the United States Pharmacopeial Convention’s
(1975) United States Pharmacopeia (U.S.P.) and A.O.A.C.
Ascorbic acid has to conform to U.S.P. specifications
(Bio-Serv, personal communication).

Chemicals such as mold inhibitors (for example sorbic
acid), methylparaben, folpet3, benomyl4, propionic acid,
phosphoric acid, Formalin (formaldehyde), and antibiotics
are added to the diet in various combinations. Ingredi¬
ents such as methylparaben must conform to the U.S.P.;
sorbic acid must conform to the National Academy of
Sciences’ (1972) Food Chemicals Codex, and Aureomycin
(chlortetracycline) must give a positive result for Quali¬
tative Test according to A.O.A.C.

Another kind of supplement is food gum. It can be one of
a diversified group of materials, including seaweed ex¬
tracts such as agar, algin, carrageenan, and furcellaran;
tree exudates and extracts such as gum arabic and traga-
canth; seed gums such as guar and locust bean; cellulose
derivatives such as carboxymethyl cellulose and micro¬
crystalline cellulose; and microbial gums such as xanthan
(Howell 1977). The function of these materials is also

W[(Trichloromethyl)thio]phthalimide.

‘Methyl l-(butylcarbamoyi)-2-benzimidazolecarbamate.

diversified; for example, uses include adhesive and gelling
(agar), bulking (gum arabic), suspending (carrageenan),
swelling and inhibition of syneresis (guar), and binding
(locust bean gum). Agar is typically analyzed by sup¬
pliers, harvesters, and manufacturers for moisture
(16%-17% normal range), total ash (1.5%), acid-insoluble
ash (0.07%— 0.09%), pH (7.2), gel strength for a 1.5% solu¬
tion (730-800 g/cm2), mesh distribution, and clarity of a
hot 1.5% solution (Perny, personal communication). Also,
a microbiological assay is done periodically; but microbes
are rarely a problem for agar, which usually is negative
for E. coli (0-10 range), has no Staphylococci spp., and
has a standard plate count of 2,000-12,000 spores/g.

There are three commercially available plant-based agar
types— Gelidium spp. (gelation point 34°-36° C), Grassi-
laria spp. (gelation point: 42°-45° C), and Terocladia spp.
(not used widely; gelation point: 34°-36° C). The type of
seaweed collected and even where it comes from affect
the quality of extracted agar (Perny 1971). Most agar is
currently processed outside the United States. The most
common method of determining its gel strength is the
Kobe test, which is performed on a 1.5% solution and
measured in grams per square centimeters.) The Fira test
measures deformation and fracturing of the agar, and gel
strength determines the final firmness and resilency of a
completed diet.

Certain nonnutritive materials, such as alphacel and corn¬
cob grits, may be added to diets to increase bulk and re¬
place some of the agar. For example, corncob grits (crude
cellulose or fiber) were used to replace 80% of the agar in
a casein and wheat germ diet for rearing Heliothis spp.
larvae (Raulston and Shaver 1970). This filler material
was monitored by the manufacturer for water absorption,
capacity for carrying water and oils, capacity for holding
water, and general physical properties (Foley 1978). These
physical properties included bulk density, specific gravi¬
ty, particle size, and flowability. Solubility in water and
organic solvents was also tested. And elements such as
structural and nonstructural polysaccharides were ana¬
lyzed. Data were given for nutritive components such as
protein, fat, crude fiber, ash, and vitamins, as were data
for biodegradability, microbiological assay, pesticide
residues, and combustibility.

Storage and Handling

There is probably no substitute for freshly processed in¬
gredients, especially if they are perishable. But generally,
a cool, dry area (for example, 15.6° C and 40% relative
humidity) will suffice for prolonged storage. Some ingre¬
dients, such as wheat germ and soyflour, are packaged in
large quantities (for example, 50- and 100-lb lots), so it is
necessary to have adequate storage space. And some in¬
gredients require refrigeration. Storage conditions should

47

be designed to provide maximum shelf life for the
ingredients.

Vitamin sources (or mixtures) and purified unsaturated
fatty acids are among the most perishable ingredients
used in insect diets. Refrigeration and desiccation extend
the shelf life of vitamins; and fatty acids may be pro¬
tected from peroxide formation by storing them in a
100%-nitrogen atmosphere. Wheat germ is also a perish¬
able item that should be stored in a moisture-proof con¬
tainer; so it is often purchased in small quantities
packaged in vacuum-sealed containers. Moisture-proof
containers are recommended for storing most dietary in¬
gredients— casein, sucrose, inhibitors, agar, minerals, etc.

Lot or batch numbers are provided by most suppliers,
but receiving date should be placed on all incoming in¬
gredients. If the ingredients are divided into smaller
quantities or used to prepare premixed diet, the date
received and lot number should be placed on all con¬
tainers. Ingredient changes in the middle of a project
should be avoided. Ingredients should be handled, mixed,
ground, sifted, etc., in well-ventilated areas, because
hazardous dusts may be produced. Equipment and
machinery used in ingredient processing and handling
should be easy to clean and disinfect.

Acknowledgments

We thank the following companies for their time and ef¬
fort in supplying valuable information for this paper:
Archer Daniels Midland, Decatur, Ill.; Bio-Serv, French-
town, N.J.; Hoffman-LaRoche, Nutley, N.J.; and Perny,
Ridgewood, N.J.

References

Agricultural Research.

1979. Tailoring vegetable proteins. 18(1): 7.
Association of Official Analytical Chemists.

1975. Official methods of analysis. 12th ed., 1094 pp.
The Association, Washington, D.C.

Davis, G. R. F.

1972. Refining diets for optimal performances. In J.
G. Rodriguez (ed.), Insect and Mite Nutrition,
pp. 171-181. North-Holland Publishing, Am¬
sterdam.

Eastman Food Laboratory.

1973. Active oxygen method for comparing fat and
oil stabilities. 5 pp. Eastman Food Laboratory
Standard Procedure #6A. Eastman Publ.
YZ-159A.

Foley, K. M.

1978. Chemical properties, physical properties and
uses of the Andersons’ corncob products. 425
pp. Andersons, Maumee, Ohio.

Howell, J. F.

1977. Codling moth: agar substitutes in artificial

diets. U.S. Agric. Res. Serv. [Rep.] ARS-W-43,
9 pp.

ICN Nutritional Biochemicals.

[Catalog for 1979/80], 37 pp. Cleveland, Ohio.
National Academy of Sciences.

1972. Food chemicals codex. 2d ed., 1039 pp. The
Academy, Washington, D.C.

Perny.

1971. All agar-agar is not alike, 14 pp. Ridgwood,

N.J.

Raulston, J. R., and Shaver, T. N.

1970. A low agar casein-wheat germ diet for rearing
tobacco bud worms. J. Econ. Entomol.

63: 1743-1744.

United States Pharmacopeial Convention.

1975. The pharmacopeia of the United States of
America (the United States pharmacopeia).

19th rev., 824 pp. Mack Publishing, Easton, Pa.
Wheeler, D. A.

1932. [Untitled article.] Oil and Soap 9: 89.

Appendix.— Insect-Diet
Ingredients and Some
of Their Sources

The list below gives commonly used diet ingredients not
likely to be available locally. Each item is listed by non¬
proprietary name. Proprietary names and other manufac¬
turer identifications, if any, descriptions, uses, amounts,
etc. are given in parentheses. Names in brackets are short
forms for manufacturers and suppliers. Full names and
addresses, keyed to these abbreviations, follow the list.
The main source for this information was Frass, the In¬
sect Rearing Group newsletter.

Diet ingredients

Acetic acid (glacial) [Balter].

Agar (fine) [Burtonite, Morehead], (granulated) [Bio-Serv],
(type 100 cm) [Perny], several grades [U.S. Biochemi¬
cal].

Alfalfa meal [Nutrilite].

Alphacel (nonnutritive bulk) [ICN],

Ascorbic acid [Roche].

Beeswax (dark crude) [Sioux],

Benomyl (Benlate) [DuPont],

Bone flour (purified) [ICN], Blended Food Product, Child
Food Supplement, Formula No. 2 or CSM (corn, soy-
flour, milk) [Krause].

Brewers’ yeast (debittered) [Vitamin Food].

Calcium alginate (Kelgin HV) [Kelco].

Calco oil red dye (American Cyanamid’s A-1700) [Cyan-
amid].

Carboxymethyl cellulose (sodium salt) [ICN].

48

/5-Carotene (crystalline) [ICN],

Carrageenan (Gelcarin HWG) [Marine Colloids].

Casein (crude, 80 mesh; lactic, 30-40 mesh) [Milk Special¬
ties, New Zealand, Erie], (Vitamin-Free) [ICN].

Cephalin [ICN],

Chlortetracy cline soluble powder (Aureomycin; 25.6 g
a.i./6.5-oz package) [Lederle, Ozark].

Cholesterol (U.S.P. grade) [Bio-Serv],

Choline chloride [ICN],

Corncob grits (60 grade) [Andersons].

Corn oil [ICN],

Cottonseed meal (41% solvent extracted) [Yazoo], (Phar-
mamedia) [Traders].

Cottonseed meats [Yazoo].

Dextrin [ICN],

Dextrose [ICN],

Egg albumen [ICN],

Ethanol [U.S. Industrial].

Fibrin [ICN],

Fish meal [ICN],

Folpet (Phaltan) [Chevron].

Formaldehyde (40% solution) [Fisher],

Fructose (U.S.P. grade) [Bio-Serv].

Fumaric acid [ICN].

Gluten, wheat [ICN],

Glutathione [ICN],

Glycerol (chemically pure) [ICN].

Glycogen (beef) [ICN],

Guar gum [ICN],

Gum arabic (acacia) [ICN],

Inositol [ICN],

KOH (45% potassium hydroxide solution) [Fisher],
Lactalbumin [ICN],

Lecithin [ICN],

Linoleic acid (65% solution) [ICN],

Linolenic acid (55% solution) [ICN],

Locust bean gum [Bio-Serv].

Maltose [ICN],

Menadione (vitamin K3) [Bio-Serv].

Methylparaben (methyl-para hydroxybenzoate, technical)
[Sigma, Tenneco 1, Tenneco 2],

Milk powder (whole 28%) [Bio-Serv], (whole) [ICN].
Penicillin G [ICN],

Peptone [ICN].

Potassium sorbate powder [Aceto-Chemical], (Sorbistat
K) [Pfizer 1],

Promine D (90%-95%, protein isolate) [Jeffards].

Propionic acid [ICN].

Propyl gallate (Tenox PG) [Eastman].

Sorbic acid [Pfizer 2, Sigma].

Sorbose (L form) [ICN],

Soybean oil [ICN],

Soyflour (toasted, Nutrisoy) [Archer],

Soy protein (Supro 610, 100 mesh) [Ralston],

Streptomycin sulfate [ICN].

Tocopherol acetate (vitamin E, DL-Alpha) [Bio-Serv],

Torula yeast [Bio-Serv, ICN, St. Regis].

Tragacanth [ICN],

Vitamin premix (Roche No. 26862) [Roche],

Wesson salts (mineral mix, also known as Salt W) [ICN],
U.S. Biochemical], (modified) [Bio-Serv],

Wheat bran [ICN],

Wheat germ (raw) [Earthwonder], (raw, flaked) [Niblack],
(toasted, flaked or regular) [Kretchmer],

Wheat germ oil [Bio-Serv],

Whole wheat flour [ICN],

Xanthan [ICN],

Zein [Bio-Serv].

Sources

Aceto-Chemical:

Aceto-Ch6mical Co.

Flushing, N.Y.

American Cyanamid:

American Cyanamid Co.

Houston, Tex.

Andersons:

The Andersons
Maumee, Ohio.

Archer:

Archer Daniels Midland Co.

Decatur, Ill.

Baker:

J. T. Baker Chemical Co.

Phillipsburg, N.J.

Bemhem:

c/o South Western Sales Association
Jacksonville, Fla.

Bio-Serv:

Bio-Serv, Inc.

Frenchtown, N.J.

Burtonite:

Burtonite Co.

Nuttley, N.J.

Chevron:

Chevron Chemical Co.

San Francisco, Calif.

DuPont:

E. I. duPont de Nemours Co.

Wilmington, Del.

Earthwonder:

Earthwonder
Springfield, Mo.

Eastman:

Eastman Chemical Products
Kingsport;, Tenn.

Erie:

Erie Casein Co.

Erie, Ill.

Fisher:

Fisher Scientific

49

St. Louis, Mo.

ICN:

ICN Nutritional Biochemicals
Cleveland, Ohio.

Jeffards:

Doug Jeffards Co.

Nashville, Tenn.

Kelco:

Kelco Co.

Chicago, Ill.

Krause:

Krause Milling Co.

Milwaukee, Wis.

Kretchmer:

Kretchmer Products
International Multifoods Corp.
Minneapolis, Minn.

Lederle:

Lederle Laboratory
Dallas, Tex.

Marine Colloids:

Marine Colloids, Inc.

Springfield, N.J.

Milk Specialties:

Milk Specialties
Dundee, Ill.

Morehead:

Morehead & Co.

Van Nuys, Calif.

New Zealand:

New Zealand Milk Products, Inc.
Rosemont, Ill.

Niblack:

Niblack Foods
Rochester, N.Y.

Nutrilite:

Nutrilite Products
Lakeview, Calif.

Ozark:

Ozark Supply Co.

American Cyanamid
Kansas City, Mo.

Perny:

Pemy, Inc.

Ridgewood, N.J.

Pfizer 1:

Pfizer Corp.

Chicago, Ill.

Pfizer 2:

Pfizer, Inc.

Doraville, Ga.

Ralston:

Ralston Purina Co.

St. Louis, Mo.

Roche:

Roche Chemical Division
Hoffman-LaRoche, Inc.
Nutley, N.J.

St. Regis:

St. Regis Paper Co.
Rhinelander, Wis.

Sigma:

Sigma Chemical Co.

St. Louis, Mo.

Sioux:

Sioux Honey Association
Sioux City, Iowa.

Tenneco 1:

Tenneco Chemicals, Inc.
Organics & Polymers Division
Piscataway, N.J.

Tenneco 2:

Tenneco Chemical Co.

Chicago, Ill.

Traders:

Traders Oil Mill Co.

Fort Worth, Tex.

U.S. Biochemical:

U.S. Biochemical Corp.
Cleveland, Ohio.

U.S. Industrial:

U.S. Industrial Chemical Co.
Louisville, Ky.

Vitamin Food:

Vitamin Food Co., Inc.
Newark, N.J.

Yazoo:

Yazoo Valley Oil Mill
Greenwood, Miss.

50

Containerization for Rearing Insects

By Robert L. Burton1 and W. Deryck Perkins2 3

History

Many different kinds of containers have been used for
rearing insects. The containers have been selected for
their suitability and their availability from a multitude of
widely used items. For example, Smith (1966) listed over
40 different containers used in many kinds of rearing pro¬
grams, including milk bottles, battery jars, plastic dishes,
metal trays, glass jars of all sizes, aquariums, trash cans,
and barrels of various sizes (fig. 1). As the technology of
insect rearing moved from natural diets to artificial diets,
containerization became more standardized. Singh (1977)
briefly surveyed 75 species from 50 families and 10
orders and found that 34% were reared in plastic cups;
22% in glass vials; 17% in petri dishes; 10% in fruit jars;
and the remainder in test tubes, plastic or metal trays,
flasks, plastic bags, plastic boxes or dishes, cardboard
containers, salve cans, garbage cans, gelatine capsules,
ice cream cups, multicell containers, and paper straws.
The 1-oz polystyrene plastic condiment cup has been one
of the more popular containers for use with synthetic
diets. These cups serve well in both small and relatively
large programs. They are fairly inexpensive, conveniently
disposable, readily available, provide a good microen¬
vironment, can be used in semiautomated equipment, and
allow for isolation of cannabalistic species. Where species
are not totally cannabalistic, another popular container,
the paper drink cup, has been used for rearing insects in
aggregate; these have many of the advantages of the
plastic cups. Petri dishes have also been widely used for
aggregate rearing. For axenic culture of insects, glass
shell vials with cotton plugs have been used. For rearing
stored-product pests on both natural and synthetic diets,
glass jars of various sizes have been used.

In a few cases, containers have been designed and de¬
veloped by researchers specifically for insect rearing. The
need for such containers may result from cost, conven¬
ience, or species uniqueness. For example, Raulston and
Lingren (1969), Morrison et al. (1975), and Hartley et al.

'Research entomologist. Wheat and Other Cereal Crops
Research Unit, Agricultural Research Service, U.S. Department
of Agriculture, P.O. Box 1029, Stillwater, Okla. 74076.

"Research entomologist, Southern Grain Insects Research
Laboratory, Agricultural Research Service, U.S. Department of
Agriculture, Tifton, Ga. 31793.

"Journal article 3757 of the Agricultural Experiment Station,
Oklahoma State University, Stillwater, Okla. 74078.

(1982) have developed multigrowth grids for lepidopterans.
Peterson (1964) describes the design and construction of
a multitude of cages and containers used specifically for
insects. Other specific rearing containers are described by
Baumhover et al. (1977), Harrel et al. (1973), and Leppla
et al. (1975).

As the technology of insect rearing moves forward, the
adequacy of the rearing container will be examined more
strenuously. Improvement of insect containerization re¬
quires an indepth look at the related aspects of container
design. This paper discusses these aspects and other con¬
siderations that might be needed in developing or im¬
proving an insect-rearing container.

Design and Selection
of the Rearing Container

General considerations

An effective container must protect the food; present the
food to the insect in an acceptable manner; provide the
proper surfaces and atmosphere for the insect; and con¬
fine, and, in some cases, separate cannabalistic insects.
Above all else, the rearing container must meet the phys¬
iological and ecological needs of the insect whether it
be environment, space, or other. At the same time, the
container must have certain general structural qualities
and specifications that conform to the activities of the
insect-rearing program such as filling, implanting insects,
storing, handling, and cleaning.

Physiological

and ecological

considerations

Size, shape, and surface.— The size of the container af¬
fects the insect in several ways. If wandering by larvae
must be prevented or reduced, the container must hold
the insect close to the diet. Population density, which
depends partly on container size, can affect such factors
as growth and fecundity. All containers must provide
enough space to permit unrestrained larval development
and pupation. Container size is important in rearing part¬
ly cannibalistic species because sufficient space reduces
this activity. When insects can be reared in aggregate,
each species probably has an optimum container size.

The shape of a container is also important. Shape may af¬
fect diet thickness or the ratio of total volume to surface
area of the diet and thus influence moisture retention.
Shape may affect pupation sites and access to the diet.

51

Figure 1.— A variety of containers that have been used for rearing insects.

Proper surfaces must be available for unrestrained
molting and pupation. For example, in many container
types, the unwaxed closure surface has traditionally pro¬
vided a surface for these activities. Both size and shape
affect air and moisture exchange, because air movement
around and between containers may be important for
proper ventilation.

Air and moisture exchange. — Air and moisture exchange
may be the most important physiological and ecological
function of the rearing container. The insect must have a
favorable microenvironment, which can be produced only
if ventilation and moisture regulation in the container
meet the insect’s maximum respiratory demands (these
may be difficult to determine) and moisture requirements
(which will vary among species). Controlled water-vapor
transmission can regulate the drying rate of the diet.
Also, and usually more important, it can eliminate ex¬
cessive moisture buildup in the container. Free moisture
promotes spore germination and growth of unwanted mi¬
crobial contaminants and is one of the primary causes of
these epizooics. Traditionally, the closure has provided

the ventilation (see “Closure Requirements”) since the
container must contain the wet diet and thus be resistant
to water movement. Common techniques for air and mois¬
ture regulation have been the use of closure materials
that ventilate (such as unwaxed paperboard) and regula¬
tion of the size of a ventilation area covered with screen,
fabric, etc. Conditions outside the container then become
important in the regulation of moisture in the container.
Moisture requirements of the microclimate vary from
species and among the various life stages of the insect
and require that optimum levels be known for each insect
to be reared.

Toxicity.— Certain plastics, such as cellulose acetate, can
be toxic to some species of insects (Chada 1962). Also,
some types of wood may be harmful. Some paints, var¬
nishes, and preservatives have toxic properties, especially
when freshly used. All these substances can also cause
toxic reactions in plants and thus affect the insects if the
plants are used as hosts. Being aware of possible toxic ef¬
fects is important, then, in choosing container materials.

52

Table 1. — Characteristics of materials for making rearing containers

Material

Cost

Durability

Weight

Paperboard .

Low

Low .

Light .

Meta!

High

High

. Heavy

Fiberglass .

High

High

. Light

Pressed

fiberboard.

Wood

Medium .

High

Medium

High

Medium

Medium

Thermoplastics:

Styrene .

Low

High

Light . .

Polyethylene

Low

High

Light . .

ABS .

Medium

High

. . . . Medium

Cellulose

nitrate.

Medium

High

Light . .

Polycarbonate

film.

Medium .

High

Light . .

Advantages Disadvantages

Rigid, strong

Rigid, strong, easily
cleaned.

Easy to make at
rearing facility.

Good for small
cups and boxes;
ideal for dispos¬
ables.

Resistant to most
solvents, oils, etc;
high density; low
maintenance.

Relatively rigid;
good solvent re¬
sistance; easily
fabricated.

Transparent; easily
fabricated; good
for plant-insect
cages.

Transparent; easily
fabricated.

Wears quickly; its
chaff is harmful;
unstable to all
liquids if un waxed.

Does not hold finishes
well; oxidizes; sharp
corners and edges
hazardous.

Prone to chipping and
cracking and may
irritate skin.

Short-term moisture
and oil resistance.

Possible toxic proper¬
ties to insects in
some woods, pre-
finished woods, and
various finishes.

Brittle.

Static charge attracts
dust.

None.

Inflammable.

Inflammable.

General structural

and physical considerations

Size and shape. — Size and shape must also be considered
in terms of structural design and container processing.
Size and shape can affect container strength. Size is im¬
portant for handling ease in terms of hand-load capacity.
Size may also affect storage efficiency, and shape certainly
does. Stacking, nesting, hanging, etc. need to be con¬
sidered, especially as ways to conserve expensive con¬
trolled environmental space during incubation and to
provide proper air circulation about the containers. Size
and shape must conform to processing equipment used in

the rearing program; close tolerances may be required by
automated equipment. Closures, seams, etc. must fit
properly to prevent airborne contamination of the con¬
tents, excess moisture loss, and escape of the insects.

Durability.— In the case of reusables, durability is im¬
portant in reducing replacement costs. Reusables must be
cleaned, so containers must resist wear due to repeated
cleaning, which may include both strong cleaning solutions
and steam heat. Reusable containers that can withstand
sterilization temperatures might be preferred if disease or
other contamination problems are anticipated. Containers
should also be resistant to appropriate diets; some are

53

Table 2.— Cost factors to consider when selecting an insect-rearing container

Container type

Cost element

Design effect

Disposable

Reusable

Purchasing .

Mass-produced containers less
expensive; prefabricated custom-
made more expensive. Initial de¬
sign important to prevent costly
die changes.

Low initial investment; continuous
costs accumulative; price
increases a problem.

Initial investment high but little ad¬
ditional costs. Initial design critical.

Shipping and storage .

Costs reduced with good nesting
and stacking qualities. Custom-
made containers may add to
shipping costs.

Costs high and continuous .

Shipping costs low; perhaps less stor-
age required.

Handling .

Properly designed containers pro-
mote handling efficiency and
reduce damage.

Few handling problems .

Much handling required.

Maintenance .

Good design promotes stability . . . .

No costs .

Some maintenance and replacement
costs.

Cleaning .

Well-designed containers more
cleanable.

No costs .

Labor, cleaning supplies, and cleaning
area costly.

Disposal .

Type of material important .

Disposal costs high; environmental
impact important; may require
compaction equipment; recy¬
cling possible.

Minimal disposal costs.

Availability .

Commonly used prefabricated
types more available.

Dependent on industry; inventory
critical.

Less dependent on industry.

dispensed hot and wet and, in some cases, have corrosive
properties. Disposables need not be durable.

Visibility and accessibility. — Container contents must be
easily seen. Frequent visual inspection is extremely
valuable to program maintenance and quality control.
Also, if host plants are used, the container must allow
adequate light transmission for the plant. Efficient
removal of insects from the containers requires good ac¬
cessibility. Both visibility and accessibility save valuable
time when insects are removed from containers at various
growth stages.

Availability.— Containers must be readily available for
programs with continuous production. (See appendix for a
list of some commercial sources for containers and ma¬
terials.) Close inventories and projected needs are
especially necessary for disposables. Mass-produced and
widely used containers, those primarily used for other
purposes, such as paper drink cups, are generally more
reliably available for future needs than customized con¬
tainers that are in less demand. Such prefabricated
availables also provide a developing program with a very
large choice of sizes, shapes, materials, and prices. Users
of handmade containers should consider continued avail¬
ability of materials.

Materials.— Materials for rearing containers are relevant
to almost every design criterion and every rearing re¬
quirement. Some of the more common materials available
for container fabrication are listed in table 1 with char¬
acteristics that may affect container design.

Cost. — The rearing container may be the most costly part
of the rearing program. Costs of disposables reflect direct
purchases, and costs of reusables reflect expensive clean¬
ing and the eventual replacement, which will also be ex¬
pensive. Size of the rearing program generally affects the
cost per container per insect, as larger quantities gen¬
erally produce lower unit costs. When programs become
large enough, equipment for container fabrication
becomes more economically realistic. But fabrication
machinery requires large initial cash outlays that are
sometimes difficult to justify, so proven container design
is especially important. The relationship of container de¬
sign to container requirements also affects container
costs as does the choice of disposable or reusable con¬
tainers (table 2). Rearing technique may dictate which
type of container to use; but, in other cases, careful con¬
sideration should be given to cost.

54

Closure requirements

Obviously, the container and the closure should be con¬
sidered as a unit, although there may be several combi¬
nations to choose from. So factors affecting container
choice also affect closure choice. Because the closure is
usually the part of the container that provides ventila¬
tion, and therefore is a device for controlling air and
moisture exchange, it is especially necessary to be aware
of the complex relationship between container and room
environment when choosing closure types and materials.
Simple materials, such as screen or cloth used to cover
vents, cardboard caps, or cotton plugs are often all that
is needed for adequate ventilation with proper room con¬
ditions. Different structure and compactness of materials
used for closures can affect the rate of evaporation from
the rearing container. Burton (1967) used unlined (un¬
waxed) cardboard caps when rearing fall armyworm,
Spodoptera frugiperda (J. E. Smith); Raulston and
Lingren (1972) used polypropylene cloth on multicell
units for rearing tobacco budworm, Heliothis uirescens
(Fabricius); Hartley et al. (1982) used rigid sheet
polypropylene on multicell units for rearing Heliothis zea
(Boddie); Kogan (1971) used layers of Cellucotton wad¬
ding to aid in moisture removal from rearing cells for
Mexican bean beetles, Epilachna variuestis Mulsant; and
Harrell et al. (1977) used Tyvek for rearing the boll
weevil, Anthonomus grandis grandis Boheman. A special
requirement for cap thickness is also necessary to pre¬
vent the corn earworm from chewing out. The use of
machines (Burton and Cox 1966, Davis 1982) for
manipulating closures adds still another dimension for re¬
quired tolerances.

The need for access into the rearing container is an im¬
portant factor in choosing closures. More durable
material and special handling techniques may be needed
when closures have to be reused as insects are observed
and fed during development. Ridgway et al. (1969)
described a method for access into Hexcel rearing con¬
tainers for repeated feedings. Cardboard lids on plastic
cups have proven useful when repeated feeding is
desirable and have also been adapted by Harrell et al
(1968) to mechanical harvest of the fall armyworm, and
similarly by Davis (1982) for the southwestern corn borer,
Diatraea grandiosella (Dyar). Some methods of harvest re¬
quire materials with special characteristics. For example,
Harrell et al. (1974) built a machine to remove Tyvek from
multicell rearing trays. Tyvek, a durable plastic product,
provided sufficient strength for efficient operation of the
machine, while paper cover material, weakened by
moisture from the diet, tended to pull apart under stress.

Close tolerances for closures are important. To prevent
closures in some types of containers (for example, lids for
plastic cups) from being pushed out or from falling off,
the container manufacturer’s specifications for closure

diameter requirements should be followed. Storage condi¬
tions may also affect the degree of fit for fibrous closures
by causing swelling under moist conditions and shrinkage
in drier environmental rooms where containers are kept
during insect development.

Technical information is seldom used but could be
valuable in selecting the proper materials for closures and
containers. For example, several give characteristics of
paper products, such as the porosity by resistance to
airflow and the water-vapor transmission rate. The Tech¬
nical Association of the Pulp and Paper Industry (TAPPI)
provides official test methods and respective TAPPI
reference numbers to member firms. These numbers pro¬
vide a standard that can be used when characteristics of
paper products are of interest.

The Future of Containerization

As costs continue to increase and demands for more and
higher quality insects grow, efficiency in rearing and in
controlling rearing variables and a need to standardize
programs will become more important. Use of the 1-oz
plastic cup has been so successful in most rearing pro¬
grams that a change here will probably not occur soon.
This slowness to change may also be true for other types
of disposables. But certain trends such as inflation and
scarcity of petroleum products will continue to have their
effect. These two factors may well bring about the
gradual end to the disposables we now so conveniently
use. When this end occurs, the need to develop replace¬
ment reusables will be even more important, and better
designs and more convenient reusable containers may be
the future of insect containerization.

References

Baumhover, A. H.; Cantelo, W. A.; Hobgo[o]d, J. M., Jr.;
Knott, C. M.; and Lam, J. J., Jr.

1977. An improved method for mass rearing the

tobacco hornworm. U.S. Agric. Res. Serv. [Rep.]
ARS-S-167, 13 pp.

Burton, R. L.

1967. Mass rearing the fall armyworm in the
laboratory. U.S. Agric. Res. Serv. [Rep.]
ARS-33-117, 12 pp.

Burton, R. L., and Cox, H. C.

1966. An automated packaging machine for
lepidopterous larvae. J. Econ. Entomol.

59: 907-909.

Chada, H. L.

1962. Toxicity of cellulose acetate and vinyl plastic
cages to barley plants and greenbugs. J. Econ.
Entomol. 55: 970-972.

Davis, F. M.

1982. Mechanically removing southwestern corn borer

55

pupae from plastic rearing cups. J. Econ. En-
tomol. 75: 393-395.

Harrell, E. A.; Hare, W. W.; and Burton, R. L.

1968. Collecting pupae of the fall army worm from
rearing containers. J. Econ. Entomol.

61: 873-876.

Harrell, E. A.; Perkins, W. D.; Sparks, A. N.; and Moore,
R. F.

1977. Mechanizing techniques for adult boll weevil
production. Trans. ASAE 20: 450-453.

Harrell, E. A.; Sparks, A. N.; and Perkins, W. D.

1974. Machine for collecting corn earworm pupae.

U.S. Agric. Res. Serv. [Rep.] ARS-S-43, 4 pp.

Harrell, E. A.; Sparks, A. N.; Perkins, W. D.; and Hare,
W. W.

1973. An insect diet filler for an inline form-fill-seal
machine. J. Econ. Entomol. 66: 1340-1341.

Hartley, G. E.; King, E. G.; Brewer, F. D.; and Gantt,

C. W.

1982. Rearing of the Heliothis sterile hybrid with a
multicellular container and pupal harvesting. J.
Econ. Entomol. 75: 7-10.

Kogan, M.

1971. Feeding and nutrition of insects associated with
soybeans: I. Growth and development of the
Mexican bean beetle, Epilachna varivestis, on
artificial media. Ann. Entomol. Soc. Am.

64: 1044-1050.

Leppla, N. C.; Carlyle, S. L.; Pons, W. S.; and Mitchell,

E. R.

1975. Use of a rodent rearing container for culturing
insects. J. Ga. Entomol. Soc. 10: 326-327.

Morrison, R. K.; House, V. S.; and Ridgway, R. L.

1975. Improved rearing unit for larvae of a common
green lacewing. J. Econ. Entomol. 68: 881-882.

Peterson, A.

1964. Entomological techniques. How to work with
insects. 435 pp. Edwards Brother, Inc., Ann
Arbor, Mich.

Raulston, J. R., and Lingren, P. D.

1969. A technique for rearing larvae of the bollworm
and tobacco budworm in large numbers. J.
Econ. Entomol. 62: 959-961.

1972. Methods for large-scale rearing of the tobacco
budworm. U.S. Agric. Res. Serv. Prod. Res.
Rep. 145, 10 pp.

Ridgway, R. L.; Morrison, R. K.; and Badgeley, M.

1969. Mass rearing a green lacewing. J. Econ. En¬
tomol. 63: 835-836.

Singh, P.

1977. Artificial diets for insects, mites, and spiders.
594 pp. IFI/Plenum Co., New York.

Smith, C. N. (ed.).

1966. Insect colonization and mass production. 618
pp. Academic Press, New York.

Appendix. — Some commercial sources for containers and materials

Containers
and materials

Use

Sources

Paper containers
for food and drink.

Insect holding .

Solo Cup Co., Urbana, Ill.; Sweetheart Cup Div.,
Owings Mills, Md.; Lily-Tulip Div., Toledo, Ohio;
Dixie Cup Div., Green Bay, Wis.

Plastic stock .

Forming film for
larval rearing.

Standard Packing Corp., Clifton, N.J.; B. F. Good¬
rich Co., Atlanta, Ga.; Foster Grant Corp., San¬
dusky, Ohio.

Tyvek and paper

Cover of rearing

Allegheny Label Inc., Cheswick, Pa.; Cellu-Craft,

stock plastics.

cells.

Inc., New Hyde Park, N.Y.; Tolas Corp., Phila¬
delphia, Pa.; Transilurap Inc., Cleveland, Ohio.

Porex

polypropylene sheet.

Cover of rearing
cells.

Porex Materials Corp., Fairburn, Ga.

Polypropylene
cloth fabric.

Cover of rearing
cells.

Chicopee Manuf. Co., Cornelia, Ga.

Polystyrene

cups.

Larval holding .

Unijax, Inc., Raleigh, N.C.; Premium Plastics, Chi-
cago. Ill.

Paper lids .

Cup cover .

Unijax, Inc., Raleigh, N.C.; Premium Plastics, Chi-
cago, Ill.; Standard Cup & Seal, Chamblee, Ga.

Polycarbonate
plastic film.

Cage construction . . .

Cadillac Plastic & Chemical Co., Oklahoma City,
Okla.

56

Section 3

Engineering for Insect Rearing

Tremendous progress has been made in the last few years
in the art of controlling insects. Most of this work with
insects has rightly been considered the province of ento¬
mologists. But engineers with the proper background and
training can provide important expertise for solving some
insect-related problems, and they are making significant
contributions.

Engineering involvement on many of the insect-related
problems has been slow to materialize. A few engineers
have been working for many years with entomologists on
specific problems of mutual interest, such as trapping
and insecticide application. Recently, other engineers
have begun working on rearing and modeling. There are
still many more engineering problems related to insects
than there are engineers working to solve them. So only a
few of the major problem areas will be covered in this
section— environmental control for insects and personnel,
control of respiratory hazards to humans, design of
insect-rearing facilities, insect-rearing automation,
materials-handling processes, quality-control procedures,
and systems analysis and modeling. Many other areas
have problems needing engineering attention— shipping;
harvesting insects; packaging; diet preparation; design of
rearing containers and covers; safety; building materials;
quality and intensity of light; photoperiod; thermoperiod;
work areas; rearing systems for parasites, predators, and
pathogens; and specific rearing systems for different
insects.

Adequate engineering involvement in all the problem
areas would require additional personnel. Those already
involved are spread too thin, and the probability is high
that the number will decrease. There are too many prob¬
lems for a reduced number of qualified personnel to handle.
A person’s academic training in engineering or entomol¬
ogy does not fully qualify him, nor does it guarantee suc¬
cess. This type of training is essential, but it must be
supplemented with experience and knowledge gained
from a hands-on operation.

Personnel already at work could probably be used more
effectively by the establishment of a central insect-
research center staffed with qualified engineers. These
engineers could be assigned to seek solutions to the most
pressing problems as they arise anywhere in the United
States. If a problem could not be solved at the center,
then the staff would go to the problem area, solve it, and
return for further assignment. Also, the staff could sup¬
ply a much-needed consulting service. In a few years,
such a center, properly staffed and supported, could prob¬
ably make tremendous progress toward solving some of
the problems, at much less cost than otherwise. A review
of recent accomplishments indicates that progress has
been rapid when engineers and entomologists cooperate,
as they complement each other well.

E. A. Harrell,

Research agricultural engineer, retired,
Agricultural Research Service

57

Controlled Environments
for Insects and Personnel
in Insect-Rearing Facilities

By Charles D. Owens1

Introduction

Environmental control involves regulation of the external
conditions that affect the growth, development, and be¬
havior of an organism. In a facility where insects are
reared, particularly in large numbers, the conditions that
must be regulated include temperature, humidity, noise,
and the movement and cleanliness of air. Experience has
shown that as the insect concentration increases in a pro¬
duction system so does the incidence of contamination,
disease, and related problems. Mass rearing often causes
allergy problems for workers; therefore they need pro¬
tection from excessive exposure to moth scales and urti-
cating or allergenic setae (Etkind 1976, Press et al. 1977).
And rearing under unsanitary conditions can result in in¬
sect death because of microbial contaminants or patho¬
gens, and workers are likely to become sensitive to these
allergens. So provision of clean air and maintenance of
sanitary conditions are especially important in the design
of a mass-rearing facility. In fact, all aspects of the en¬
vironment in a mass-rearing facility can be made to per¬
form satisfactorily with proper engineering design. This
paper surveys equipment and design factors important
to control of rearing environments for both insects and
personnel.

Air Cleaners

Air must be clean before it is brought into a rearing
room. And undesirable airborne particles and gases must
be removed from air that is recirculated. Equipment to
clean or filter air includes: air washers, viscous fiber or
dry filters, electrostatic precipitators, and cyclones. A
more detailed description of air cleaners and air perform¬
ance can be found in the handbooks of the American So¬
ciety of Heating, Refrigerating, and Air-Conditioning
Engineers (1972, 1973, 1975, 1978).

Air washers are dust-collecting devices that pass the air
through a spray of water to wash out the contaminants.
They are used to control temperature and humidity in en¬
vironmental rooms. But they are relatively inefficient in

‘Agricultural engineer, retired, Otis Methods Development
Center, Agricultural Research Service, U.S. Department of
Agriculture, Otis Air National Guard Base, Mass. 02542.

removing fine particles, so they are used primarily to
remove undesirable gases and wettable particles.

Viscous fibrous filters are flat panels of coarse fibers. The
fibers are coated with a viscous substance that acts as an
adhesive to the impinging particles. Some of these filters
can be cleaned and reused, and some are disposable.

Dry filters are usually made of fiber- or blanket-like ma¬
terial of varying thicknesses. The removal of airborne
particles depends on the closeness of the fibers. Indus¬
trial cloth bags are used as filters for air having a high
dust load with a particle size of 0.5 nm or larger. The effi¬
ciency of industrial filters varies and is rated according to
the specifications of the U.S. National Bureau of Stand¬
ards or standard dust-particle test. Air filters used in
ventilating systems are usually rectangular dry-fiber
units. Absolute or HEPA (high-efficiency particulate air)
filters are used for very low dust loads and have a high
efficiency for collecting particles down to 0.3 pm. They
are rated by a smoke test known as the DOP (dioctyl
phthalate) penetration test. The HEPA filters are used
for cleanrooms and laminar-flow hoods.

Dry centrifugal collectors called cyclones are the most
common type of collector for large particles. They
separate particles from the air by radial acceleration or
centrifugal force. Air entering the cyclone is transformed
into a vortex, causing centrifugal action to force the par¬
ticles against the wall. Ultimately, the particles travel to
the lower end of the cone where they are collected.
Cyclone air cleaners are generally used when particles are
present in large quantity and their size exceeds 50 pm
(for example, moth scales). Although some facilities use a
series of cyclone collectors to remove scales from the air,
a cyclone combined with a bag filter is more efficient.

Electronic air cleaners used in cleaning ventilated air are
designed as electrostatic precipitators. The air passes
through a high potential ionizer field that gives a charge
to the particles; these are then attracted to ground plates.
The particles can be removed from the collector plates by
washing. Since the charged particles form clumps on the
plates, regular cleaning is necessary.

Selection of an air filter depends on the amount and size
of contaminants in the air and requirements for air clean¬
liness. As more insects are reared, the need for clean air

58

Table 1. — Number of particles (>0.5 m) per 0.03 m3 of air outside
and inside the insect-rearing facility at Otis Methods Develop¬
ment Center

Location

Filter

Average

particle

count1

Outdoors .

None

50,000

Laboratory .

Holding room:

None

90,000

1 V2 minutes per air change

. 95% HEPA

200

3 minutes per air change . .

95% HEPA

500

Infesting room workroom .

95% HEPA

. . 1,500

'Reading taken with particle counter.

/

increases because of the increased occurrence of patho¬
gens and microbial contaminants. If the insects are
highly susceptible to airborne pathogens, the 99.9%
HEPA filters should be used with at least one air change
every 2 minutes. Generally, the 95% HEPA filter (hospi¬
tal grade) will maintain sufficiently clean conditions in
insect-rearing and other work areas (table 1). Also the
more efficient the filter, the greater the air pressure dif¬
ferential across the filter. So high-efficiency filters require
care in installation to prevent air bypass leaks.

Controlling Environment
for Insects and Personnel

Control of air, temperature, humidity, and other environ¬
mental factors is required in all areas of an insect-rearing
facility, but how much control and how it is accomplished
vary for different areas and insects. And, unless insects
are held for prolonged periods in workrooms, control of
some or all of these factors in these rooms can be de¬
signed for worker comfort.

Controlling air for personnel

Because dust generated from the body scales of adult
moths is a primary cause of human allergies, the air
should be free of contaminants in areas where workers
must handle insect stages that are potentially hazardous
to their health. So, during handling of insects, the air
should flow past the workers, over the work area, and
through a filter system before returning to the room. A
velocity of 30-40 m/min is required to capture and con¬
vey the scales and hairs. A velocity of 15 m/min or higher
will transport the smallest ( -Min) and lighter particles.

The easiest way to protect the worker is to use a work¬
table or clean-air station, such as the one developed for
handling larvae and pupae of the gypsy moth, Lymantria
dispar (Linnaeus), (fig. 1). Air is drawn past the worker,

TO VACUUM CLEANER

Figure 1.— Cross section of a clean-air work¬
table.

into slots in the center of the table, and through an
industrial-grade filter; it is then discharged back into the
room. The industrial filter has a glass-fiber pack for the
prefilter and an overall efficiency rating of 80%. The air
velocity, varying from 15 m/min near the workers to 107
m/min at the opening, causes the air current to flow
toward the center of the table. Because of the open de¬
sign of the system, large particles settle on the table and
are removed with a separate vacuum system. Two
vacuum outlets connected to one central vacuum cleaner
can remove particles such as webbing, cast skins, and
clumps of scales when both outlets are operating. For
small operations, commercial clean-air hoods can be used
if the air current is drawn into them; most horizontal
laminar-flow hoods (a kind commonly used in labora¬
tories) blow clean air out toward the operator and are not
suitable for small operations.

Disposal of live moths requires a relatively high air veloc¬
ity that can be provided by using a hood that tapers
toward a slot (fig. 2). The air velocity should exceed 30
m/min at the front and range from 305 to 457 m/min at
the exhaust. With this type of hood, the front part is
used to transfer or handle moths, and the area nearest
the discard slot conveys moths or other heavy particles
into the vacuum. This arrangement is used in the adult¬
handling room of the gypsy moth rearing facility at the

59

Figure 2.— Diagram of a high-velocity work
hood connected to a cyclone collector.

Otis Methods Development Center, Otis Air National
Guard Base, Mass., for housing mating cages, removing
excess moths, and handling egg masses. The high-
velocity outlet is connected to a cyclone that collects the
moths, while the lighter particles are removed by a bag
filter. Before the air returns to the room, it is passed
through a 95% HEPA filter (Bell 1981). Some rearing
facilities have moth cages placed over funnels that are
connected to a cyclone air cleaner. This arrangement also
extracts the scales that become dislodged from the moths
and reduces the number of airborne particles. In addition
to the clean-air work stations, each workroom should
have all air pass through a HEPA filter at least once
every 3-4 minutes.

Controlling air for insects

HEPA filters are necessary to reduce the incidence of air¬
borne bacterial, fungal, and viral pathogens, thereby
decreasing the probability of diet contamination and dis¬
ease. Also, increasing the number of air changes per unit
of time (for example, to 2-3 minutes per change) gen¬
erally decreases the number of particles. A slight positive
pressure should be maintained in larval-development
rooms so that contaminated air will flow out when the

doors are opened. Positive pressure is accomplished by
having a filtered fresh-air intake. Larval-development
rooms do not require much outside air, and the more
times air passes through a filter, the cleaner it should be.
Rooms that are normally contaminated can be operated
under pressure so air will flow into them (Shapiro 1980).
Negative pressure is accomplished by using an exhaust
fan to exhaust air through a HEPA filter. The air in
insect-rearing rooms must be controlled so that the tem¬
perature, humidity, and air velocity are fairly uniform.
This uniformity is difficult to accomplish.

Air distribution can be provided by standard heating and
ventilating arrangements in work, diet-preparation, and
insect-handling areas. But holding rooms require a sys¬
tem that produces more uniformity. Standard vertical
laminar-flow cleanrooms have the air blowing through a
perforated ceiling to a grated floor. This design is ex¬
pensive, and keeping the area below the grating clean is
difficult. A better arrangement, adapted from a design by
Klassen (1971), provides good air distribution by using a
wall plenum return and a perforated ceiling to allow air
into the room (fig. 3). Moving the air horizontally, as in a
clean-air, horizontal laminar-flow system (Harrell 1979b),
also provides superior air distribution. Air enters through

60

SUPPLY DUCT

'

^RETURN DUCT

J

LIGHTS'

J/k

A

PERFORATED CEILING

EXHAUST PORT

Sk

Figure 3.— Diagram showing airflow in a larvae
rearing room.

Figure 4.— Typical arrangement of air-
conditioning and filtering equipment in a lar¬
vae room. A. Air-conditioner bypass system.
B. Oversized duct system.

holes in one sidewall and exits through a similar arrange¬
ment on the opposite side. This design works better in
narrow rooms.

A 2-3 minute air-exchange rate requires more airflow
than is needed to maintain the temperature and humidi¬
ty. Therefore, to conserve on installation and operating
costs, the air-handling system may be designed to allow
50%-75% of the air to bypass the conditioning equip¬
ment (fig. 4). This bypass can be accomplished by making
the duct area larger than the heating and cooling coils or
by locating the conditioning equipment in a bypass par¬
allel to the main duct. The latter arrangement was used
effectively for the Otis Methods Development Center’s
gypsy moth rearing facility. With the bypass system, the
air-conditioning equipment was sized to the heating and
cooling load and not to the airflow rate. This arrange¬
ment was less expensive than a system without a by¬
pass would have been and provided better temperature
regulation.

Controlling temperature

and humidity

The longer the larval growth period, the more nearly
uniform the temperature has to be for developmental syn¬
chrony. In a large room, the temperature can be con¬
trolled by a good (±1° C or better) thermostat. But small
cabinets or walk-in chambers require thermostats with
sensing devices that respond more quickly to changing
environmental conditions. Since energy requirements are
high for large larval development rooms, it is more eco¬
nomical to have a ±2° C differential between the heating

and cooling cycles rather than the typical on/off control
system. A regular fluctuation (l°-2° C) in room temperature
does not hamper insect development. One way to compen¬
sate for differences in the room environment is to routinely
rotate the insects around the room so that all are exposed
to similar conditions. This rotation can be done with a con¬
tinuous conveyor system.

The temperature and humidity of the room are not neces¬
sarily the same as they are in the rearing container. But
the room environment affects the diet’s drying rate (Hare
et al. 1973). The amount of diet, number of insects per
container, the material the containers are constructed from,
and density of containers also affect humidity and air ex¬
change. Therefore, room conditions should be adjusted
for all these influences so that the desired environment
will be maintained in the container (Harrell 1979a).

The best way to add humidity to the environment is to
inject steam into the airstream in front of a recirculating
fan. Spray or atomizer devices often discharge excess
water into a room and become clogged with mineral de¬
posits. And the hot-air-furnace humidifier is not satis¬
factory in insect-rearing facilities because it requires a
large heat differential to operate, and that is not present
in a rearing room.

The cooling coils of air-conditioners can be used to de-
humidify some rooms. Low airflow and low coil tempera¬
ture are required to maintain a low dewpoint temperature.
So large refrigeration units with special compensating
devices are used to avoid frosting problems. If humidity

61

is to be held below a 10° C dewpoint, some type of ab¬
sorption dehumidifying system must be used. The most
common system uses silica gel or activated alumina as
the desiccant. Generally, the equipment incorporates a
regeneration system to reactivate the desiccant.

Controlling lighting

Work areas need nonglare overhead lighting; low-level
general illumination is adequate for holding rooms. Light
intensity experienced by the insects will vary consider¬
ably depending on types of containers and on the way the
containers are distributed in the rooms. Generally, insects
respond to very low light intensity, so just enough is pro¬
vided to entrain their rhythms. Supplementary lighting
of 50-100 fc is required for tasks such as handling newly
hatched larvae or counting eggs. This lighting is similar
to the requirements of desk and office work. The place¬
ment, shielding, and type of light, and also the texture of
work surfaces, can affect the amount of eyestrain caused
by glare or improper illumination.

Controlling noise

In rearing and handling rooms, noise is produced by me¬
chanical devices and high-velocity airblowers. An airflow
system has two sources of noise— the low frequencies
from the fan and the high frequencies produced by air
flowing through the outlet or inlet grills. Centrifugal fans
with airfoil blades produce less noise than other types.
Sound transmission can be reduced by isolating the fan
and duct with a flexible coupling. HEPA filters also re¬
duce noise levels. Designing ducts and air outlets so that
air velocity is less than 200 m/min reduces the high-
frequency sound generated by grills. Increasing the mass
and stiffness of hoods will dampen their vibrations.
Sometimes, as with the high-velocity exhaust hood used
in moth rooms, the noise level cannot be reduced to de¬
sired levels. In such cases, either the length of exposure
should be reduced or earplugs should be used. If the
sound is annoying, but not hazardous, structural changes
can be used to shift it to a less objectionable frequency,
or a more pleasant sound can be added to mask the orig¬
inal. Noise can also be reduced by decreasing the energy
that drives the vibrating components, changing the coup¬
ling between the energy source and the acoustical radiating
system, altering the structure that is radiating the sound,
and using attenuators.

Other Design Considerations

Operating costs, as well as initial costs, must be con¬
sidered in designing a facility. For example, more energy
is required to move air through a more efficient filter. If
industrial-grade prefilters are used, HEPA filters will last
longer. Increasing the number of air changes per unit of

time requires more energy and increases the noise level.
Rooms should be well insulated (R-ll or better) to reduce
the amount of air-conditioning needed and to moderate
temperature changes during electrical power outages. The
walls, floors, and ceiling should be finished with materials
that facilitate cleaning and sanitizing (smooth and wash¬
able surfaces such as concrete floors and painted walls
and ceilings). A central vacuum-cleaning system or a
portable cleaner with an absolute filter should be used to
remove dirt from the floor. Workers should take precau¬
tions to prevent transporting contaminants into the facil¬
ity. And the work should be performed in a sequence
ranging from clean to dirty.

Recommendations

To best control the environment of insect-rearing facili¬
ties: Provide individual air-conditioning units and con¬
trols for all rooms; have a backup holding room, so insects
can be moved in case of mechanical failure or contamina¬
tion; insulate rooms to reduce temperature change during
an electrical power failure; properly install HEPA filters
in all rooms; use industrial-grade filters as prefilters to
the conditioning equipment; recirculate the air every 2-3
minutes; maintain low air velocities to reduce noise; pro¬
vide uniform airflow in the rearing rooms; distribute rear¬
ing containers so airflow affects them equally; use steam
humidifiers; install exhaust hoods equipped with clean-air
filters; and monitor air cleanliness with a particle counter.

References

American Society of Heating, Refrigeration, and Air-
Conditioning Engineers.

1972. ASHRAE handbook and product directory,

1972. Fundamentals. The Society, New York.

1973. ASHRAE handbook and product directory,

1973. Systems. The Society, New York.

1975. ASHRAE handbook and product directory,
1975. Equipment. The Society, New York.

1978. ASHRAE handbook and product directory,
1978. Applications. The Society, New York.
Bell, R. A.; Owens, C. D.; Shapiro, M.; and Tardif,

J. G. R.

1981. Development of mass rearing technology. In
C. C. Doane and M. L. McManus (eds.), The
Gypsy Moth: Research Toward Integrated Pest
Management, pp. 154-161. U.S. Dep. Agric.
Tech. Bull. 1584.

Etkind, P. H.

1976. The allergenic capacities of the gypsy moth
Lymantria dispar L. Ph.D. thesis, Yale Uni¬
versity, New Haven.

Hare, W. W.; Harrell, E. A.; and Perkins, W. D.

1973. Moisture removed from rearing cells during
incubation of com earworm on an artificial

62

diet. J. Econ. Entomol. 66: 283-285.

Harrell, E. A.; Perkins, W. D.; and Mullinix, B. G.

1979a. Effects of temperature, relative humidity, and
air velocities on development of Heliothis zea.
Ann. Entomol. Soc. Am. 72: 222-223.

1979b. Environmental rooms for insect rearing.
Trans. ASAE 22: 922-925.

Klassen, W., and Gentz, G.

1971. Temperature-constant and temperature gra¬
dient-face insectary design and operation. J.
Econ. Entomol. 64: 1334-1366.

Press, E.; Googins, J. A.; Poareo, H.; Jones, K.; Perlman,
F.; and Everett, J. R.

1977. Health hazards to timber and forestry work¬
ers from the Douglas-fir tussock moth. Arch.
Environ. Health 33: 206-210.

Shapiro, M.; Bell, R. A.; and Owens, C. D.

1981. In vivo mass production of the gypsy moth
nucleopolyhedrosis virus. In C. C. Doane and
M. L. McManus (eds.), The Gypsy Moth: Re¬
search Toward Integrated Pest Management,
pp. 633-655. U.S. Dep. Agric. Tech. Bull.
1584.

63

Controlling Respiratory
Hazards in Insectaries

By Wayne W. Wolf1

Introduction

Present laws in this country mandate that the employer
must provide a safe working place for employees. The
first responsibility lies with the employer in recognizing
hazards and providing appropriate equipment and work
procedures. Further responsibility lies with supervisors
and workers in providing instruction, maintaining equip¬
ment, and performing work in a safe, prescribed manner.

The objectives of programs for respiratory protection are
to save workers from suffering ill effects, prevent lost
time, and prevent permanent injury. These objectives
may be accomplished by recognizing that respiratory
diseases can be prevented like other diseases. For a
disease to occur, there must be a susceptible host, a
means of transmitting the disease, and a successful trans¬
fer. Preventing transmission will be the primary defense
against respiratory diseases in an insectary.

Respiratory Hazards
in Insectaries

The respiratory hazards often present in insectaries are
particulate matter and gases that adversely affect the
human body. These hazards may originate from various
insect stages, insect waste products (such as frass), mold
spores, materials used in insect diets, or cleaning and
sanitizing chemicals. These substances have various
modes of action and may produce skin injuries, allergic
reactions, asphyxiation, or damaged internal organs.
Allergic reactions may involve the skin, eyes, and respira¬
tory tract, and prolonged exposures may produce
irreversible pulmonary damage. Allergic reactions may
develop after repeated exposure over a period of years,
and some individuals may become very sensitive to low
concentrations. Routine work such as counting, sexing,
dissecting, marking, weighing insects, and handling and
cleaning insect cages can increase the air contamination.
Moth scales are a primary source of contamination.

Particles greater than 214-3 ^m in diameter are mostly
deposited in the upper respiratory system, while particles
about 1 (im in size are deposited most efficiently in the

‘Agricultural engineer, Southern Grain Insects Research
Laboratory, Agricultural Research Service, U.S. Department of
Agriculture, Tifton, Ga. 31793.

alveolar cells of the lungs. Conditions such as rhinitis,
laryngitis, and bronchitis can develop in the upper res¬
piratory system. In the lower system, conditions such as
emphysema, pleurisy, pneumonia, and pneumoconiosis
can develop (Olishifski 1978).

Airborne concentrations of substances that workers may
be safely exposed to day after day and the acceptable
conditions of exposure are called threshold-limit values
(TLV). The TLV guides that have become the most widely
accepted are those issued by the American Conference of
Governmental Industrial Hygienists; they are reprinted
in National Safety News (1979a). The TLV’s listed in¬
clude various chemical and physical hazards that may be
encountered in an insectary. Hazards such as insect scales
or allergy-inducing materials are not covered.

Detecting Contaminants
in Insectaries

Assessing the severity of hazards or effectiveness of con¬
trol equipment requires some method of detection or meas¬
urement. Measurement of many gases can be accomplished
quickly and inexpensively with small “grab” samplers.
Such a sampler may consist of a small glass tube and a
pump that forces a measured quantity of air through the
tube. The tube contains granular chemicals that change
color to indicate the concentration of a specific gas such
as chlorine or formaldehyde. Sources of sampling equip-

Table 1.— Particle size distributions in various locations at the
Western Cotton Research Laboratory1

Particle

size

w

Distribution (particles/m3)

Air in pink

bollworm

cages2

Outdoor

air3

Filtered air
in moth room4

0.5-1

6,600,000

1,400,000

74,000

1-2

2,600,000

560,000

16,600

2-5

1,600,000

233,000

14,800

5-10

350,000

7,800

5,300

>10

2,900,000

3,500

9,200

‘Measured with Climet Model 250 particle counter.

2 Average of 8 cages, one run per cage.

“Average of 6 runs.

4 Average of 12 runs (3 runs every 4 hours) after room reached
equilibrium. Filter system shown in figure 4.

64

30 D

i

ment may be obtained from offices of the U.S. Occupa¬
tional Safety and Health Administration (OSHA) or from
various safety-product buyer’s guides such as National
Safety News (1979b).

Distributions of concentrations and sizes of airborne par¬
ticles may also be measured (Cadle 1975). The measure¬
ments may be made with filtration, sedimentation, centri¬
fugal, impaction, electrostatic-precipitation, or optical
techniques. For high concentrations of large particles,
filtration through fine mesh filters provides quick, inex¬
pensive sampling. The filter may be weighed before and
after sampling to provide concentrations expressed as
weight per unit of volume of air. This method may be
useful around diet-preparation areas where concentrations
are high. In rearing areas, the nature and concentrations
of particles such as insect scales require equipment ca¬
pable of measuring very small particles at relatively low
concentrations. Particle counters using light-scattering
principles can count individual particles as small as 0.3
fim, and some of these instruments can indicate numbers
of particles in various size ranges (Cadle 1975).

This type of equipment was used to sample particles at
the U.S. Agricultural Research Service’s Western Cotton
Research Laboratory, Phoenix, Ariz. (table 1). The egg-
laying room contained moths of tobacco budworm, Heli-
othis virescens (Fabricius); pink bollworm, Pectinophora
gossypiella (Saunders); beet armyworm, Spodoptera exi-
gua (Hiibner); and cabbage looper, Tricoplusia ni
(Hiibner). Air was sampled from inside pink bollworm
cages because they were the smallest and most numerous
moths in the room. Air was also sampled near the middle
of the egg-laying room and outside the laboratory.

These measurements indicated that very small particles
were present in the moth cages and that dry, 95%-efficiency
filters could remove them. The particle concentration at
the outlet of the filter in the moth room was below the in¬
strument’s detection threshold. Many particles in the
moth room probably entered from outdoors; counts in the
moth room varied with outdoor particle counts during the
day.

Reducing Human Susceptibility
to Insectary Contaminants

A person’s susceptibility to an allergen or an irritant is
difficult to determine. Also, different people exposed to
similar environments respond differently. This variation
may be due to variations in the rate of clearance from the
lungs, the effects of cigarette smoking, existing pulmo¬
nary diseases, and genetic factors. Preemployment
screening may keep susceptible people from being ex¬
posed, and annual allergy tests and tests of pulmonary
function may be used to monitor workers to detect de-

Nr

Figure 1.— Distances from blowing and ex¬
hausting nozzles where air velocity is less than
10% of velocity at nozzle opening. D=nozzle
diameter.

veloping symptoms or susceptibility. Desensitization
treatments benefit some individuals. These treatments
and tests must be performed by qualified medical person¬
nel. Low contamination levels reduce the probability of
an individual acquiring a sensitivity to a specific sub¬
stance. Engineering controls and proper work procedures
can lower contamination levels.

Engineering Controls
of Insectary Contamination

Engineering controls include mechanical equipment such
as filters, ducts, containers, blowers, motors, and hoods
to prevent contamination from reaching a worker. The
selection, design, and use of such equipment are the most
important steps in controlling respiratory hazards. Since
particles that cause respiratory problems settle slowly,
natural settling is not a solution. For example, l-am sized
sawdust particles need 75 hours to settle 1.5 m (McDer¬
mott 1977). Small particles remain suspended in the air
almost like gas molecules.

Applying controls near the source of contamination gen¬
erally uses less energy, costs less, and works best. The
use of air jets to move particles away from a work station
usually worsens a problem because turbulence associated
with the jet spreads the contaminants downstream and
may even entrain more particles. Exhausting air from
around the source removes contaminants so they may be
filtered from the airstream or safely discharged. The air
velocity should be at least 0.5 m/s at the point where con¬
taminants are picked up or entrained in the exhaust air-
stream. Unfortunately, air velocity is less than 10% of

65

Figure 2.— Insect cages mounted on exhaust
ducts to control moth scales. Air passes through
cages and is filtered by cyclone and dry filters
shown in figure 4.

nozzle velocity at a distance of 1 nozzle diameter from an
opening (fig. 1). This effect mandates that exhaust open¬
ings be located close to the contaminant source and also
makes overhead canopy hoods ineffective in collecting
particles from workbenches.

Many insects have been collected with oral aspirators
that inject contaminants directly into a worker’s lungs.
Oral aspiration should not be attempted without a filter
capable of stopping 99% of particles 0.3 gm in diameter.

Enclosures

Enclosures surround the source of contaminants to pre¬
vent them from mixing with room air. There may be
provisions for filtering air in the enclosure, filtering recir¬
culated room air, or exhausting dirty air outdoors. Enclo¬
sures include reach-in cabinets, enclosed shelves, and
special containers. Enclosures offer good protection as long
as the access openings are closed. When they are open, tur¬
bulence from work procedures, room air-conditioners, or
the transfer of material into or out of the enclosure can
mix contaminated air with room air.

Insect cages do not provide protection unless their walls
retain small particles or unless air is exhausted from the
cage. Cages can be made with material permeable to
moisture and vapor but able to block small particles (ma¬
terial such as fine knit cloth or perforated plastic). Ex¬
hausting air from screened cages removes particles before
they mix with room air, does not restrict access to cages,
and removes particles disturbed during feeding and egg
collecting (fig. 2). The air velocity through the screened

Figure 3.— Cabinets for separating egg-laying
cages of different strains of insects. Each
cabinet has air circulation from right to left
across each shelf and has 35%-efficiency filters
to collect scales.

walls should be 0.5 m/s or greater so that insect movement
inside the cage does not mix particles from the cage with
room air. The cages can rest on ducts with an opening to
each cage. Room air flows through the cage, through the
ducts, and is filtered before being discharged into the
room. Duct sizes should change at each branching point
to maintain adequate airflow between cages. Since chang¬
ing the duct size at each cage is not practical, the duct
for each row of cages should be large enough so the end
cage gets adequate ventilation. When several strains of
one species need to be separated, egg-laying cages may
have to be placed in separate cabinets (fig. 3).

Receiving hoods

Receiving hoods similar to those used above household
stoves are generally not satisfactory for dust control be¬
cause the air velocity at the work level is too low. These
hoods are used when the work process creates enough
heat that particles are carried into the hood as the hot air
rises.

66

Capturing nozzles

Capturing nozzles normally consist of a duct with a fared
opening. The fared opening (nozzle) is placed near the
contamination source, and the exhaust air carries par¬
ticles into the duct. Single nozzles are not satisfactory for
most insectary work stations because the air velocity
decreases so rapidly with distance from the nozzle. Add¬
ing more nozzles increases the capture zone but still may
not provide protection if multiple insect cages or contam¬
ination sources are at the work station or are being trans¬
ferred to the station.

Slotted ducts and perforation surfaces are also classed as
capturing nozzles. Slotted ducts are normally located
near the rear of the work station. Perforated surfaces on
the rear of the work station or a perforated countertop
provides the best airflow because clean air passes the
worker’s breathing zone, picks up particles in the work
zone, and is exhausted.

More working space is generally available when perfo¬
rated counters are used, because the exhaust ducts are
below the table. Adding partitions to sides and tops of
work stations creates a tunnel effect and improves the
airflow near the worker’s breathing zone. Supplying
laminar-flow clean air above a work station and exhaus¬
ting the air through a perforated counter provides the
best protection.

An approximate method for determining quantities of air
that must be exhausted to produce a desired capture
velocity is presented in American Society of Heating, Re¬
frigeration, and Air-Conditioning Engineers (1976, pp.
22.1-22.12). Air-duct design, loss coefficients, and recom¬
mended practices are described by Stomper (1979). Air¬
flow should be verified periodically with tracers such as
smoke and air-velocity meters.

Filters

If dirty air cannot be exhausted outdoors, then it must
be filtered before being recirculated in a room. The
amount of air, dust-loading capacity, type of contaminant
and smallest size of particle to be removed determine the
type of filter needed. Dust collectors are used when the
dust concentration is high and dust particles large, as in
a large moth-holding room. They can collect large quan¬
tities of dust that would quickly load more efficient
filters. Cyclone separators are the least costly dust collec¬
tors for medium-to-coarse granular dusts. They rely on
centrifugal force to separate dust particles from the air-
stream, are relatively small, and are inexpensive to main¬
tain. High air velocities in the cyclone chamber require a
high pressure drop across the collector; so operating costs
and noise levels tend to be high. Few small particles are

Figure 4.— Cyclone separator on right with
35%- and 95%-efficiency dry filters; charcoal
filter on left. Used in moth room to filter
scales collected from moth cages shown in
figure 2.

removed by cyclone separators; therefore they are almost
exclusively used upstream of more efficient filters
(Stomper 1979). Cyclones in series use large amounts of
energy because of the large pressure drop across each
unit. Since a single cyclone removes the bulk of material,
the remaining small particles can be removed with dry
filters (fig. 4).

Fabric dust collectors use cloth tubes or envelopes. Par¬
ticles larger than the fabric interstices are deposited by
simple sieving action. A mat of dust forms on the fabric
surface and improves the filtering efficiency. The collec¬
tors have built-in features to periodically remove accu-

67

mulated dust from the cloth surfaces. Common methods
of dust removal include mechanical shaking, reverse-air
collapse, and pulse jet. Airflow is stopped during clean¬
ing. Dust is removed from the cloth and then removed
from a drawer or hopper below the filter. These filters re¬
quire less energy and are more efficient than cyclones,
but initial expense and maintenance is greater. Filter
cloth can be selected to remove 99% of particles 1.0 /xm
in diameter.

Dry filters are the broadest category of air filters for a
variety of designs, sizes, and shapes. The most common
filter medium is glass fiber because of its low cost and
because the fiber diameter can be controlled during
manufacture. Generally, the finer the diameter, the higher
will be the air-cleaning efficiency. The fiberglass filters
used in home furnaces are not efficient enough to remove
most respiratory hazards. Hospital grade (95% efficiency)
or industrial grade (99.97% efficiency) HE PA (high-
efficiency particulate air) filters are recommended to ade¬
quately eliminate allergy, asthma, and pulmonary hazards
(Zeterberg 1973). (Stomper 1979 discusses filter types and
includes a guide to filter selection.)

Electronic filters of the type commonly sold for air-
conditioning systems tend to lose efficiency as they re¬
move dirt; they must be kept clean. They also generate
ozone. They have a higher initial cost and are not as effi¬
cient over long periods of time as less expensive dry
filters. Electronic filters maintain their initial low airflow
resistance, even when loaded with dirt.

Activated charcoal filters remove many gaseous contam¬
inants. They act as a catalyst to remove ozone and can
remove most odors associated with insectaries. Charcoal
filters are normally placed downstream of the last filter in
the system. The types of gases removed and life expec¬
tancies of filters can be obtained from manufacturers’
specifications.

Filter testing and rating is standardized industry wide
with the American Society of Heating, Refrigeration, and
Air-Conditioning Engineers test standard 52-76 (Stomper
1979). This standard describes tests for atmospheric dust
spots and arrestance. But the tests do not indicate the
removal efficiency for various sized particles. Some manu¬
facturers provide information on filters that relates filter
efficiency to particle sizes removed. HE PA filters are
tested with a homogeneous fog of dioctyl phthalate
(DOP), and particle concentrations upstream and down¬
stream are determined to a high degree of accuracy with
optical methods (Stomper 1979).

Respirators

to Prevent Contamination
of Insectary Personnel

OSHA mandates that engineering solutions be tried be¬
fore respiratory equipment is used. But some work pro¬
cedures, such as transferring insect cages to cleaning
stations, may not have practical engineering solutions.
Temporary jobs may involve transient exposure; and, if
engineering solutions such as plastic bags, water baths,
or special transfer carts are not possible for such jobs,
then respirators may be necessary.

Respirators may be obtained for protection against dusts
or gases. They should not be used in atmospheres that
would pose an immediate health threat to a worker with
no respirator. Only respirators that are certified effective
for the type of contaminant encountered should be se¬
lected, and they must cover the mouth and nose. If the
hazard is absorbed through or irritates the skin, then the
skin should also be covered. The respirator should fit the
individual user and not interfere with the work. Cloth or
paper dust respirators are generally more comfortable to
wear but provide less protection than cartridge res¬
pirators. Respirators are available that supply clean air to
the worker via a hose from a remote supply or from a
self-contained demand air supply.

An acceptable respiratory program should include the
following elements;

1. Written standard operating procedures should be
available for the selection and use of respirators.

2. Users should be instructed on proper use and limita¬
tions of respirators.

3. Certified respirators should be selected for the hazards
involved.

4. The respirator must be properly fitted.

5. Respirators should be cleaned and disinfected as often
as necessary to insure protection of the wearer.

6. Respirators must be used at all times when protec¬
tion is required.

7. Respirators should be stored in a convenient, clean,
and sanitary location.

8. Respirators used routinely should be inspected during
cleaning.

9. Defective respirators should be replaced or repaired
by experienced personnel before use.

10. There should be regular inspections and evaluations
to determine the Continued effectiveness of the res¬
piratory protection program.

Cloth facelets, beards, glasses, or goggles that interfere
with the sealing edges of the facepiece or valve action
preclude the wearing of a respirator. Contact lenses should
not be used by someone wearing a full facepiece or hood

68

respirator because the wearer would have to expose him¬
self to recover or adjust them (Day 1979).

Conclusion

The unique nature of each insectary requires special haz¬
ard assessments and appropriate controls. Some insects
tolerate various contaminants, so equipment can be de¬
signed to operate only when workers are present. Other
insects require HE PA filters and equipment to protect
them from bacterial, mold, and viral contamination. Such
equipment provides some protection from respiratory
hazard for insectary workers and may simplify the design
of respiratory-protection equipment.

References

American Society of Heating, Refrigeration, and Air-
Conditioning Engineers.

1976. ASHRAE handbook and product directory,
1976. Systems. 1,120 pp. The Society, New
York.

Cadle, R. C.

1975. The measurement of airborne particles. 342
pp. John Wiley & Sons, New York.

Day, R. O.

1979. Basic elements of a respirator program. Natl.
Saf. News 119(2): 44-45.

McDermott, H. J.

1977. Handbook of ventilation for contaminant con¬
trol (including OSHA requirements). 368 pp.
Ann Arbor Science, Ann Arbor, Mich.

National Safety News.

1979a. Threshold limit values for chemical sub¬
stances in workroom air. Natl. Saf. News
119(1): 66-73.

1979b. Product index. Natl. Saf. News 119(3): 138-
258.

Olishifski, J. B.

1978. Lung function and respiratory protection.
Natl. Saf. News 118(1): 47-56.

Stomper, E.

1979. Handbook of air-conditioning, heating, and
ventilating. 3d ed., 1420 pp. Industrial Press,
New York. (See especially section 5.)

Zeterberg, J. M.

1973. A review of respiratory virology and the spread
of virulent and possibly antigenic viruses via
air-conditioning systems. Ann. Allergy 31:
291-299.

69

General Requirements

for Facilities That Mass-Rear Insects

By Jack G. Griffin1

Introduction

A thorough knowledge of the unique combination of pro¬
cedures, equipment, space, and environment requirements
for rearing any particular insect is necessary for adequate
planning of the rearing facility. Therefore, describing an
ideal facility for rearing all insects is impossible. Most
structures currently used for insect rearing were designed
originally for other purposes (see, for example, several ar¬
ticles in Leppla and Ashley 1978) and modified later to
house small laboratory colonies. So these facilities vary
widely in type, floor plan, and construction method (see,
for example, several articles in Leppla and Ashley 1978).
And many facilities for large-scale rearing were planned
and constructed with small-scale rearing systems as
models and with general knowledge of the structural per¬
formance of similar buildings. Many of these facilities
were modified almost immediately after becoming opera¬
tional because some drawbacks that can be tolerated for
rearing small colonies become serious in mass rearing, es¬
pecially where contamination by micro-organisms is a
problem. Unfortunately, not much research has been done
on how facilities affect insect production. Despite these
handicaps, we know enough about the general needs and
problems of mass rearing insects to be able to describe
certain common requirements for insect-rearing facilities.
These include general space requirements, structure
types, construction materials and methods, and environ¬
mental control and equipment.

Space Requirements
for Rearing Facilities

Space requirements are not the same for all insects, but
most need places for storage, diet preparation, egg pro¬
duction, egg or larval implantation, insect development,
and adult emergence. For example, with the knowledge
and experience gained in mass rearing boll weevils, An-
thonomus grandis grandis Boheman, at Mississippi State
University’s Robert T. Gast Boll Weevil Rearing Facility
at Mississippi State, Miss., I recommend that a facility
for mass rearing this insect have 14 separate areas, each
consisting of one or more rooms. This number is needed

'Agricultural engineer, retired, Boll Weevil Research
Laboratory, Agricultural Research Service, U.S. Department of
Agriculture, Box 5367, Mississippi State, Miss. 39762.

to provide proper environments, sanitation and disease
control, photoperiod, and security. Unfortunately, stand¬
ards are not available for determining the space needed in
insect-rearing facilities. The size of each area varies with
use, insect to be reared, and production capacity. And,
adequate room must be provided for equipment, work
and storage space, and traffic lanes.

Structures

for Rearing Facilities

Structures for rearing facilities must meet the building
code of the location, provide the sanitary conditions and
environments needed for the rearing operations, provide
safety for workers, have adequate space, and be designed
to permit efficient arrangement of each operation. Some
types of buildings are better adapted and more economi¬
cal than others for rearing insects. These structures may
have metal or wood framework enclosed with various
materials, or walls of cinder block and other masonry.

The roof may be sloped or flat to accommodate air-
conditioning and associated handling equipment. (Placing
the air-conditioning equipment on the roof saves space,
and the roof might be the most efficient location.)
Modified mobile home units are also being used (J. Rober¬
son, personal communication, and see, for example, Gantt
et al. 1978), but these are not suitable for large, per¬
manent facilities.

Considerations in planning rearing-facility structures in¬
clude operation sequence, sanitation and disease control,
production operations, workflow, and security require¬
ments. These factors vary in importance with different in¬
sects. Sanitation, for example, is critical in boll weevil
rearing and is given priority. Control of pathogens and
other micro-organisms is probably one of the most impor¬
tant factors in producing large quantities of high-quality
insects. Therefore, areas where contamination control is
critical (rooms for handling sterile diet and rooms for in¬
sect development) should be separated from areas need¬
ing less isolation (egg-production colony, emergence room,
and areas where spent diet or adults are handled). Also,
some areas must be divided into separate rooms to fur¬
ther isolate the more critical contamination sources.
Because of high levels of contaminants in some diet in¬
gredients and their spread by air currents during proces¬
sing, diet-preparation areas should be separated. Airlock
vestibules reduce the amount of air that enters clean-
rooms. Places for personnel to shower and dress may also
be needed.

70

The functions and sequences of each operation must be
considered in arranging areas in a facility. For instance,
diet sterilization and larval or adult diet preparation and
dispensing should be located near each other. Areas
should also be arranged to provide the most efficient flow
patterns for materials and personnel. Good flow patterns
reduce backtracking, save time, prevent traffic conges¬
tion, and prevent interference between locations. A cen¬
tral corridor adjoining all production areas and extending
to the outside, if properly planned and arranged, can pro¬
vide good traffic and workflow patterns. Or, all the pro¬
duction areas can be joined by a covered outside walkway.
This approach is used at the R. T. Gast Rearing Facility.
But items moved from one area to another would be ex¬
posed to outside contaminants, and insects would be
more likely to escape. And the amount of travel between
areas might be greater than with a central corridor. A
combination of these two arrangements may be used to
best advantage.

Construction Materials
and Methods

Construction materials must provide permanency, clean-
ability, serviceability, economy, and safety. Usually,
however, no one material rates high for all these criteria.
For example, concrete flooring is the first choice for a
permanent facility. If smoothly finished, it is slippery
when wet; otherwise it is hard to clean and sanitize. It
also produces dust particles when dry and can be dam¬
aged by some acids. So concrete finished rough or with
an antiskid material applied to the surface can be used
for floors that are wet much of the time and where sani¬
tation is not a problem. Dressing and restroom floors can
be plain concrete with a filler applied to the surface to
reduce dust, or with a resilient floor cover added for
better appearance. Shower-, equipment-, and maintenance-
room floors that receive much wear, are frequently
washed, require a surface that is not slippery when wet,
and must provide a high degree of sanitation, should
have a quarry-tile cover over a concrete base. A chemical-
resistant grout should be used, and tile may be placed
whole or broken to reduce its cost. These floors should
also contain waterproof pans and drain outlets.

Materials are available for making a monolithic, seamless
floor topping that can be used on concrete. These have not
been tested widely in insect-rearing facilities; but one, an
acrylic resin mixed with dry components and troweled on a
concrete base in two layers, has delaminated from some
floors at the R. T. Gast Rearing Facility. Another of these
materials, applied on some floors at the U.S. Agricultural
Research Service’s Southern Field Crop Insect Manage¬
ment Laboratory, Stoneville, Miss., as a resinous mixture
without any dry components, has also peeled from the
base (C. W. Gantt, personal communication).

Some floors must be washed and rinsed with water and
sanitizing solution to keep them clean. This operation is
easier if floors slope to drains. P-traps should be used to
prevent sewage gas from entering the rooms, and trap
boxes are necessary to stop particles from clogging the
lines. Floors should be insulated to prevent surface con¬
densation in rooms that are maintained at warm temper¬
atures and high relative humidities during winter. Cold
floors also contribute to undesirable stratification of air
temperature.

Cinder blocks have been satisfactory as interior walls in
the R. T. Gast Rearing Facility. Originally, their surfaces
were rough and porous, but three coats of epoxy enamel
paint gave a smooth and durable finish. These block
walls withstand the occasional physical abuse that in¬
evitably occurs where equipment is moved. Plywood or
Masonite sheets with a plastic layer laminated on one
side also provide a surface that is tough and easy to
clean. They are used in food-processing areas but are
suitable elsewhere. Gypsum board or plaster painted with
epoxy enamel has served satisfactorily in areas where the
walls are not subjected to excessive moisture or wear.
Surfaces of plaster and gypsum board should be finished
smoothly for easy cleaning and sanitizing. Walls for
shower stalls and maintenance rooms usually have ceram¬
ic tile on a masonry base. Preferably, restroom walls
should have 1.2-m-high ceramic tile wainscots, or the en¬
tire wall may be covered with tile, tempered Masonite, or
exterior plywood boards with a layer of plastic laminated
to their interior surfaces. Metal walls are suitable in
storage rooms unless they are exposed to corrosive ma¬
terials. Stainless steel can be used in some areas, but it is
not recommended. Joints between the floor, walls, and
ceiling must be without crevices, so sometimes they are
joined by a masonry base. The base should be recessed
flush with the wall surface to avoid formation of a ledge.
Metal beads should not be used because they will cor¬
rode. Holes for utility lines that enter the wall cavity
should be sealed to prevent entrance of dust and micro¬
organisms. Walls should be a minimum of 2.4 m high,
and their exterior sides should be insulated to reduce
heat transfer, room-temperature stratification, and con¬
densation.

Gypsum board or plaster coated with epoxy paint will
usually suffice for ceilings in areas not subjected to high
moisture or overhead leaks. Otherwise, exterior-grade ply¬
wood or tempered Masonite with a laminated plastic
covering should be used. A good, tight fit, or a specially
fabricated metal connector strip, is needed with this type
of construction. Also, any openings for light fixtures or
utility lines should be sealed. Ceilings should be insulated
to facilitate temperature control in heated and cooled
rooms.

71

Windows are usually used for light and esthetics rather
than for ventilation. They should be kept to a minimum
and be nonopening, stationary panels with frames made
of heavy-duty metal. Windows in heated or cooled rooms
should be insulated. All hinged doors should be hollow
core with a glass panel near eye level. Both frames and
doors should be made of heavy-duty metal. Outside doors
should have sturdy door closers and weatherstrip seals.
Doors between rooms that are maintained at different
levels of cleanliness, air temperature, and relative humidi¬
ty should have automatic bottom seals. Oversized outside
doors are essential for transferring equipment. Interior
doors should also be wide enough to accommodate equip¬
ment, at least 1.1 by 2 m. For restrooms and showers,
0.9-m-wide doors are adequate.

Environmental Control
and Equipment

Some rearing facilities use individual electric or gas
heaters and window air-conditioner units for each room.
But this approach is neither the most desirable nor most
economical for large operations. Central heating and cool¬
ing are preferable. Typically, hot water is the source of
heat, and cold water is used for cooling. The water flows
through coils that are positioned in the ducts of the air¬
handling system; a blower circulates the air. The water is
heated with steam or some type of fuel or is cooled to
about 8° C with a refrigeration-liquid chiller. Controls are
available for modulating the flow of water through the
coils to maintain the proper air temperature. In egg-
production, insect-development, and adult-emergence
rooms, the differential of the controlling device should
not be more than 2° C and about 4% relative humidity.
Individual air-handling systems can be used to provide
different temperature, relative humidity, and sanitation
levels in the facility. For example, 12 units are installed
in the R. T. Gast Rearing Facility. Fresh air is brought in
and conditioned before being mixed with the recirculated
air. An exhaust system is essential, and return outlets
must be located to provide for a uniform air movement,
temperature, and relative humidity throughout the
rooms.

Cooling coils may be used to partially dehumidify. But,
since the vapor pressure of air passing the cooling coils is
not reduced very much, supplementary dehumidifying
equipment is usually necessary. The heating coil is placed
downwind from the cooling coils when they are used to
dehumidify the air. The most desirable way to add
moisture is to use humidifiers that inject steam into the
duct. An override humidistat should be located in the air
duct on the room side of the humidifier to prevent occur¬
rence of too much moisture. Fogging nozzles and rotating
watering wheels or drums are less desirable than a steam
injector because they often cause excessive fungal and
algal growth. Also, it is important to insulate the outside

rather than the inside of the duct. The system’s control
sensors for air temperature and relative humidity should
be located to sense average conditions of the air in a
room.

Light should furnish proper illumination for work to
satisfy requirements of the insects, either from natural
light through windows, from artificial light, or from both.
Light fixtures should meet their use requirements (be
dust, moisture, and insect proof). Fluorescent lighting is
more efficient than incandescent in output per unit of
electrical energy. For insects that require both light and
darkness during each 24-hour period, lights can be con¬
trolled by a time switch. Care must be given to the inten¬
sity and distribution of light if the insects are photo-
trophic. A natural or artificial high-intensity source can
cause some insects, such as the boll weevil, to crowd
toward the light. Weevils spread more uniformly over the
diet pellets when exposed to uniform, indirect lighting or
to total darkness. But high-intensity artificial light is
sometimes helpful in insect rearing. For example, it is
used to attract weevils from the rearing medium after
they have developed to the adult stage.

Air movement is critical in production rooms. For ex¬
ample, too much air movement around rearing trays in a
boll weevil development room dries the rearing medium
too fast and causes nonuniform development and lower
yields. Similar problems occur in weevil oviposition and
emergence rooms. In rooms where sanitation is critical,
air supply outlets from the conditioning system should be
covered with hospital grade (minimum) HEPA (high-
efficiency particulate air) filters. The return-air opening in
all rooms must have a standard furnace filter. The added
static pressure caused by the HEPA filters must be con¬
sidered in selecting blowers for the air-handling system.
Some HEPA filters require 2.54-cm wg static pressure to
maintain their rated airflow. Laminar-flow cleanrooms
should be used for handling diets and other materials
that must remain free of microbial contamination. Also,
the rearing facility should be kept as clean as possible by
good housekeeping practices to minimize dependence on
air filters. Some insects shed scales and other particles
that can be harmful or at least annoying to workers
(Mangum et al. 1969). If such insects are present, air
must be recycled through filters often enough to keep it
clean.

Airlock, passthrough cabinets may be used for passing
materials between rooms in areas where traffic is re¬
stricted for sanitation purposes. These passthroughs may
contain ultraviolet lamps to help reduce transfer of air¬
borne micro-organisms. Proper types and amounts of
weather-stripping and sealing materials must be installed
on all doors. Rearing facilities often require autoclaves
and ethylene oxide fumigation chambers. These units can

72

have a door on each end and can be used between rooms
as passthroughs. Formaldehyde gas fumigation chambers
used in the R. T. Gast Rearing Facility are gastight and
exhausted through the roof. Each fumigating chamber is
controlled automatically with a timer and an actuator on
the dampers of the exhaust and fresh-air outlets.

A central vacuum system for wet-dry cleaning of floors
would be helpful, especially for insects that require a high
degree of sanitation. The lines for the vacuum system
should be made of a noncorrosive material. Compressed
air is needed for some operations and equipment. There
should be a backup compressor and motor for the system.
An air dryer and filter should be used in the lines.

Several details are important to maintaining the neces¬
sary environment and equipment in a rearing facility.
Control of the environment is lost if central heating or
cooling breaks down; a backup unit is recommended for
each of these. Some rearing operations require pure
water; so a distilling unit, holding tank, and distribution
lines have to be installed. Sinks and benches should be
made of heavy-duty stainless steel with rounded edges
for cleaning and safety; drainboards may also be useful.
Smaller pieces of equipment and materials should be
stored on open racks or tables. Refrigerators and freezers
should be made of stainless steel to withstand the clean¬

ing and sanitizing agents. The compressors for walk-in
units located in dusty places (such as those units used to
store or prepare diet ingredients) should be installed in an
adjacent clean area. Fire extinguishers of proper type
should be located conveniently in the facility. Safety
lights that activate automatically should be provided. An
internal communication system that can be used without
any manual operation is important for coordinating work
performed in separated areas (such as diet sterilization
and placement); workers should have outside communica¬
tion equipment when they are restricted to a certain area
of the building.

References

Gantt, C. W.; Brewer, F. D.; and Martin, D. F.

1978. Modified facility for host and parasitoid rear¬
ing. In N. C. Leppla and T. R. Ashley (eds.),
Facilities for Insect Research and Production,
pp. 70-72. U.S. Dep. Agric. Tech. Bull. 1576.
Leppla, N. C., and Ashley, T. R. (eds.).

1978. Facilities for insect research and production.
U.S. Dep. Agric. Tech. Bull. 1576, 86 pp.
Mangum, C. C.; Ridgway, W. O.; and Brazzel, J. R.

1969. Large scale laboratory production of the pink
bollworm for sterilization programs. U.S.
Agric. Res. Serv. [Rep.] ARS 81-35, 7 pp.

73

Automation in Insect Rearing

By E. A. Harrell1 and C. W. Gantt2

Insect-rearing procedures are slowly but surely being auto¬
mated, or mechanized, because demands for laboratory-
reared insects are growing and costs are increasing rapid¬
ly. “Automation” is the substitution of mechanical or
electrical devices for human labor; “mechanization” is
basically the same process, and we will use the two terms
interchangeably here. The decision to automate, in most
instances, results from attempts to complete a difficult
task as quickly and cheaply as possible, not necessarily
to replace labor as is sometimes thought. In insect rear¬
ing, automation may increase speed, reduce labor,
eliminate human error, make operations uniform, improve
insect quality, and increase the number of insects reared.

Many industries, especially electronics, have made
tremendous progress in automation. Recently, the elec¬
tronics industry has developed sophisticated solid-state
controls, chips, and microprocessors. Solid-state controls
to regulate temperature are becoming common in insect
rearing. Microprocessors are not used so commonly, but
they will be in the near future.

As late as the midsixties, most insects reared in small
numbers were laboratory-reared on plant parts for food.
With the advent of laboratory diets (meridic media),
many techniques and apparatuses were developed for
their preparation and handling. As the diets, equipment,
and techniques became more sophisticated, usually the
number of insects reared increased, and the cost per in¬
sect decreased.

For instance, an output of corn earworm, Heliothis zea
(Boddie), was improved from about 500 to 2,000 per day
by modifying and adapting a food-packaging machine to
dispense a diet (Burton et al. 1966). This machine posi¬
tioned 30-ml cups and metered diet into them in a con¬
tinuous operation. To further use the potential of the
packaging machine, equipment was designed and built to
place a predetermined number of eggs in a cavity
(pickout) on the cap (Harrell et al. 1970) and restack the
caps mechanically so that they could be placed on the
cups by the packaging machine. This innovation in¬
creased the corn earworm output to about 30,000 per day.

'Research agricultural engineer, retired, Southern Grain Insects
Research Laboratory, Agricultural Research Service, U.S.
Department of Agriculture, Tifton, Ga. 31793.

“Research agricultural engineer, retired, Southern Field Crop In¬
sect Management Laboratory, Agricultural Research Service,
U.S. Department of Agriculture, Stoneville, Miss. 38776.

With the increased rearing capability, collecting pupae by
hand from rearing cups became almost impossible. So
researchers designed and built a mechanical pupae collec¬
tor with a capacity of about 5,000 cups per hour (Harrell
et al. 1969). It required one operator and increased his
output tenfold. The mechanical pupae collector was 90%
efficient, which was adequate but not as good as manual
collection. Up to this point in the mechanization of com
earworm rearing, production was almost doubled and the
cost per insect reared reduced by about half. Corn ear-
worm rearing was further automated by adapting a 381-
cm-long by 64-cm-wide by 127-cm-high inline form-fill-
seal machine (Sparks and Harrell 1976). Powered elec¬
trically and pneumatically, the machine forms plastic into
a continuous web of rearing cells, heat-seals a cover over
them, and shears the web into desired lengths. The
machine has been synchronized to accomplish the simul¬
taneous operations of forming, sealing, and shearing at a
maximum of 17 strokes per minute. A diet- and egg¬
filling station was designed, built, and installed between
the forming and sealing heads to make a continuously
automated process with a capacity of about 160,000 com
earworm rearing cells per 7-hour run (Harrell et al. 1973;
Harrell, Sparks, Perkins, and Hare 1974). A diet propor¬
tioning, mixing, and sterilizing system was semiauto-
mated to supply the form-fill-seal machine (Harrell,
Sparks, Hare, and Perkins 1974). Also, a collector was
designed and built to remove pupae from this type of
cell. Corn earworm diet is dispensed into individual rear¬
ing cells while it is hot (42° C) and fluid. Eggs are placed
in small, adjacent connecting cells where they are pro¬
tected from the hot diet and microbial inhibitors. Our un¬
published research has shown that these inhibitors injure
eggs of com earworm; fall armyworm, Spodoptera
frugiperda (J. E. Smith); cabbage looper, Trichoplusia ni
(Hiibner); and boll weevil, Anthonomus grandis grandis
Boheman, when they are in contact with the diet before it
solidifies. After the eggs hatch, small larvae move
through the connecting tunnel into the diet-filled cells
and feed. This connecting-cell method works well for in¬
sects with mobile larvae and eggs that can be separated
but not for insects, such as the boll weevil, that are
almost immobile in the larval stage. Several eggs (three
to four) are used per cell, but, because corn earworm lar¬
vae are cannibalistic, usually only one survives per cell.

Perhaps the most widely known and successful effort in
automating insect rearing is that for the program rearing
the screwworm, Cochliomyia hominivorax (Coquerel). The
original program, begun in the midfifties and improved
until the early sixties, produced enough flies to eradicate
the screwworm from the southwestern United States (for

74

details about the rearing procedures, see Goodenough and
Brown 1976).

Between 1975 and 1978, boll weevil rearing received an
extraordinary amount of engineering effort for automa¬
tion. The petri dishes that had served as rearing con¬
tainers, and the associated manual manipulations, were
replaced by plastic trays that were formed, filled, and
sealed in a continuous process (Harrell et al. 1977). The
equipment was similar to that used for rearing the com
earworm except that the auxiliary devices and rearing
trays were designed specifically for the boll weevil. Also,
the diet was rapidly cooled to form a surface skin before
the eggs were added. The eggs were sprayed on the diet
in a uniform pattern and covered with a granular material
that absorbed the carrier liquid and provided support for
the young larvae. Equipment currently developed has the
capacity to process enough trays to yield about 1 million
weevils per hour (Harrell et al. 1980).

Progress has also been made in mechanizing processes
associated with the weevil brood colony. Adult diet is
prepared, sterilized, made into pellets, and covered with
wax semiautomatically (Griffin and Lindig 1974). In this
process, diet is pumped through a sterilizer and into the
pellet-making machine where it is cooled, solidified, pushed
out of pipes, and sheared into the desired lengths. The
pellets then pass through a vat of hot wax where they are
coated. These wax-coated pellets provide food and
oviposition sites for the boll weevil. After the eggs are
laid in the pellets, the wax is separated, recovered, and
returned for recycling. The eggs are then mechanically
removed from the pellets (Griffin and Lindig 1977).

Some of the time-consuming and tiresome jobs of rearing
systems such as implanting eggs in cells by hand, hand¬
ling the rearing trays on a conveyor, and metering
specified numbers of eggs into the cells are being auto¬
mated. A system has been developed for handling eggs
without hand labor (McWilliams et al. 1980). It uses
oviposition cages that provide for adult emergence,
feeding, and oviposition. The cages are housed in a
chamber that confines and collects moth scales, which are
a potential human health hazard.

Many other insect-rearing procedures have been partly
automated. For example, Gantt et al. (1976) reported that
a valveless piston-type pump was developed and used in
a project for suppressing the sugarcane borer, Diatraea
saccharalis (Fabricius). The pump dispensed maggots col¬
lected from a tachinid Lixophaga diatraeae (Townsend) in¬
to host-rearing containers (plastic cups, 22.5-30 ml). This
development increased production 17 times over previous
manual methods.

A procedure that needs to be automated is the stacking

of trays from the boll weevil form-fill-seal machine.
Presently, two or three workers are required to stack the
trays when they are processed at speeds of 30 to 35 trays
per minute. The food industry has the technology
available to mechanically fill a room with racks and
remove processed packages from a machine and stack
them on the racks. Perhaps this system can be adapted
to the needs of boll weevil rearing.

The environment is very important to the survival of an
insect and its progeny; therefore the insectary must be
monitored and controlled within specified limits. Perhaps
the most critical areas are the rooms for oviposition, pro¬
cessing of rearing containers, and larval incubation.
Computer-operated electronic controls are available, but
the degree of control is no better than the sensors. Un¬
fortunately, however, progress on sensor development has
not been as rapid as that on controllers. Sensors for dry-
bulb measurements such as thermocouples, thermistors,
resistance bulbs, and thermostats are dependable and ac¬
curate. Commercially available proportioning controllers
will maintain dry-bulb temperatures very precisely when
used with resistant-type electrical heaters. Control of
relative humidity is more difficult than control of
temperature; it is usually obtained by measuring relative
humidity or dewpoint temperatures. Some of the more
precise sensors require considerable attention and
maintenance. Also, their location must be selected
carefully to satisfy their operational characteristics.
Perhaps the best method for monitoring and controlling
the environment in a room is to saturate air at a known
temperature with water vapor and then change the air
temperature to obtain the desired relative humidity.
Steam can be added to increase it, or it can be lowered
with desiccants or refrigeration equipment.

Automation will probably pay for itself. For example,
mechanization of com earworm rearing (diet preparation,
tray processing, pupal collection, etc.) at the U.S. De¬
partment of Agriculture’s Southern Grain Insects
Research Laboratory at Tifton, Ga., reduced costs by
$8.91/1,000 (late 1970’s). And replacement of petri dishes
with trays for rearing the boll weevil saved about $75 per
hour. Additional savings and increased capacity were
achieved by mechanizing the operations of filling the
trays with diet, adding the sand, and applying the cover.
In these cases, automation was essential because the pro¬
grams required large number of insects daily.

Although those of us involved in rearing insects have
made tremendous progress in automating our procedures,
we have not kept pace with industry, nor have we
adopted all appropriate industrial technology. Unfor¬
tunately, no one has capitalized on opportunities to
design and build an automated rearing facility. Instead,
automation has been piecemeal: the few engineers active-

ly researching this capability have concentrated on
isolated problems rather than on the design of complete
systems. This approach can cause imbalances and bot¬
tlenecks in the sequence of operations; some areas may
not have the capacity to handle the additional workload
created by mechanizing another. When mechanizing some
part of a rearing system, one should account for the com¬
plete rearing regime to achieve a smoothly operating and
efficient system for mass producing quality insects.

References

Burton, R. L.; Harrell, E. A.; Cox, H C; and Hare, W. W.

1966. Devices to facilitate rearing of lepidopterous
larvae. J. Econ. Entomol. 59: 594-596.

Gantt, C. W.; King, E. G.; and Martin, D. F.

1976. New machines for use in a biological insect-
control program. Trans. ASAE 19: 242-243.
Goodenough, J. L., and Brown, H. E.

1976. Screwworm eradication program— procedures
and problems. ASAE Pap. 75-3047.

Griffin, J. G., and Lindig, O. H.

1974. Mechanized production of boll weevil diet
pellets. Trans. ASAE 17: 15, 16-19.

1977. System for mechanical harvesting of boll weevil
eggs from diet pellets. Trans. ASAE 20: 454-
465.

Harrell, E. A.; Burton, R. L.; Hare, W. W.; and Sparks,

A. N.

1969. Collecting corn earworm pupae from rearing
containers. U.S. Agric. Res. Serv. [Rep.] ARS

42-160, 7 pp.

Harrell, E. A.; Burton, R. L.; and Sparks, A. N.

1970. A machine to manipulate corn earworm eggs in
a mass-rearing program. J. Econ. Entomol.

63: 1362-1363.

Harrell, E. A.; Perkins, W. D.; and Sparks, A. N.

1980. Improved equipment and techniques for mech¬
anizing the boll weevil larval rearing system.
Trans. ASAE 23: 1554-1556.

Harrell, E. A.; Perkins, W. D.; Sparks, A. N.; and Moore,
R. F.

1977. Mechanizing techniques for adult boll weevil
Coleoptera : Curculionidae production. Trans.
ASAE 20: 450-453.

Harrell/ E. A.; Sparks, A. N.; Hare, W. W.; and Perkins,
W. D.

1974. Processing diets for mass rearing of insects.
U.S. Agric. Res. Serv. [Rep.] ARS-S-44, 4 pp.
Harrell, E. A.; Sparks, A. N.; Perkins, W. D.; and Hare,
W. W.

1973. An insect diet filler for an inline form-fill-seal
machine. J. Econ. Entomol. 66: 1340-1341.

1974. Equipment to place insect eggs in cells on a
form-fill-seal machine. U.S. Agric. Res. Serv.
[Rep.] ARS-S-42, 4 pp.

McWilliams, J. M.; Gantt, C. W.; and Stadelbacher, E. A.
1980. Heliothis virescens (F.): oviposition and egg col¬
lection system. J. Ga. Ent. Soc. 16: 386-391.
Sparks, A. N., and Harrell, E. A.

1976. Corn earworm rearing mechanization. U.S. Dep.
Agric. Tech. Bull. 1554, 11 pp.

76

Materials Handling in Insect Rearing

By J. L. Goodenough1

Introduction

Insect rearing involves many and varied materials-
handling processes. These range from implanting eggs in
small dishes to receiving semitrailer loads of diet. This
paper considers materials handling as processes; these
are described by action verbs such as “move,”

“combine,” “mix,” and “separate.” Such actions are in¬
volved in the warehousing of materials in diet prepara¬
tion, and in the rearing, packaging, and distribution of in¬
sects. Rearing is divided into egg handling, larval rearing,
pupal handling, maintenance of adult colonies, egg
production, and associated operations. Examples are in¬
cluded here to illustrate present technology (developed
mainly by engineers and biologists working jointly) and
to support recommendations for further work. The refer¬
ence to physical and mechanical properties of both diet
and insects and rheological properties of diet materials
should increase biologists’ understanding of terms used
by engineers to describe properties of materials. Data on
these properties, which are essential to food engineers,
barely exist in the field of insect rearing.

Warehousing of Materials

Is the warehouse nothing more than that leftover space
where we pile junk, or is it intimately related to packag¬
ing, maintenance, and procurement? Careful planning is
required to provide an adequate facility that will include
the necessary space; proper environmental control; and
procedures needed for receiving, storing, and retrieving
all required materials. And planning must account for the
multitude of ways materials are packaged. For storage
facilities and procurement sections to be properly coor¬
dinated, adequate records and inventory control must be
provided. Records and control must insure that the pro¬
curement section is not surprised by last-minute orders
and that the warehouse is not caught without space or
personnel to receive a shipment.

Emphasis on warehousing is evident in reports of large-
scale insect-rearing systems, especially the need to store
diet materials with minimal loss of quality. Harrell and
Griffin (1981) described special warehousing facilities for
storage of the dried diet products, corncob grits, and

‘Agricultural engineer, Pest Control Equipment and Methods
Research Unit, Agricultural Research Service, U.S. Department
of Agriculture, Room 231, Agricultural Engineering Building,
Texas A&M University, College Station, Tex. 77843.

sand needed for mass rearing the boll weevil, An-
thonomus grandis grandis Boheman. They recommend a
building located separately from the main rearing facility
to reduce transfer of dust particles and micro-organisms
into the rearing area and to discourage personnel from
moving between storage and rearing areas without follow¬
ing proper sanitation procedures. Two cool, dry rooms
would be necessary to store the various diet materials;
one of these would have temperature and humidity con¬
trol. A gasoline-engine-driven forklift would be used to
transport diet materials from storage to preparation
areas. Bell et al. (1981) reported that the expanded gypsy
moth, Lymantria dispar (Linnaeus), rearing and virus-
production system included a cool room for storage of
diet ingredients in bulk and a reach-in commercial freezer
for storage of vitamins in bulk and for holding pre¬
weighed batches of dry diet ingredients. The screwworm,
Cochliomyia hominivorax (Coquerel), rearing program
(Goodenough and Brown 1976) had a central warehouse
(remote from the mass-rearing facility) for receiving and
storing dried diet products, packaging materials, and mis¬
cellaneous tools, repair parts, and supplies in bulk and for
dispensing items to the many program elements.

Handling Materials
in Diet Preparation

In the study of materials handling in insect rearing, the
greatest effort has been spent in trying to simplify and
streamline diet preparation. The materials-handling ac¬
tions researched most often include measurement, steril¬
ization, mixing, conveying, and dispensing. Most pro¬
cesses include heating and cooling. Some diets are
pelletized and some are encapsulated. Harrell and Griffin
(1981) reported that grinding and sifting is necessary in
preparing boll weevil diets. Some diets are held in cold
storage after preparation until used (Barnes 1976,
Baumhover et al. 1977).

Various types of controls are used to measure or meter
diet and to adjust diet temperature. Griffin and Lindig
(1974) operated a manifold and controlled valves by hand
in making diet pellets for boll weevils. This operation was
later automated with electric and pneumatic control of
ball valves (Griffin 1979a). An adjustable timer and sole¬
noid valve have been used to control the rate of diet dis¬
pensed with a multiple-species diet dispenser (Gantt and
King 1981). Harrell et al. (1974) metered dry diet
materials with a vibrator-agitator. Gantt and King (1981)
designed and built a special check valve to control drip¬
ping of the applicator nozzle between filling of diet cups.
Temperature control of diet has been reported necessary

77

by Baumhover et al. (1977), Harrell et al. (1977), and
Gantt and King (1981). Bell et al. (1981) noted the ad¬
visability of controlling the entire diet-preparation pro¬
cedure to help maintain uniformity among batches.

Many materials are routinely sterilized during diet
preparation. For rearing the gypsy moth. Bell et al. (1981)
boiled all diet ingredients during processing to inactivate
autolytic enzymes and microbial contaminants that pro¬
mote food spoilage. Diet has been sterilized in production
of the boll weevil (Harrell et al. 1977) as have been gran¬
ular material (Griffin 1978) and diet materials, corncob
grits, and sand (Harrell and Griffin 1981). Griffin et al.
(1974) reported flash sterilization and automatic tempera¬
ture control. Also, Griffin and Lindig (1974) reported
sterilization of diet equipment for producing boll weevil
food pellets, and Griffin (1979b) reported sterilizing the
cooling tunnel used in producing boll weevil diet.

An element of diet preparation common to nearly all
insect-rearing programs is mixing. Bell et al. (1981)
blended gelling agents for gypsy moth diet; then the dry
mixture was blended into the gel with a steam-jacketed
kettle mixer. Small experimental batches were prepared
in a blender. Barnes (1976) reported that in a program for
rearing the Natal fruit fly, Pterandrus rosa (Kuschel), diet
ingredients were mixed first; then water was added, and
the aggregate was mixed well. Mixing of dry diet
materials and then adding water is a procedure also used
by Harrell et al. (1974) for corn earworm, Heliothis zea
(Boddie), diet and by Harrell and Griffin (1981) in prepar¬
ing boll weevil diet. In contrast, Goodenough and Brown
(1976) reported that, in preparation of screwworm diet,
dry products were mixed into water by suction (via a
modified venturi section). And, as the water-nutrient mix¬
ture was recirculated from one of two mix tanks
(2,300-liter capacity each) through the top of an inverted
T-section, measured amounts of dried nutrient products
were vacuumed into the mixture through the stem of the
T; this method provided uniform mixing of the dry ingre¬
dients. Another mixing operation is required in some pro¬
grams to incorporate preservatives.

Large volumes of diet are usually moved by pumping
after they have been mixed. Griffin and Lindig (1974) and
Harrell and Griffin (1981) reported boll weevil diet being
pumped into a pelletizing machine and also from a
sterilizer to the holding tank of a filler for larval rearing
trays. Harrell et al. (1974) used cool sterile diet pumped
into a diet filler for rearing the corn earworm. Gantt and
King (1981) pumped high-temperature insect diet into
rearing cups with a multispecies diet dispenser. Good-
enough and Brown (1976) reported that liquid diet for
screwworms was pumped from mixing tanks through a
recirculating system and sprayed onto larvae-rearing

vats; continuous agitation in the recirculating system
kept the diet materials in suspension.

The final stage in diet preparation for at least two species
is pelletization. Griffin and Lindig (1974) reported
mechanization of this process for boll weevil diet. Griffin
(1979a) reported an automatic control and manifold for
producing the food pellets. Also, to provide an oviposi-
tion substrate, Griffin and Lindig (1974) reported coating
adult boll weevil diet with wax. A diet formulation for
rearing the common green lacewing, Chrysopa carnea
Stephens, was reported by Martin et al. (1978) who
encapsulated the diet in a shell of four waxes.

Egg Handling
Collecting eggs

There are nearly as many methods of collecting eggs as
there are systems to produce them. Griffin and Lindig
(1977) and Harrell and Griffin (1981) reported harvesting
boll weevil eggs after adult weevils had oviposited in food
pellets. Egg-implanted pellets were vibrated into a
“cracker,” which broke the wax coating either with a
rotary chopper or by forcing pellets between two rollers.
After the wax was skimmed or floated off, water, diet
particles, and eggs were pumped into an egg-diet
separator, where the eggs were recovered by a series of
wire-mesh screens and water rinses and cleaned with
saturated NaCl-brine solution. Harrell and Griffin (1981)
included a system for recycling the reclaimed wax. Bell et
al. (1981) reported egg masses of the gypsy moth being
scraped from removable paper liners placed inside heavy-
duty paper oviposition containers. Morrison and Hoffman
(1976) collected Angoumois grain moth eggs by vigorous¬
ly brushing them from nylon screen. Reeves (1975)
reported common green lacewing eggs being collected
from paper cage liners; a cage-liner cutter and two mobile
racks held cage liners after diet application until installa¬
tion in cages; the racks were also used to hold liners
removed from cages until eggs could be collected, thus
significantly reducing space and labor required to handle
them. Screwworm eggs were scraped by hand from
wooden oviposition frames with kitchen spatulas (Good-
enough and Brown 1976). Barnes (1976) reported most
eggs of the Natal fruit fly laid through filter paper being
easily soaked off in a beaker of water and collected by
filtration. Tobacco hornworm eggs laid on artificial leaves
can be easily removed by hand (Baumhover et al. 1977).

Harvest of Angoumois grain moth eggs included remov¬
ing insect scales mixed with eggs by using a screen
shaker (Reeves 1975); scales were eliminated from the air
in oviposition and egg-collection rooms with an air-filter
system; hoods were placed over the cage-holding area and

78

egg-collection table; a high-pressure centrifugal fan pulled
air laden with insect scales from the hoods through ducts
and through a single-stage, impingement-type wet-gas
scrubber, that collected 99% of the scales. Mitchell et al.
(1975) cleaned fruit fly (Tephritidae spp.) egg-production
cabinets with steam because, in addition to cleaning, it
killed any remaining adults and prevented their escape.

Surface sterilization of eggs

Egg preparation for implanting on diet material may in¬
clude surface sterilization. For example, Bell et al. (1981)
reported using a 10% Formalin (formaldehyde) solution to
remove virus from gypsy moth eggs, and Harrell and
Griffin (1981) found that boll weevil eggs have to be
surface-sterilized. Baumhover et al. (1977) reported tobac¬
co hornworm eggs being surface-sterilized, rinsed, and
dried.

Measuring eggs

Eggs are usually measured in some way. For example,
measurement of tobacco hornworm eggs before implant¬
ing was reported by Baumhover et al. (1977) to be by
weight or volume. Goodenough and Brown (1976) re¬
ported screwworm eggs being weighed on a pan balance,
and Mitchell et al. (1965) reported measuring fruit fly
(several species of Tephritidae) eggs volumetrically in
water.

Dispensing eggs

Sparks and Harrell (1976) reported the first mechanized
dispensing of com earworm eggs; casein glue was
dispensed on paper bottle caps, and then six to eight
eggs were placed on the glue. The machine then inverted
and stacked the caps. In a later development, a vibratory
feeder was used to add eggs to cells formed by a form-fill-
seal machine. Bell et al. (1981) reported gypsy moth eggs
being placed into petri dishes. Barnes (1976) placed Natal
fruit fly eggs on artificial diet medium just before they
hatched. Goodenough and Brown (1976) reported 7-9 lots
of screwworm eggs being placed onto damp paper towel¬
ing in petri dishes. And boll weevil eggs were sprayed on¬
to diet as part of a system using a form-fill-seal machine
(Griffin 1978, Harrell and Griffin 1981); the machine first
formed plastic trays and dispensed diet into them; then,
measured amounts of eggs were sprayed onto the diet;
the eggs were uniformly suspended by maintaining the
proper specific gravity of a starch solution (Gast 1966,
Griffin 1978); after eggs were dispensed, the diet was
covered with a granular material. This unit appears to be
the most automated system yet developed for dispensing
rearing medium and implanting eggs on it.

Incubating or storing eggs

Egg-handling procedures may include only incubation.
Barnes (1976) reported Natal fruit fly eggs being held in
a saturated atmosphere until just before hatching. Then,
newly hatched larvae were placed on artificial medium.
Screwworm eggs hatch after being held about 12 hours in
an incubator. Excess production may be chilled and held
for short periods (Goodenough and Brown 1976). The
longest period for chilling and holding egg masses that I
have found is that of 5-6 months in rearing gypsy moths
(Bell et al. 1981).

Handling Materials
for Rearing Larvae

Handling larvae

Goodenough and Brown (1976) reported that in the screw¬
worm rearing program, soon after eggs hatched, the lar¬
vae were transferred from petri dishes to small pans con¬
taining starting medium. After 24 hours, these larvae
were transferred from the small starting pans to large
pans and later to 1.2- by 1.8-m vats. The rearing vats
were stacked five per rack onto racks suspended from a
monorail. Developing screwworms were moved pro¬
gressively through rearing, pupation, and pupae holding
areas on the monorail system. In the gypsy moth rearing
program, newly hatched larvae were placed into 180-ml
diet cups by brushing (Bell et al. 1981) and then trans¬
ferred to the main rearing room. After 21 days, larvae
were transferred to 500-ml cups that contained fresh diet
(see below). In contrast, Harrell et al. (1974) and Sparks
and Harrell (1976) reported that food was already pro¬
vided before corn earworm eggs were metered into rear¬
ing cups. To facilitate transporting the rearing medium
after eggs were added, manual stacking was required for
com earworm rearing cups and trays (Sparks and Harrell
1976) and for boll weevil rearing trays (Harrell et al.

1977; Harrell and Griffin 1981).

Feeding larvae

Supplemental feeding is required in rearing screwworm
larvae. Goodenough and Brown (1976) reported that liq¬
uid screwworm diet was added to larval rearing vats at
4-hour intervals. The liquid diet was added by feeder lines
of a system that continuously circulated the diet. Several
researchers have reported adding diet only at the initial
filling of rearing containers (Baumhover et al. 1977, in
rearing tobacco hornworms; Barnes 1976, in rearing
Natal fruit fly; Sparks and Harrell 1976, in rearing
Heliothis spp.; and Harrell et al. 1977 and Griffin et al.
1979, in rearing boll weevils). Because of high amount of
labor required and difficulty in applying conventional

79

automated rearing technology, Bell et al. (1981) desired
not to change diet during the long feeding period of gyp¬
sy moth larvae; but, because no available diet was nutri¬
tionally stable the required length of time, insects were
usually transferred to fresh diet after 21 days. Although
Bell et al. (1981) had not completed their tests, they did
find that doubling the vitamin mix and boiling all diet in¬
gredients during processing resulted in successful rearing
without transfer of larvae to fresh diet.

Temperature control in rearing larvae

In some rearing systems, as larvae mature, the increasing
amount of metabolic heat produced may cause problems
that require special materials-handling procedures.

Tanaka et al. (1972a) found that metabolic heat had raised
the rearing-vat temperature enough to reduce the rate of
larval growth in Mediterranean fruit fly, Ceratitis
capitata (Wiedemann); they corrected the problem by
moving the trays of larvae into a cooler rearing room,
which resulted in increased yields. Or they could add
water to rearing trays if the cooling units failed. Good-
enough and Brown (1976) reported that rearing-medium
temperature was controlled by disconnecting vat heaters
when screwworm larvae were large enough to produce an
appreciable amount of metabolic heat.

Harvesting larvae

A wide variety of materials-handling procedures have
been used to harvest larvae. Screwworm larvae were
harvested by adding water to the center of the rearing
vats, which induced the mature larvae to crawl off (Good-
enough and Brown 1976). These larvae fell into water
flowing in a gutter, were elevated by jet action (injector),
and then were separated from the water with a shaker.
Larvae moved across the vibrating rack of the shaker and
fell into a bin. They were then scooped onto wire-mesh
screen placed over pupation trays. The larvae crawled
through the screen into a sawdust pupation medium,
leaving trash on the screen. Trays of larvae were trans¬
ferred to the pupation room via monorail. Barnes (1976)
used a combination of inverting rearing trays and low
temperature to induce Natal fruit fly larvae to crawl out
of the medium through gauze and fall into a box of damp
sand. Baumhover et al. (1977) reported that, because of
their cannibalistic behavior, tobacco hornworm larvae had
to be removed from the rearing trays and placed in indi¬
vidual pupation cells.

Sanitation in rearing larvae

Sanitation was reported as critical in corn earworm rear¬
ing by Sparks and Harrell (1976) and has been imperative
in increasing production of gypsy moths and reducing the
necessity to replace larval diet (Bell et al. 1981). General

sanitation operations include cleaning rearing vats (Good-
enough and Brown 1976), cleaning floors and walls,
fumigating the rackveyor (Harrell and Griffin 1981), and
cleaning media cabinets with steam (Mitchell et al. 1965).
In tobacco hornworm rearing (Baumhover et al. 1977), all
equipment was routinely soaked overnight or longer in a
0.5% Chlorox (0.026% sodium hypochlorite) solution for
cleaning and sterilization; the stored equipment was
soaked again just before use; any units that had
unhealthy larvae or active colonies of bacteria or fungi
growing on the diet or fecal pellets were removed from
the rearing area. Sengupta and Yusuf (1974) reported
silkworm, Bombyx mori (Linnaeus), rearing beds being
cleaned once a day and dead and diseased worms being
removed. Sanitation to provide suitable holding condi¬
tions sometimes includes filtering of air. Incoming air is
filtered to remove bacteria and fungi before passing over
boll weevil rearing trays (Griffin 1979b). Filtering is done
within the airflow system used in the room for rearing
gypsy moth larvae to remove insect hairs and scales and
to reduce the incidence of airborne microbial con¬
taminants (Bell et al. 1981).

Materials Handling for Pupation

Handling pupae

Generally, after trays of pupae are loaded into holding
racks, they are transported into holding rooms and held
for further development. Some systems require that
pupae be separated from holding medium before being ir¬
radiated, packaged, or returned to the adult colony.
Young et al. (1980) separated corn earworm pupae from
unused diet, spent diet, and frass residue by floating the
insects in water at negative pressures of 380- to 610-mm
Hg. And Gross and Young (1978) reported that dead corn
earworm pupae and pupae that produce deformed adults
float without vacuum. Tobacco hornworm pupae were
held in pupation cells for 7 days and then transferred to
screen trays in single layers, or prepupae were placed in a
moist soil substitute (vermiculite and sand, 1:1) and
later sifted (Baumhover et al. 1977).

In the screwworm rearing program, after 12-20 hours in
the pupation room, trays containing pupae, larvae, and
sawdust were emptied onto a shaker-separator, which
separated sawdust from pupae and larvae (Baumhover et
al. 1966, Goodenough and Brown 1976). Sawdust was
recycled until it became too dirty to use. Larvae and
pupae were conveyed on a continuous belt through a tun¬
nel of fluorescent lamps, which caused many of the re¬
maining larvae to crawl off, fall into pans, and be recy¬
cled through the pupation room. Pupae were loaded into
screen trays, stacked on racks, and moved through the
holding room for 5Vi days by monorail. After this holding
period, they were either recycled to the adult colony or

80

loaded into stainless steel canisters and irradiated (before
leaving the fly security area). After irradiation they were
conveyed directly to the packaging area. Mechanical
shakers have also been used to separate corn earworm
pupae from trash (Sparks and Harrell 1976), and fruit fly
pupae from vermiculite (Mitchell et al. 1965).

Sexing pupae

One of the most tedious materials-handling tasks re¬
quired in rearing insects is the separation of pupae by
sex. Several workable aids have been developed, but
much data are needed to aid in the design of this equip¬
ment. A mechanical sexing aid (Wolf et al. 1972) was
used to orient cabbage looper, Trichoplusia ni (Hiibner),
pupae for rapid identification through a microscope. A
sizing machine developed by Schoenleber et al. (1970)
used rollers to divide codling moth, Laspeyresia pomon-
ella (Linnaeus), pupae into 10 groups by size as an aid in
sexing them. Whitten (1969) reported mechanically sex¬
ing pupae of the Australian sheep blow fly, Lucilia
cuprina Wiedemann, by differentiating between the light
transmission of the male (both sexes are normally brown)
and female (genetically induced black) pupae.

Sanitation in handling pupae

Sanitation procedures in pupal handling have included
screwworm pupal trays being sterilized in a steam
cabinet while still on the monorail (Goodenough and
Brown 1976) and soaking of tobacco hornworm pupation
units (Baumhover et al. 1977). Since protection provided
by soaking in a sodium hypochlorite solution was short
lived, the tobacco hornworm units were held for 2-3
seconds in a 1.5 g/liter solution of streptomycin sulfate to
provide a residue of antibiotic.

Materials Handling
for the Adult Colony

A special facility may be provided for maintaining an
adult colony and obtaining eggs. It may include holding
cages, feeding devices, oviposition stimuli, and equipment
for disposal of spent insects and materials. Holding cages
range in size from 11.4-liter cylindrical containers made
of fiber and lined with paper (ice-cream cartons, see
Reeves 1975 or Bell et al. 1981) to 1.2- by 1.8- by 2.4-m
cages on rollers (Goodenough and Brown 1976). Some
are equipped with special oviposition devices (see, for
example, Baumhover et al. 1966, 1977; Barnes 1976;
and Goodenough and Brown 1976).

The cleaning of holding cages may require them to be
placed in a chillroom for immobolization of insects
(Goodenough and Brown 1976); cleaning may also re¬
quire special facilities for cleanup (Harrell and Griffin

1981) and equipment for disposing of spent adults and
diet (Morrison and Hoffman 1976). Environmental con¬
trol may include special equipment to remove dust and
scales (Sparks and Harrell 1976); microbes (Harrell and
Griffin 1981); or hair, scales, and moths (Bell et al.

1981).

Materials Handling in Packaging

Materials-handling procedures for packaging depend on
whether individual or bulk handling is desired, stage of
the insect, distance to be moved, and time needed for
moving. An appropriate container can be designed when
these and the range in ambient weather conditions are
known. Richards (1961) expressed density of packing as
numbers of nearest neighbors surrounding an atom in a
crystal. The hexagonal close-pack configuration is the
densest possible arrangement and has 12 nearest equi¬
distant neighbors. Thus, Petterson and DeBolt (1976)
reduced space needed to rear and transport cabbage
looper larvae by using Hexcel trays instead of cups.
Mclnnes et al. (1976) reported that tachinid puparia
were held in individual gelatin capsules that, in turn,
were placed in holes in polystyrene blocks; much hand
labor was needed to place the puparia in the blocks;
but moving, storage, shipment, and identification of
emerged and hyperparasitized puparia was easy.

Ridgway et al. (1977) observed that bulk handling is
desirable because it saves packages, labor, and space.
Bulk handling was first applied to the screwworm and
later, by Higgins (1970), to pink bollworm, Pectiniphora
gossypiella (Saunders). Now it is also being used for
other species. Screwworm pupae were shipped in bulk to
packaging centers where machines formed the box and
automatically dispensed measured amounts of pupae in¬
to it (Goodenough and Brown 1976). But adding a
separator and food, closing the boxes and placing them
on trays, and placing the trays on racks had to be done
by hand. Wolf and Stimmann (1971) developed a
machine for transferring live insects in bulk by a
cyclone principle.

Baumhover et al. (1977) reported that vermiculite and
several other packing materials were tested for shipping
tobacco hornworm pupae and that “sandwiching tobac¬
co hornworm pupae in a 12-layer absorbent paper
cushion gave the best results (only 2.3% loss).” Tanaka
et al. (1972a) reported that using evacuated poly¬
ethylene bags reduced the Mediterranean fruit fly’s
metabolic rate so that pupae remained at ambient
temperature. In fact, emergence was delayed about 2
days, shipping costs were reduced from $10 to $0.50 per
million, and only about half the original space was need¬
ed. The previous method was to ship pupae in shallow,
screened containers placed in cardboard frames.

81

Improved packaging techniques have greatly reduced
labor required to distribute a Trichogramma sp. egg
parasite. Previously distributed by having parasitized
Angoumois grain moth eggs put in small packages,
Trichogramma pretiosum Riley has more recently been
released in bulk. Reeves (1975) reported that a
triangular paper package developed for packaging coffee
cream was incorporated into a machine to mechanically
release T. pretiosum from aircraft. Next, Angoumois
grain moth eggs infested with T. pretiosum pupae were
attached to bran flakes and dispensed with a ground
broadcast unit (Jones et al. 1977). Later, Jones et al.
(1979) and Bouse et al. (1980) reported bulk aerial
release of infested eggs attached to bran flakes. Recent¬
ly, cooled, gearmotor-driven units have been developed
that allow bulk handling and distribution of the eggs
from aircraft without any carrier material (Bouse et al.
1981).

Materials Handling
in Distribution of Insects

Insects are usually moved from rearing facilities to
distribution points by aircraft, especially if long
distances are involved. Distribution may be by ground
or aerial methods, but aerial release is preferred for
large areas and if a high degree of release uniformity is
required. Screwworm flies in boxes were released as
soon as possible when 80% had emerged (Goodenough
and Brown 1976). If release was delayed because of in¬
clement weather, mechanical problems, etc., boxes were
held at 13° C after the 80% level was reached. The
boxes were metered from aircraft by a variable-speed
conveyor. During long periods of inclement weather,
flies could be released from open trucks.

A system for bulk release of screwworm flies that would
use a whirling mass of air has been designed (vortex
principle, C. Husman, personal communication), but not
built. The U.S. Animal and Plant Health Inspection
Service, Mission, Tex. (H. C. Hofmann and others, un¬
published data) has developed and field-tested the re¬
lease of chilled screwworm flies. The chilled-fly tech¬
nique could be used in the regular program relatively
quickly. Some research has also been done on the aerial
release of screwworm pupae (A. B. Broce, P. Nichols,
and associates, unpublished data), but additional tech¬
nology will be needed if pupal releases are to be made
regularly.

Aerial transport of screwworm pupae was reported by
Baumhover et al. (1955) and by Coppedge et al. (1978).
The melon fly, Dacus cucurbitae Coquillet, was trans¬
ported to other programs by aircraft (Tanaka et al.
1972b). Other insects successfully transported by air¬
craft include tobacco hornworm pupae (Baumhover et

al. 1977); corn earworm pupae (Sparks and Harrell
1976); olive fruit fly, Dacus oleae (Gmelin), pupae (Re-
mund et al. 1977); Trichogramma spp. in parasitized
eggs (Ridgway et al. 1977); and eggs in bulk (Bouse
1981). Ridgway et al. (1977) also reported a free drop of
Lixophaga diatraea (Townsend) adults. These flies,
recovered after being chilled and then dropped from air¬
craft, lived as long as untreated flies.

Effects of Physical Properties
of Materials on Handling

Physical properties affect the ease and method of hand¬
ling and storage and the amount of damage that will oc¬
cur during handling and storage. Generally, the greater
a material’s mechanical strength, the more it resists
damage during handling. Mohsenin (1970) discussed
various measurements and tests that can be used to de¬
scribe the structure, physical characteristics (shape,
size, volume, surface area, density, color, and ap¬
pearance), and mechanical and rheological properties of
plant and animal materials; the influence of these prop¬
erties on texture of foods; and their relationship to
aerodynamic and hydrodynamic characteristics and fric¬
tional properties of plant and animal materials.

Mechanical properties are those describing the behavior
of the material under applied forces, such as stress and
strain of a material under static and dynamic loading
and those describing flow characteristics of a material
in air or water. “Rheological” is a term used to describe
mechanical properties when applied forces result in de¬
formation and flow in the material. It is appropriate to
describe diet materials and sometimes insect materials
by either mechanical or rheological properties on the
basis of the type of deformation that results from the
applied loading. The factors involved in rheological
behavior are force, deformation, and time. Properties
used to express changes in materials that may result
from these factors are time-dependent stress and strain
behavior; creep; stress relaxation; viscosity; acoustics;
electromagnetic (including dielectric and magnetic),
photometric, solar, and ionizing energy; and effects of
vibration. As an aid to developing mechanical devices
for processing insects rapidly without injury, Stimmann
et al. (1972), after studying the effects of simulated free
fall on cabbage looper pupae, reported the need for addi¬
tional studies to determine how rheological properties
affect pupae, how abrasion affects pupae, and how the
impacts of pupae affect surfaces softer than those used
in the free-fall simulation tests.

Data on physical properties of materials are useful to
designers of nearly all materials-handling equipment.

The Standard Handbook for Mechanical Engineers
(Baumeister et al. 1978) includes a guide for selecting

82

the type of materials-handling system and a table of what
types of conveyors and elevators are best, depend¬
ing on the physical conditions of the material. Data
useful to formulating insect diets come from the food in¬
dustry. Similar data are needed about physical prop¬
erties of the insects; such data would be especially
useful in designing equipment used when vibration or
shocks are encountered (as during separation from pupa¬
tion medium; during transportation to storage; during
packaging and shipping; and in development of bulk¬
handling systems). Present data and the techniques for
obtaining them are incomplete; so pioneering research is
needed before an adequate data base will be available.

Also important is that the data be reported in a stand¬
ard format so that the values will be readily usable by
others. (This was suggested by Mohsenin 1970; see also
the Agricultural Engineers Yearbook, American Society
of Agricultural Engineers 1982, for tables of “Preferred
Units for Expressing Physical Quantities” and “Radia¬
tion Quantities and Units.”)

Materials-Handling Needs

Great improvements could be made in materials-handling
efficiency through alteration of the biological system. If,
through genetic selection for instance, cannibalistic
behavior in Heliothis spp. or tobacco hornworm larvae
could be reduced or eliminated, rearing efficiency could
increase by an order of magnitude or more. These insects
could be used to produce beneficial insects and virus; and
they might still be suitable for certain sterile-insect pro¬
grams. Other basic simplifications would be enabled by
changes in the actual diet materials, such as complete
removal of agar from the gypsy moth diet to cut costs (a
major problem cited by Bell et al. 1981).

Technology to automatically sex insects would greatly
reduce labor required and permit expanded work in many
programs. Rossler (1975) reported that sexing was re¬
quired for strain research such as single-pair matings,
rearing of hybrids, and field release of only one sex of
Mediterranean fruit fly. Similarly, Baumhover et al.

(1977) reported the need to sex pupae in rearing the
tobacco hornworm. White and Mantey (1977) reported
that sexing was required for codling moths for release-
recapture experiments of irradiated and nonirradiated
laboratory and native moths. The same needs were
prevalent in the screwworm program. The immense task
of manually sexing large numbers of irradiated flies need¬
ed for release and recapture of different sexes of flies in
different areas has caused several studies to be aban¬
doned by the U.S. Agricultural Research Service. The U.S.
Animal and Plant Health Inspection Service in Mission,
Tex., attempted to use the machine described by Schoen-
leber et al. (1970) to sex screwworm pupae by size, but this
technique was unsatisfactory because groups of larvae

reared on different vats varied too much from one another.
A similar problem was reported by Wolf et al. (1972) for
cabbage loopers and by Carlton and Hardee (1974) for boll
weevils and, in both cases, led to development of
mechanical sexing aids for pupae. Bell et al. (1981)
reported that a virus removes gypsy moth females
biologically from some of their wild strains; but the virus
is too detrimental to the mass-production system. Because
female gypsy moths are about three time larger than
males, separation on the basis of size is practical for that
species. Godbee and Franklin (1978) reported that sexing
of the black turpentine beetle, Dendroctonus terebrans
(Olivier), by sound is possible because 87% of the males
but no females make high-pitched chirping sounds. They
reported that the regular method of sexing these beetles is
by shape of the posterior seventh tergite but that extreme
care must be exercised in sexing young adults in this man¬
ner because of possible injury.

Many aspects of handling in rearing should be evaluated
in terms of quality control. Because this subject is treated
in detail by others (see “Putting the Control in Quality
Control in Insect Rearing,” by D. L. Chambers and T. R.
Ashley, and “The Closed-Loop System of Quality Control
in Insect Rearing,” by J. C. Webb.), I do not include it
here.

A variety of other materials-handling needs have been
cited by various authors. Bell et al. (1981) reported that
automating the implanting of eggs on diet and the
harvesting of pupae in the gypsy moth rearing program
could reduce costs from $12/1,000 insects to about
$7/1,000. Harrell et al. (1977) called for the establishment
of optimal larval holding environments. They added that
optimal shape, size, and wall thickness needed to be de¬
termined for larval rearing trays and that a mechanical
system was needed for handling the processed trays
through the larval stage and for harvesting and using the
weevils.

I

Key needs in materials handling for the screwworm rearing
program and other, similar programs are automatic feeding
of larvae and removal of spent media from larval-rearing
vats, mechanized handling of dry materials in the diet-
preparation room, elimination of chilling of oviposition
cages during cleanup, development of technology for a
bulk-release system, application of linear programing to op¬
timize planning or use of other techniques of operations
research (see Gass 1958 and Hillier and Lieberman 1974;
and see “Systems Analysis and Automated Data Pro¬
cessing for Quality Control . . .,” by D. H. Akey, for an ex¬
ample of computerized management of data from an
insect-rearing program), an improved quality-control pro¬
gram including feedback of field data to rearing (see “The
Closed-Loop System of Quality Control in Insect Rearing,”
by J. C. Webb), computerized handling of field-evaluation

83

data, development of techniques allowing release of only
one sex of insect in any particular area (by a biological or
mechanical sexing method), further development of
automated packaging procedures, and close attention to
worker safety (see Snook 1978).

References

American Society of Agricultural Engineers.

1982. Agricultural engineers yearbook. 832 pp. The
Society, St. Joseph, Mich.

Barnes, B. N.

1976. Mass rearing the Natal fruit fly Pterandrus
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Soc. South Afr. 39: 121-124.

Baumeister, T.; Avallone, E. A.; and Baumeister, T., Ill
(eds.).

1978. [Marks’] Standard handbook for mechanical

engineers. 8th ed. 1864 pp. McGraw-Hill Book
Co., New York.

Baumhover, A. H.; Cantelo, W. A.; Hobgo[o]d, J. M.;
Knott, C. M.; and Lam, J. J., Jr.

1977. An improved method for mass rearing the to¬
bacco homworm. U.S. Agric. Res. Serv. [Rep.]
ARS-S-167, 13 pp.

Baumhover, A. H.; Graham, A. J.; Bitter, B. A.; Hopkins,
D. E.; New, W. D.; Dudley, F. H.; and Bushland, R. C.
1955. Screw-worm control through release of steril¬
ized flies. J. Econ. Entomol. 48: 462-466.
Baumhover, A. H.; Husman, C. N.; and Graham, A. J.
1966. Screw-worms. In C. N. Smith (ed.), Insect Col¬
onization and Mass Production, pp. 533-554.
Academic Press, New York.

Bell, R. A.; Owens, C.; Shapiro, M.; and Tardif, J. R.

1981. Development of mass rearing technology. In C.
C. Doane and M. L. McManus (eds.), The Gyp¬
sy Moths: Research Toward Integrated Pest
Management, pp. 599-633. U.S. Dep. Agric.
Tech. Bull. 1584.

Bouse, L. F.; Carlton, J. B.; Jones, S. L.; Morrison, R. K.;
and Abies, J. R.

1980. Broadcast aerial release of an egg parasite for
lepidopterous insect control. Trans. ASAE 23:
1359-1363, 1368.

Bouse, L. F.; Carlton, J. B.; and Morrison, R. K.

1981. Aerial application of insect egg parasites.
Trans. ASAE 14: 1093-1098.

Carlton, J. B., and Hardee, D. D.

1974. Boll weevils: improved techniques in subjective
sexing. Trans. ASAE 17: 656-657.

Coppedge, J. R.; Goodenough, J. L.; Broce, A. B.; Tanna-
hill, F. H.; Snow, J. W.; Crystal, M. M.; and Petersen,

H. D. V.

1978. Evaluation of the screwworm adult suppres¬
sion system (SWASS) on the island of Curacao.
J. Econ. Entomol. 71: 579-584.

Gantt, C. W., and King, Edgar G.

1981. Diet dispenser for a multiple-insect-species
rearing program. Trans. ASAE 24: 194-196.
Gass, Saul I.

1958. Linear programming. 280 pp. McGraw-Hill
Book Co., New York.

Gast, R. T.

1966. A spray technique for implanting boll weevil
eggs on artificial diets. J. Econ. Entomol.

59: 239-240.

Godbee, J. F., Jr., and Franklin, R. T.

1978. Sexing and rearing the black turpentine beetle
(Coleoptera : Scolytidae). Can. Entomol.

110: 1087-1089.

Goodenough, J. L., and Brown, H. E.

1976. Screwworm eradication program— procedures
and problems. ASAE Pap. 76-3047, 11 pp.

Griffin, J. G.

1978. A system for the egg planting operation in boll
weevil mass rearing. Trans. ASAE 21: 469-
472.

1979a. Actuator system for operating small ball
valves. U.S. Sci. Educ. Adm. Adv. Agric.
Technol. South. Ser. 2, 4 pp.

1979b. Equipment for cooling larval diet in a boll

weevil mass-rearing operation. U.S. Sci. Educ.
Adm. Adv. Agric. Technol. South. Ser. 1, 3 pp.
Griffin, J. G., and Lindig, O. H.

1974. Mechanized production of boll weevil diet
pellets. Trans. ASAE 17: 15-16, 19.

1977. System for mechanical harvesting of boll
weevil eggs from diet pellets. Trans. ASAE
20: 454-456.

Griffin, J. G.; Lindig, O. H.; and McLaughlin, R. E.

1974. Flash sterilizers: sterilizing artificial diets for
insects. J. Econ. Entomol. 67: 689.

Griffin, J. G.; Lindig, O. H.; Roberson, J.; and
Sikorowski, P.

1979. System for mass rearing boll weevil in a
laboratory. Mississippi Agric. For. Exp. Stn.
Tech. Bull. 95, 14 pp.

Gross, H. R., and Young, J. R.

1978. Heliothis spp.: method for selecting mature
pupae for irradiation, and performance of H.
zea irradiated as pupae. J. Econ. Entomol.

71: 403-406.

Harrell, E. A., and Griffin, J. G.

1981. Facility for mass rearing of boll weevils.

Engineering aspects. U.S. Sci. Educ. Adm.

Adv. Agric. Technol. South. Ser. 19, 77 pp.
Harrell, E. A.; Perkins, W. D.; Sparks, A. N.; and Moore,

R. F.

1977. Mechanizing techniques for adult boll weevil
Coleoptera : Curculionidae production. Trans.
ASAE 20: 450-453.

84

Harrell, E. A.; Sparks, A. N.; Hare, W. W.; and Perkins,

W. D.

1974. Processing diets for mass rearing of insects.

U.S. Agric. Res. Serv. [Rep.] ARS-S-44, 4 pp.

Haupt, A., and Busvine, J. R.

1968. The effect of overcrowding on the size of

houseflies ( Musca domestica L.). Trans. R.
Entomol. Soc. London 120(15) : 297-311.

Higgins, A. H.

1970. A machine for free aerial release of sterile pink
bollworm moths. U.S. Agric. Res. Serv. [Rep.]
ARS 81-40.

Hillier, F. S., and Lieberman, G. J.

1974. Operations research. 800 pp. Holden-Day, San
Francisco.

Jones, S. L.; Morrison, R. K.; Abies, J. R.; Bouse, L. F.;
Carlton, J. B.; and Bull, D. L.

1979. New techniques for the aerial release of

Trichogramma pretiosum. Southwest. Entomol.
4: 14-19.

Jones, S. L.; Morrison, R. K.; Abies, J. R.; and Bull, D. L.

1977. A new and improved technique for the field
release of Trichogramma pretiosum.

Southwest. Entomol. 2: 210-215.

Mclnnes, R. S.; Albert, D. J; and Alma, P. J.

1976. A new method of shipping and handling
tachinid parasites of Paropsini
(Col. : Chrysomelidae). Entomophaga
21: 367-370.

Martin, P. B.; Ridgway, R. L.; and Schuetze, C. E.

1978. Physical and biological evaluation of an encap¬
sulated diet for rearing Chrysopa camea. Fla.
Entomol. 61: 145-152.

Mitchell, S.; Tanaka, N.; and Steiner, L. F.

1965. Methods of mass culturing melon flies and
oriental and Mediterranean fruit flies. U.S.
Agric. Res. Serv. [Rep.] ARS 33-104, 22 pp.

Mohsenin, N. N.

1970. Physical properties of plant and animal

materials. 755 pp. Pennsylvania State Univer¬
sity Press, University Park, Pa.

Morrison, R. K., and Hoffman, J. R.

1976. An improved method for rearing the

Angoumois grain moth. U.S. Agric. Res. Serv.
[Rep.] ARS-S-104, 5 pp.

Petterson, M. A., and DeBolt, J. W.

1976. Reusable plastic trays for rearing beet ar-
myworms and cabbage loopers. J. Ga. En¬
tomol. Soc. 11: 183-186.

Reeves, B. G.

1975. Design and evaluation of facilities and equip¬
ment for mass production and field release of
an insect parasite and an insect predator.

Ph.D. dissertation, 180 pp. Texas A&M
University, College Station, Tex.

Remund, U.; Boiler, E. F.; Economopoulos, A. P.; and

Tsitsipis, J. A.

1977. Flight performance of Dacus oleae reared on
olives and artificial diet. Z. Angew. Entomol.
82: 330-339.

Richards, C. W.

1961. Engineering materials science. 546 pp.

Wadsworth Publishing Co., San Francisco.

Ridgway, R. L.; King, E. G.; and Carrillo, J. L.

1977. Augmentation of natural enemies for control of
plant pests in the Western Hemisphere. In R.
L. Ridgway and S. B. Vinson (eds.), Biological
Control by Augmentation of Natural Enemies,
pp. 379-416. Plenum Press, New York.

Rossler, Y.

1975. Reproductive differences between laboratory-
reared and field-collected populations of the
Mediterranean fruit fly, Ceratitis capitata.

Ann. Entomol. Soc. Am. 68: 987-991.

Schoenleber, L. G.; Butt, B. A.; and White, L. D.

1970. Equipment and methods for sorting insects by
sex. U.S. Agric. Res. Serv. [Rep.] 42-166, 7 pp.

Sengupta, K., and Yusuf, M. R.

1974. Studies on the effect of spacing during rearing
on different larval and cocoon characters of
some multivoltine breeds of silkworm, Bom by x
mori L. Ind. J. Seric. 13: 11-16.

Snook, S. H.

1978. The design of manual handling tasks.
Ergonomics 21: 963-985.

Sparks, A. N., and Harrell, E. A.

1976. Corn earworm rearing mechanization. U.S.

Dep. Agric. Tech. Bull. 1554, 11 pp.

Stimmann, M. W.; Wolf, W. W.; and Huszar, D. K.

1972. Effect of mechanical impacts on pupae of the
cabbage looper: survival of pupae and condi¬
tion of imago. J. Econ. Entomol. 65: 102-103.

Tanaka, N.; Hart, R. A.; Okamoto, R. Y.; and Steiner,

L. F.

1972a. Control of the excessive metabolic heat pro¬
duced in diet by a high density of larvae of the
Mediterranean fruit fly. J. Econ. Entomol.

65: 866-867.

Tanaka, N.; Ohinata, K.; Chambers, D. L.; and Okamoto,

R.

1972b. Transporting pupae of the melon fly in poly¬
ethylene bags. J. Econ. Entomol. 65: 1727-
1730.

White, L. D., and Mantey, K. D.

1977. Codling moth: mating of irradiated and unir¬
radiated laboratory-reared and native moths in
the field. J. Econ. Entomol. 70: 811-812.

Whitten, M. J.

1969. Automated sexing of pupae and its usefulness
in control by sterile insects. J. Econ. Entomol.
62: 272-273.

Wolf, W. W., and Stimmann, M. W.

85

1971. Cyclone separator for transferring live insects,
and its bological effects on Trichoplusia ni and
Voria ruralis. J. Econ. Entoniol. 64: 1544-1547.

Wolf. W. W.; Stimmann. M. W.; and Parker. G. K.

1972. A mechanism to aid sorting of male and female
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65: 1044-1047.

Young, J. R.: Harrell. E. A.: and Gross. H. R.. Jr.

19S0. Effects of negative pressure on Heliothis zea
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Entomol. Soc. .Am. 73: 78-SO.

S6

The Closed-Loop System of Quality
Control in Insect Rearing

- By J. C. Webb1

1 In recent years, insect rearing has changed from being
| mainly the maintenance of small laboratory cultures to

I operations producing millions of insects each week. With
increases in both the number of insects reared and in the
ij total cost of rearing them, it has become evident that

I good quality-control practices are needed. The objective
of quality control is to insure that a product conforms to
predetermined specifications or standards. To achieve
i this end. the desired characteristics must be known. De-
' velopment of this information is the joint responsibility
I of engineers, entomologists, and administrators.

! A systematic approach is essential in maintaining any
! good quality-control program (King 1975). Such an ap¬
proach would include development of systems and tech-
; niques for conducting and interpreting the quality-control
j process. Many systems and techniques are available: they
range from the simple (making an observation and push¬
ing a button) to the complex (computer-controlled sys¬
tems). But. regardless of its complexity, some basic
elements are common to any system. Control over quality
| depends on the availability of pertinent information on
I which to base decisions. A quality-control system that
regularly provides this information is a feedback or
closed-loop system.

MacFarlane (1964) describes the feedback system as com¬
prising four basic elements: the input, output, feedback
loop, and comparator (fig. 1). The reference input is the
standard that has been set for the product. A sample of
the output is measured and the results fed back for com¬
parison to the standard. Then, if necessary, adjustments
are made in the error variable. In insect rearing (fig. 2),
the main system would consist of all phases of the rear¬
ing process such as egg collection, larvae rearing, and
pupal development. The output, adult insects, would be
sampled and tests conducted on selected factors such as
flight propensity, vision, sound, and mating ability.

These results would be compared to the reference input,
or standard: if they were within the tolerances, no change
would be required in the rearing system. But, if one or all
factors fell outside the specified limits, then some adjust¬
ment would be required.

Research leader, Insect Biophysics Research Unit. Insect
Attractants, Behavior, and Basic Biology Research Laboratory,
Agrictdtural Research Service. U.S. Department of Agriculture.
Gainesville, Fla. 32604.

The concept of a feedback system is not new: in fact,
elements of it have been around for hundreds of years. It
is used in most quality-control programs in industry and
is adaptable for insect rearing, provided the appropriate
input information is known. This kind of feedback system
can be described by the general equation that

Hd‘St3)+B(dS/d^)-f-kS =(5i).

This general equation governs the behavior of any closed-
loop system: the left side describes the output and the
right side the input. The uniqueness of the equation to a
specific system is in the coefficients (I, B, and k). These
are factors that are fed into the system and also the out¬
put that is measured and compared to the standards. So
values for the coefficients must be as accurate as possi¬
ble. (See MacFarlane 1964 for a full discussion of this
kind of equation.)

In insect rearing, the main closed-loop system is com¬
posed of many subsystems (fig. 3). Some subsystems de¬
pend on other subsystems: some are independent. The
combined subsystems maintain each of the variables that
directly affect quality of the adult insects (for example,
pupal weight, temperature of larval medium, light level in
egging cages). The same general equations and feedback
apply to the main system and its incorporated sub¬
systems. For a system to be effective in feedback and
control, we must know what factors are important and be
able to measure them, set their tolerances, and collect
data on them.

Selection of factors to be controlled is the first step in
developing the quality-control system. This selection is
done routinely in any insect-rearing program, whether
systematically or not. The factors selected must have a
direct relationship to the quality of the final product. For
example, there are probably several points along the pro¬
duction line where the temperatures should be recorded
and controlled. It may be important also to control light
quality and quantity in the egg-collection cages. There
may be other subsystems where pupal or larval size are
important. We know that insects reared for a sterile-male-
release program must be able to seek out and mate with
their native counterparts. So they must be able to propel
themselves from one location to another and also to per¬
form the normal courtship ritual. Some of the measurable
factors that may be important for sterile-release insects,
then, are ability to fly, walk, see, call, and produce
sounds. Once a factor has been identified and selected to

87

OUTPUT

FEEDBACK

CONTROLLED

OUTPUT

-

Figure 1.— The ideal feedback system.

STANDARDS
INPUT
ID-

COMPARATOR

MAIN SYSTEM

EGGING

LARVAE REARING
PUPAE DEVELOPMENT

FEEDBACK

SELECTED FACTORS
FLIGHT PROP,
VISION . SOUND,
OTHERS

ADULT

INSECTS

-*-0

Figure 2.— Feedback system for insect rearing.

MAIN SYSTEM

Figure 3.— A main insect-rearing system com¬
posed of several subsystems.

88

go into the system, its quantity and tolerances must be
set. Each factor that is known to affect the quality of the
final product should be included in the system.

Progress has been made in recent years in the basic studies
of insect behavior and behavioral modifiers. From these
studies have emerged some simple tests that can be con¬
ducted to measure aspects of insect behavior. Several of
these tests are now included in the quality assessment of
some mass-rearing and laboratory-rearing programs.

Tests are being done of mating propensity, flight ability,
flight rate, and pupal weight. Research is being con¬
ducted in many other areas. For example, sound is pro¬
duced by many insects, and it has several measurable
variables such as wing beat frequency, pulse information,
and waveform distribution. Visual acuity is another fac¬
tor that holds real promise as an indicator of insect quali¬
ty. Other areas being researched are pheromone response,
pupal size and weight, factors affecting reproduction, and
factors affecting motility. There are two requirements

associated with factor selection; one, that the selected
factors must be quantifiable, and two, that these factors
must relate to the quality of the insect. Once these re¬
quirements are met, the selected factors can be included
in the model.

This has been a discussion of systems theory. In an ac¬
tual operation, many systems, simple or complex, have to
be modified to fit the circumstances.

References

King, J. R.

1975. Production planning and control. An introduc¬
tion to quantitative methods. 403 pp.

Pergamon Press, New York.

MacFarlane, A. G. J.

1964. Engineering systems analysis. 272 pp. Addison-
Wesley Publishing Co., Reading, Mass.

89

Systems Analysis and Modeling in Mass
Rearing and Control of Insects

By D. G. Haile and D. E. Weidhaas1

Introduction

Systems analysis is applicable to practically any field of
endeavor and certainly to mass rearing and control of in¬
sects. Systems analysis and modeling can be particularly
useful in implementation of I PM (integrated pest
management) procedures because the biological systems
and control techniques are so complex. This paper is
limited to application of the systems approach to model¬
ing, or simulation, of insect life histories and population
dynamics with references and examples from our work
with mosquitoes. The basic techniques that we emphasize
can be used as the initial stage in the modeling process
for insect populations in general. These techniques can be
expanded into very complex, computerized models of
population dynamics. We do not try to present a com¬
prehensive review of the many plant-pest models and
computerized systems for implementation of large-scale
I PM techniques.

Modeling can be used for learning and information
assimilation, for focusing attention on the important
variables and components of a complex system and pro¬
moting a better understanding of their interrelationships,
for evaluation of knowledge limitations and research
needs, for simulation experiments, and for prediction of
system behavior under various conditions. These are not
necessarily independent results and certainly do not in¬
clude all possible uses of models. Various authors, such
as Naylor et al. (1966), have discussed these and other
potentially useful aspects of simulation studies in detail.
The rationale for development of simulation models is
generally obvious to most researchers; but in many cases,
the full value or potential of these studies is not fully ap¬
preciated until one is directly involved in development of
a fairly complex model that confirms existing data or
theories. In general, the use of models is directly related
to application of the scientific method. For example,
when a system is so complex that simple observations
cannot be made, models can be used to substitute for
direct observations and aid in development of hypoth¬
eses. Likewise, model results can be compared to actual
data in testing hypotheses. The concept of testing hy-

'Agricultural engineer and laboratory director. Insects Affecting
Man and Animals Research Laboratory, Agricultural Research
Service, U.S. Department of Agriculture, Gainesville, Fla.

32604.

potheses is important when models for insect populations
are considered, since data on the effects of several im¬
portant variables will generally be limited or nonexistent.
The modeling process can be used to combine existing
data with estimates or questionable data on key vari¬
ables. Using modeling in this way will encourage research
on the most important variables where data are question¬
able, until models with sufficient complexity and accu¬
racy for practical applications evolve.

Simplified Modeling Techniques

Perhaps the simplest modeling technique applicable to in¬
sect population dynamics and control uses a growth po¬
tential, or reproduction rate, per generation, Ro, and a
factor representing the effect of a control technique, S, to
relate the population density in one generation P, gen¬
erally thought of as the parent generation, to the next
generation, F. This relationship is

F=P(Ro)(l-S). (1)

If no control is visualized, S=0 and

F>=P(R0). (2)

In this case, the actual change in density (FJP) is equal
to Ro. When a control technique is applied, a distinction
must be made between Ro and the actual growth rate or
change in density (Ra=FJP). So

Ra=FJP=R0{l-S). (3)

In a population with a growth potential of 5 X and a con¬
trol action that is 90% effective for one generation
(S=0.9), the actual growth would be 0.5—
i?a=5(l— 0.9)=0.5— and F,=0.5P. The model can be used
repeatedly for successive generations to establish the
trend of a population over time for a given control
measure. Knipling (1964) used this type of model effec¬
tively to illustrate the difference between the sterility and
insecticidal approach to insect control.

Effective use of this type of model depends on a know¬
ledge of the key variables, Ro and S. The accuracy re¬
quired for the variable estimates depends on the purpose
of the modeling effort and the uses to be made of the
result. Intuitive estimates based on experience or limited
data can be very useful in models to demonstrate the
theoretical potential of a particular control procedure.

90

But, for more critical applications or prediction of actual
population changes, more precise estimates will be re¬
quired, and the results will only be as good as the data
used to develop the estimates. Estimates of Ro for natural
populations may be obtained when data on population
density are available over a period of time and the
generation time is known. In this procedure, the density
ratios at generation intervals provide values of Ro. Cur¬
rently, many survey methods are used for sampling the
various stages of insects, and a large amount of data is
available. Unfortunately, much of this quantity of data is
inadequate for use in analytical models. To be useful, the
survey method must be applied consistently over a suffi¬
cient period of time, and the sampling efficiency of the
method must be constant over time and for different loca¬
tions. Also, the population being surveyed must be un¬
controlled, or the effect of any control actions must be
known. We should not be discouraged by the difficulty of
obtaining meaningful data on these variables and aban¬
don the modeling approach. Instead, we should continue
to build a basic set of data that will allow the modeling
process to become more and more practical.

The equations above represent a useful but elementary
approach to modeling insect population dynamics and
control. More complex models will be needed for the
many systems and control measures required for I PM.
The above equations, then, should be considered as a first
step in a modeling process that can be made as complex
as the system under consideration requires. An extension
of the above modeling approach can be made by in¬
cluding additional variables related to the life cycle or life
history of the insect under study. In a classic life-history
analysis (Birch 1948), the reproduction rate (Rj is related
to survival and progeny production so that

R0=Z(lxmx), (4)

where 1^ =probability of survival to age x from birth
or oviposition

and mx = female progeny produced per live female at
age x.

This relationship can be used to calculate values of Ro
from normal life-table data. Another useful equation for
mosquito populations and several other insects can be
derived from this relationship by applying certain
generalizations common to many insect life histories
(Weidhaas and Haile 1978). First, since oviposition occurs
only in the adult stage, immature survival can be includ¬
ed as a separate variable. Then, assuming that the rates
of adult survival and oviposition are constant with age
and that oviposition occurs at discrete intervals after a
preoviposition period,

R=SjjnKS*)H-Sa', (5)

where S( =probability of survival from egg to emerging
adult,

Sa= average daily survival rate of adult females,

m=average number of female eggs per live female
per oviposition,

d =preoviposition time in days,
and o = laying cycle in days.

In this equation, S, can be subdivided into age classes if
desired. For example, if the development time (in days)
for eggs, E; larvae, L; and pupae, P, are known and a
daily survival rate for each stage (se,s;, and s ) is known,
then s1.=(seB)(s/i)(spp). Also, if a daily survival rate, st, is
given for the complete immature development period, I,
then S.=s/. As a numerical example for equation 5, con¬
sider a mosquito population where S.= 0.853, Sa= 0.85,
m=60, d= 6, and c=3. Then jRo(0.853)(60)(0.856)/1 -0.853
=5. Again, a model such as this is no better than the
estimated variables that are used. And, in most cases,
field data on these variables may not be available. But
the method is available for theoretical calculations and
will be more applicable for practical field problems as the
data base on field populations is accumulated. Currently,
good estimates can be made for values in this equation
for many insect species. Note that the values for Ro in
equation 5 can be used in equations 1 and 3 to calculate
density trends for control measures that limit the repro¬
duction rate.

Since equation 5 includes survival rates, we can make
theoretical calculations from it about other control tech¬
niques that might affect the survival rate of specific
stages. The effect of a control technique must be ex¬
pressed as a constant daily mortality, M, in specific daily
age classes in the insect life cycle. Determining M is
possible for many control techniques, such as continuous
treatment with biological agents, insecticides, or traps
that kill a constant proportion of given stages. For a
given M, a survival rate due to control, Sc, can be
calculated (Sc=l—M) and used in equation 5 to reflect the
actual growth rate, Ra, after the control technique is ap¬
plied. For control techniques that affect the adult popula¬
tion, where Sa is already a daily survival rate, Sa would be
replaced by Sa.Sc and

i?a=(S,)(m)(SQ.Sc)d/l ~(Sa.Scr. (6)

In the case of control techniques that cause deaths in the
immature stages, S; would be multiplied by ScN where N
represents the number of daily age classes affected by the
control. So

Ra=R0ScN. (7)

For example, consider a mosquito population where the
egg, larval, and pupal development times are 2, 6, and 2

91

EGGS

LARVAE

PUPAE

(STERILE EGGS)

NORMAL

MALES

Sa

TOTAL

STERILE

MALES

DAILY

STERILE

MALE

RELEASE

Figure 1.— Diagram of a life-history model of a mosquito with 1-day age classes, including the effect
of sterile-male release. (Stages include: E=e ggs; 1L = first-instar larvae; 2L=second-instar larvae;
3L=third-instar larvae; 4L=fourth-instar larvae; P=pupae; Abnormal adults; S=sterile adults.)

days. If the control affects all larvae equally, then N=6
and Ra=RoSc6. Likewise, if the control affects all eggs or
pupae, Ra=RoSc2. If Sc affects only one age class (1 day),
such as the last day of larvae, then .Ro=ffaSr.

These modeling techniques allow us to make theoretical
calculations interrelating dynamics and control from one
generation to the next. But the mathematics for these
equations require that daily average mortality or sur¬
vivals be used. Unfortunately, many control agents are
not applied every day or for some other reason prevent
the use of average daily survival or mortality. So more
complex modeling techniques are required to give added
realism to theoretical calculations and to simulate the ac¬
tual effects of complex IPM control strategies more
accurately.

Computer Simulation Models

Computer technology can be used to extend the complexi¬
ty and versatility of the life-history approach to modeling
of insect populations. Munro (1973) pointed out that
development of dynamic life-table models represents the

most direct application of computers to insect popula¬
tions. Basically, this technique involves use of a com¬
puter to store an insect life table, which represents the
number of individuals present in each age class for a
given unit of time. A computer program can then be writ¬
ten to calculate changes in numbers and age class, for
each successive time unit based on survival rates and in¬
formation on progeny production from a given initial
population. Haile and Weidhaas (1977) used this approach
for a computer model of a mosquito ( Anopheles
albimanus Wiedemann) population with daily age classes.
This model (fig. 1) also included provisions for simulating
control by release of sterile males. Other control tech¬
niques can easily be programed for this type of model by
establishing how the technique affects the number of in¬
dividuals in given stages, the survival rates, or fecundity.
The structure and complexity of this type of model can
be modified to fit many insects and simulation situations.
But its realism is limited mainly by the use of fixed de¬
velopmental rates and discrete transfers between age
classes (where all individuals of the same age change to
different stages at the same time).

92

= E

t

-EM
t e

- E

t

Q

e

+

A Q
t a

t + 1

* L.

- L,M,

Lt

Q,

+

E,Qe

t + 1

* P,

- P,Mp

- p,

QP

+

L,Q,

't + 1

* A.

- A,Mo

+ p,

QP

Figure 2.— Elementary insect life-cycle model
that used a mortality factor, M, and a flow
rate, Q, to establish the proportion of indivi¬
duals transferring from one stage to the next
during one increment of time, t.

Several approaches have been used to incorporate
variable developmental rates into population models.
Generally, the rate of development is a function of tem¬
perature and can be expressed in different ways, such as
the degree-day approach. Models of the type represented
in fig. 2 use development data to establish the flow rate
of individuals, Q, from one stage to the next during a
given time unit, t. This type of model is simple to use
and easy to program in a computer. Also, the structure is
amenable to various specialized simulation languages
that allow very complex and versatile models to be im¬
plemented easily. This type of model is mainly limited by
the fact that age of individual insects in the various
stages is not differentiated; so a realistic representation
of the development of one cohort of insects is difficult.
Modifications, such as an increase in the number of
stages considered, can reduce this effect; but increasing
the number of stages results in models similar to the
dynamic life table discussed above. A model using a
dynamic life table can be modified to include variable

development and still realistically simulate the develop¬
ment of one cohort as well as overlapping generations.
Fine et al. (1979) developed such a model by varying the
length of time units with the development rate of the in¬
sect. And we are developing a dynamic life-history model
that allows age within stage to vary according to ac¬
cumulated development, which is temperature dependent.
A similar approach is being used for stable fly, Stomoxys
calcitrans (Linnaeus), populations by I. L. Berry (personal
communication).

Basic life-cycle models of insect populations can be the
framework for more comprehensive models that have
many relationships between environmental variables
(such as temperature, humidity, rainfall, and habitat) and
insect variables (such as development, mortality, preda¬
tion, parasitism, fecundity, and density). Many of these
relationships are complex, and the insect variables are
generally interrelated. Also, models of crop pests general¬
ly include some form of a model of plant development
(see, for example, comprehensive plant-pest models, for
cotton insects, reported by Hartstack et al. 1976 and Fye
1979). The various approaches for such models depend
largely on the model objectives and the available or ex¬
pected data base.

Conclusions

Computer simulation is a valuable research and analytical
tool that can and should be used in the development of
insect-control technologies, including those that depend
on mass-reared insects. Models provide valuable insight
into complex systems and promote development and
testing of theories. Because of the complexity of the
systems involved in insect control and the difficulty of
obtaining adequate data, an interdisciplinary team ap¬
proach is essential to effective development and use of
models. Effective model usage also depends on our ability
to identify appropriate problems, establish objectives,
and allocate sufficient resources. With enough field vali¬
dation, population-dynamics models can be used in prac¬
tical I PM programs for predictions of population levels
and timing of appropriate control actions.

References

Birch, L. C.

1948. The intrinsic rate of natural increase of an in¬
sect population. J. Anim. Ecol. 17: 15-26.

Fine, Paul E. M.; Milby, M. M.; and Reeves, W. C.

1979. A general simulation model for genetic control
of mosquito species that fluctuate markedly in
population size. J. Med. Entomol. 16: 189-199.
Fye, Robert E.

1979. Cotton insect populations. Development and

impact of predators and other mortality factors.

93

U.S. Dep. Agric. Tech. Bull. 1592, 65 pp.

Haile, D. G., and Weidhaas, D. E.

1977. Computer simulation of mosquito populations
(. Anopheles albimanus ) for comparing the effec¬
tiveness of control techniques. J. Med. En-
tomol. 13: 553-567.

Hartstack, A. W., Jr.; Witz, J. A.; Hollingsworth, J. P.;

Ridgway, R. L.; and Lopez, J. D.

1976. MOTHZV-2: A computer simulation of

Heliothis zea and Heliothis virescens popula¬
tion dynamics. Users manual. U.S. Agric. Res.
Serv. [Rep.] ARS-S-127, 55 pp.

Knipling, E. F.

1964. The potential role of the sterility method for in¬
sect population control with special reference to
combining this method with conventional

methods. U.S. Agric. Res. Serv. [Rep.] ARS
33-98, 54 pp.

Munro, J.

1973. Some applications of computer modeling in
population suppression by sterile males. In
Computer Models and Application of the Sterile
Male Technique, pp. 81-94. International
Atomic Energy Agency, Vienna.

Naylor, T. H.; Balintfy, J. L.; Burdick, D. S.; and Chu, K.

1966. Computer simulation techniques. 352 pp. John
Wiley and Sons, New York.

Weidhaas, D. E., and Haile, D. G.

1978. A theoretical model to determine the degree of
trapping required for insect population control.
Bull. Entomol. Soc. Am. 24: 18-20.

94

Section 4

Control of Pathogens and Microbial
Contaminants in Insect Rearing

In general, disease has been one of the greatest impedi¬
ments to successful mass rearing of insects, and much of
the history of insect pathology is centered on diseases of
two economically important insects: the honey bee, Apis
mellifera Linnaeus, and the silkworm, Bombyx mori
(Linnaeus). The intent of this section is to emphasize the
impact of micro-organisms on insect cultures and the
measures available to minimize or eliminate the patho¬
gens or contaminants. Special attention is devoted to the
recognition of diseases (and micro-organisms) in insect
rearing since the success of a given control measure may
depend on the micro-organism involved.

In a rearing facility or insectary, the sources of contam¬
inants are numerous and include the diet, the insect, the
rearing facility, and the rearing personnel. The sources of
microbial contaminants are described here in detail as are
measures that may be used to minimize or eliminate the
micro-organisms. In many instances, descriptions of
control measures include discussion not only of how they
affect contaminants and pathogens but also of how they
affect the insect.

Martin Shapiro,

Research entomologist,
Agricultural Research Service

95

Recognition and Diagnosis
of Diseases in Insectaries
and the Effects of Disease
Agents on Insect Biology

by Ronald H. Goodwin1

Introduction

Diseases, or any of several other possible departures from
the normal healthy state, are almost certainly more com¬
mon among insectary-reared insects than among insects
that develop in the field. This is to be expected, since an
insectary environment can never be an exact duplication
of field conditions and usually involves a degree of insect
crowding that never occurs in the field. We must assume
that any artificial rearing system produces a definable
stress on the confined population; and we know that
environmental stress either causes disease directly or
increases susceptibility to disease. Invertebrates such as
insects have a rather rudimentary immune system, and
they usually depend on population dispersal to remain
free of disease. So insectary confinement insures that any
communicable disease agents present will have the best
chance possible to spread in the population. The stress of
the rearing environment insures that individuals contact¬
ing such agents will be easily and rapidly invaded by any
pathogens present in the field-collected starting stock or
in the insectary itself.

Some of the insect diseases to be described and under¬
stood earliest were, not surprisingly, from the first two
major insect-rearing systems, apiculture and sericulture.
Sericulture was the source of the first scientific demon¬
strations of the disease process and of the importance of
microbes in diseases; it was also the basis for the
development of Koch’s postulates (Steinhaus 1949). But
most insect diseases are still unknown, and results of
many present research programs involving reared insects
have been compromised by these undiscovered diseases,
especially the enzootic diseases, in the experimental
stock. Where such diseases have gone undetected, pub¬
lished results contain wrong conclusions about basic
biological descriptions, differential competitiveness, cli¬
matological resistance, parasitoid adaptability and effec¬
tiveness, and other factors such as insecticide resistance
and sterile-male viability. Some recent rearing programs

‘Research entomologist, Rangeland Insect Laboratory, Agricul¬
tural Research Service, U.S. Department of Agriculture, Montana
State University, Bozeman, Mont. 59717.

have included disease prevention, monitoring, and elim¬
ination only after costly failures and much wasted effort.

Many of the major disease agents were originally found
among reared insects rather than in the field. For exam¬
ple, the widely used and highly lethal HD-1 strain of
Bacillus thuringiensis Berliner was discovered in an in¬
sectary for pink bollworms, Pectinophora gossypiella
(Saunders). Because of stock crowding and the other in¬
evitable stresses of suboptimal artificial diets and envi¬
ronmental conditions, insectaries have been a unique
source of new pathogens. The more lethal agents are
relatively easy to determine and eliminate from insec¬
taries. But the enzootic or debilitating diseases are usual¬
ly more of a problem since they often escape discovery
until an insectary-based research program is well under¬
way or, occasionally, even after it has been completed.

Pathogens such as the microsporidia, which apparently
produce insect hormone analogs, have no doubt compro¬
mised many physiological and basic biological studies
where their presence has gone undetected. Indeed, the
microsporidia are probably the most common undetected
pathogens; they are often transmitted transovarially and
may persist indefinitely in a stock colony without ever
quite destroying it. Meanwhile, they produce subtle and
sometimes confusing effects in the host population.

Where imported foreign insects are being quarantined
before release as weed-control agents or as parasitoids for
insect control, particular care is necessary to free the in¬
itial stock from whatever pathogens are present (Bucher
and Harris 1963). Diseased insects of foreign source may
not be able to survive when exposed to a new climate and
to field ecological systems that are inevitably alien to
them and therefore initially stress producing. As a result,
promising species may be prematurely abandoned be¬
cause they were not disease free when introduced. (It
would be counterproductive to introduce pathogens of
natural enemies simultaneously with attempts to intro¬
duce and establish the natural enemy.) Conversely, some
introductions may have failed because the stock was
without its associated symbiotic micro-organisms (that is,
were rendered aposymbiotic: Krieg 1971b, Boush and
Coppel 1974). For example, Stoltz and Vinson (1979)
discovered that certain baculoviruses present in the

96

oviducts of all investigated braconid and ichneumonid
parasitoid wasps may suppress the encapsulation reac¬
tion or otherwise modify the physiology of the usual
wasp hosts; these viruses, therefore, must not be lost
from introduced foreign stock insects through improper
handling if we expect the insects to adapt successfully to
their pest hosts in the new habitat. In general, though
they are widespread in the Insecta, not much is known
about symbiotic micro-organisms in insectary insects or
their potential for insect control (available studies include
Roth and Willis 1960, Brooks 1963, Buchner 1965, Batra
1979, and Houk and Griffiths 1980).

Also, where an investigator is concerned primarily with
importation or colonization of insect predators or
parasites, monitoring of the condition of the prey or host
species may often be inadequate. Some host diseases do
not seem to affect even internal larval parasitoids, which
emerge as apparently normal adults; more often, the
parasitoids are physiologically impaired to some degree.
Again, such relationships have not been studied
adequately.

Programs involving the release of insects that are sterile
or contain a lethal gene depend on the health and com¬
petitive ability of the production stock. So the insectary
should be able to produce a disease-free, physiologically
superior sterile insect since it would have a critical and
decisive advantage when the target adult field population
is large (Hutt 1979).

While individual rearing may be too time consuming for
regular practice, it should be used initially for at least
two or three generations so that the introductory insec¬
tary quarantine will be effective. After the newly col¬
lected insects have been isolated and observed in a
separate quarantine room and adequate procedures have
been developed to prevent cross infections from occurring
in the insectary (see “Micro-organisms as Contaminants
and Pathogens in Insect Rearing,” by Martin Shapiro),
we may assume that the more obvious diseases have been
separated from our insectary seed colony. While some
diseases are highly lethal or otherwise readily obvious,
most diseases are not so easily recognized, and many are
quite cryptic. Insects that are apparently healthy may
often carry these cryptic or enzootic disease agents,
spreading them through contact or contaminated frass or
scales throughout the colony where insects are reared in
groups. Only through close monitoring of the colony,
both macroscopically and microscopically (by sacrificing
unusual or excess specimens), will clues emerge that in¬
dicate the presence of such diseases. Once the offending
disease has been recognized in individually-reared seed-
colony stock insects, it should be eliminated through
careful destruction of the insects containing it, followed
by the rigorous disinfection of contaminated cages, equip¬

ment, and insectary environment (see “Micro-organisms
as Contaminants and Pathogens in Insect Rearing,” by
Martin Shapiro). It is usually not possible to separate
diseased insects from their pathogens. Such attempts are
to be generally discouraged (but see Vavra and Maddox
1976 for microsporidia control).

Disease Recognition

Since the objective of the insect culturist is mainly the
exclusion of disease (see Steinhaus 1953, Helms and
Raun 1971, and “Micro-organisms as Contaminants and
Pathogens in Insect Rearing,” by Martin Shapiro), he is
justifiably more interested in disease recognition (used
here to mean identification of the causal agent) than in
the more complex study of disease diagnosis. The insec¬
tary environment and the artificial diet provided are
often the primary predisposing factors for any given
disease. Therefore, here I will discuss in detail only the
diseases most likely to be a problem in an insectary or in
a similarly artificial insect-production system. Other
disease factors will not be considered; but they must be
taken into account in any final diagnosis decision, which
must include all related factors, not just apparent
primary pathogens. For such additional information and
help in specific areas of disease diagnosis, refer to the
following authors: Ainsworth (1971), a dictionary of fungi;
Batra (1979), insect-fungus symbiosis; Boush and Coppel
(1974) biology of symbiotes; Brooks (1963), micro¬
organisms of healthy insects; Buchner (1965), endosym-
biotes; Burges (1981), eight chapters on the identification
of insect pathogen groups include three on bacteria, one
on viruses, three on fungi (with keys), and one on micro¬
sporidia (with a key); Cantwell (1974), honey bee diseases;
Davidson (1981), symptomatology, pathogenesis, host
resistance, and disease cycles of pathogens and toxins in
insects and other invertebrates; Kramer (1976), dis¬
semination of microsporidia; Lipa (1975), diagnostics and
prevention of insect diseases, chapter 4; Longworth
(1978), small isometric viruses of invertebrates; Mara-
morosch (1977), photomicrographs of viruses; Morse
(1978), honey bee diseases including viruses, rickettsiae,
bacteria, protozoa, and fungi; Muller-Kogler (1965),
diagnosis of insect fungus pathogens, pp. 67-75; Nickle
(1974), nematode infections and biology of nematode fam¬
ilies, with many photographs; Poinar (1975), manual and
host list of insect-nematode associations, with illustrated
keys to families and genera, plus nematode study tech¬
niques; Poinar and Thomas (1978), identification techniques
for fungi, bacteria, viruses, protozoa, and rickettsiae, with
keys to groups and common genera, representative photo¬
micrographs of all pathogen groups mentioned; Roth and
Willis (1960), cockroach microbial relationships including
symbiotes; Shimanuki and Cantwell (1978), diagnosis of
honey bee diseases; Sprague (1977a), current classifica¬
tion of the microsporidia; Sprague (1977b), annotated list

97

of species of microsporidia; Sprague (1977c), zoological
distribution of microsporidia; Steinhaus (1949), symptoms
and pathologies, chapter 8, illustrations from many
microbial groups; Steinhaus (1963), background for
diagnostic work plus a complete step-by-step guide to
diagnostic procedures; Steinhaus (1964), definitions in
diagnosis with a discussion of the history and present un¬
fulfilled requirements for adequate diagnosis of inverte¬
brate diseases; Steinhaus and Martignoni (1970), glossary
of invertebrate pathology; Thomas (1974), a general guide
to the diagnostic equipment and procedures necessary for
each major pathogen group; Torre-Bueno (1973), glossary
of entomology; Vavra and Maddox (1976), the examina¬
tion of insects infected with microsporidia and several
techniques in investigating the microsporidia, with a
short section outlining the control of microsporidial
infections in insects; Weiser (1961), microsporidia in in¬
sects, contains many life-cycle drawings and photomicro¬
graphs showing the developmental stages and spores of
many of the known microsporidians; Weiser (1969), a col¬
lection of photomicrographs of the many agents infec¬
tious to insects, with a few photographs of diseased
whole larvae; Weiser and Briggs (1971), general keys to
pathogen groups, to genera, and to some species based on
gross and microscopic appearance of diseased specimens
and the appearance of diagnostic pathogen stages, dimen¬
sions of micro-organism groups— table 1, (few photo¬
graphs, keys to groups and genera partly outdated);
Whitcomb and Williamson (1979), insect pathogenic
mycoplasmas and spiroplasmas.

Steinhaus (1964) defined symptoms and signs of disease
in insects to include specific and distinctive factors. Dis¬
ease symptoms include abnormal movements, abnormal
responses to stimuli, abnormal functional development,
abnormal body rhythms, digestive disturbances,
reproductive disturbances, variations in longevity, and
odors. Disease signs include discoloration and alteration
of color pattern, abnormalities of body size (and the size
of parts), abnormalities in body shape, abnormalities in
external structures, abnormalities in texture and
sculpturing, abnormalities in consistency, traumata and
wounds, prolapses and hernias, and observable presence
of the pathogen (either macroscopically or microscopical¬
ly). In many diseases, these indicators overlap, so care
must be taken to overlook nothing that could lead to the
correct identification or diagnosis of a given disease.

The first step in diagnosis is to make an external exam¬
ination. This is followed by an internal gross dissection
under a dissecting microscope; the insect is cut open with
a midventral incision and laid out flat with pins so the in¬
ternal anatomy can be seen and studied easily (this is
termed an open dissection).

To make useful fresh wet-mount smears, use only a small
amount of tissue. Suspend it in water on a glass slide and
gently crush the sample with a coverslip placed on top.
The aim is to break up the tissue into individual whole
and broken cells. In some cases (for example, blood and
fat body), the coverslip pressure alone is sufficient to ade¬
quately flatten and spread the cells of a small lump of
tissue so associated microbes can be viewed clearly with
phase objectives. Many pathogens, such as the larger coc-
cidian and gregarine cysts and spores, need no more
treatment than this. Accurate pathogen identifications
can be made from photomicrographs of such preparations
if the magnifications are recorded.

There are several simple-to-complex insect diagnostic
histological and staining techniques (see, for example,
Weiser 1961, Miiller-Kogler 1965, Weiser and Briggs
1971, Cantwell 1974, Thomas 1974, Lipa 1975, Vavra and
Maddox 1976, Poinar and Thomas 1978, and Shimanuki
and Cantwell 1978). Of the many techniques available,
the most generally useful are: (1) fresh wet-mount smears
of blood, fat body, gut, and other tissues (in that order)
observed under high-power and oil-immersion phase ob¬
jectives and (2) Giemsa-stained smears of the same
tissues. If there is an apparent fungus infection, Guegen’s
solution is recommended for help in identification (Thomas
1974, Poinar and Thomas 1978)). Or lactophenol cotton
blue may be used for fungi or for the entomopoxviruses
(Poinar and Thomas 1978; see the preparation technique
for this stain given in the Entomopoxvirus “Microscopic
Examination” section.

Either Giemsa powder (Sigma Chemical Co., St. Louis,
Mo., stock No. G4507) or Giemsa stock solutions are
widely available for use with the Giemsa stain procedure.
If Giemsa powder is used, dissolve 1.25 g of powder in
41.7 g of glycerol (warm glycerol to 60° C and shake
thoroughly to dissolve the powder). Add 125 g of ab¬
solute methanol, shake to mix, and allow to stand over¬
night at room temperature. Filter with a No. 2 Whatman
filter in a vacuum-operated Buchner funnel. This solution
will store indefinitely if water is not added. For a
repeatable stain reaction, dilute 1 ; 20 (or 1 : 25 when the
technique concentration is listed as 1 drop/ml of water;
this equals 4 ml stain stock plus 100 ml of water or buf¬
fer) with buffer (Na2HP04 . 2H20, 1.707 g; KH2P04, 1.88
g; distilled water, 500 ml), not with water. The stain is
unstable when combined with the buffer (or water) and
should be discarded within 24 hours. Air-dried thin
smears on glass slides are first fixed in methanol for 3-5
minutes and then stained in 1 : 20 Giemsa buffer for
about 1 hour (try more or less time depending on results).
Wash smear gently with buffer or distilled water and blot
or air-dry. Giemsa stain modifications and the expected
results with each pathogen group are indicated in the sec¬
tions on microscopic examination.

98

The key below (key 1) to the insect diseases is designed
for use with living insects in an insectary or under condi¬
tions where continuous observation of living specimens is
possible. Starvation, asphyxiation, overheating, drown¬
ing, wounding (including gut or integument breakage),
and poisoning or other chemical assaults must be
separately considered before use of this key (Steinhaus
1953). Dead larvae will usually contain several bacterial
types; once bacterial rotting has occurred, the correct
identification of the primary pathogens that may be pre¬
sent is usually more difficult. Although the described
diseases are usually found in larvae, some of the same
conditions may be found in adults of the holometabolous
insects and in any growth stage of heterometabolous in¬
sects. Where open dissections are described, dissection of
a normal larva should be performed at the same time for
a direct comparison of the condition of the internal
tissues.

Key 1. — Symptoms and signs of insect disease ae seen with the naked eye or a dissecting microscope
below X 80

1 . Stunted larvae, abnormally whitened or yellowed . 2

Stunted larvae, no discoloration . 3

Normal-sized insects showing nervous activity and trembling followed by lack of coordination and

paralysis . Picomaviridae

Normal-sized insects showing abnormal regurgitation and anal discharge (diarrhea) . 4

Normal-sized insects showing discolorations (yellow, pink, red, purple, brown, green, blue) . 5

Insect fails to molt or pupate normally . 6

2. Light-colored stunted larvae (open dissection):

With midgut abnormally whitened . Reoviridae or microsporidia

With fat body abnormal in appearance (distended, fragile, fragmented, or discolored) .

. Entomopoxvirus, microsporidia, Neogregarinida, or Baculoviridae

3. Stunted larvae (no other signs or symptoms) . Chlamydia 2

Stunted larvae showing both regurgitation and anal discharge (sometimes with violent muscular

contractions) . Enter ella (rickettsia)

Stunted larvae showing white fecal exudate . microsporidia

4. Sawfly larvae showing white regurgitate (open dissection reveals abnormally whitened gut) .

. Baculoviridae

Insects showing both regurgitation and copious anal discharge (death may occur in hours after
symptom appearance) . Picomaviridae

5. Pronounced or subtle color changes followed by solidification (mummification) at time of death

(aquatic insects do not solidify) . fungi

Subtle color changes followed by liquefaction of larval contents after death .

. Baculoviridae (nuclear polyhedrosis)

Color changes followed by rotting of cadaver with sweet or putrid odors . bacteria

Color changes persisting after death (decomposition may be delayed) .

. Rickettsiella or Baculoviridae (granulosis)

Larvae with transparent cuticles may show progressive translucence of normally white internal
organs, finally becoming a dirty mottle brown . Coccidia (protozoa)

6. Insect becomes abnormally swollen-elongate (often lighter-colored than normal) .

. Entomopoxvirus, Rickettsiella, Baculoviridae, or microsporidia

Insect dies while molting or pupating or emerges as a deformed adult (chalky appearance) .

. Neogregarinida (protozoa)

Insect dies while molting or pupating, then darkens and rots (sweet or putrid odor) . bacteria

2See “Rickettsiales and Chlamydiales”; see also “Note Added in Proof,” p. 117.

Protozoa (and Microspora)

The one-ceUed animals, or protozoa, until recently con¬
tained among them the subgroup Microsporidia, which
seems to be the most important and widespread group of
pathogens found in insectaries. While Sprague (1977a)
has placed these microbes in the separate phylum Micro¬
spora, this classification may or may not be accepted in
the future. They will be considered as protozoans here for
convenience since all of the earlier literature treats them
as protozoans. Microsporidia and other protozoan patho¬
gens are usually enzootic in field populations. Insects in¬
fected in the field or in the insectary may survive to
reproduce and spread the infection throughout the envi¬
ronment. Often the signs and symptoms are obscure,
especially in insects not having a transparent cuticle.
While originally thought to be rather host specific, proto¬
zoan species in the microsporidia, Coccidia, and Neo-
gregarinida are now known to be able to infect at least
several related insect species and, in some cases, to cross
familial or ordinal host divisions. Indeed, some micro-
sporidians are now known from parasitic Hymenoptera
(Brooks 1974), which implies that their elimination from
introduced parasite species could be a practical means of
improving parasite activity in new habitats.

Microsporidia (Microspora)

Disease cycle.— After microsporidian spores (often from
fecal material of infected insects) are ingested, each for¬
cibly extrudes a hollow polar filament that injects the
first-stage sporoplasm into the gut cell; the sporoplasm
grows intracellularly into the intermediate stage, a
meront. Meronts divide to give further meront stages or
sporonts and sporoblasts, which develop into mature
spores. The meronts may be restricted to the gut or may
pass into other tissues. Specific infections may then be
characterized in part by the presence of meront stages
and/or spores in certain tissues of the host. Many are con¬
fined to the gut and/or the malpighian tubules; others can
be found specifically or more generally infecting the fat
body, tracheal matrix, hypodermis and connective tissue,
blood cells, silk glands, muscles, or gonads. When the in¬
fection is confined to one or two of these tissues, separate
tissue dissections are the only way to confirm the
microsporidian presence if a whole-body smear (possible
with small larvae) is not feasible.

Lysed gut-cell contents release spores from the gut cells
that survive in the fecal material or in regurgitate. Spores
may also escape from the infected insect via egg-cementing
fluids (in female lepidopterans), on the ovipositors of
parasitoids, through cannibalism, or through the rupture
of the infected cadaver and subsequent environmental
contamination. Vertical passage of nonsporulated stages

and spores commonly occurs within the eggs of many in¬
vertebrates (Brooks 1974, Kramer 1976, Vavra 1976).

Symptoms and signs.— According to Vavra and Maddox
(1976), although most infections outwardly demonstrate
no characteristic signs or symptoms, transparent insects
infected with microsporidia are often wholly or partly
milky white within because of spores and growth stages
packed into the infected tissues (for example, fat body,
hypodermis, tracheal matrix, or blood cells; figs. 1A-1D).
Where the insect is not transparent, a gross dissection
may reveal a white distended gut or similarly discolored
malpighian tubules or other tissues as compared to a
similar dissection of a healthy individual (see Cantwell
1974, fig. 11). Infected larvae may show black spotting
(fig. IE), but such spots may also indicate a fungus or
rickettsial infection. The infected insect may be stunted
(fig. IF) or, if molting is delayed, may become an abnor¬
mally large larva or a malformed pupa (figs. 1G and 1H).
Sluggishness is common, as are uncoordinated move¬
ments, abnormal postures, and movements such as
twitching. Infected individuals may reduce their food con¬
sumption or produce a white fecal exudate. Reproduction
rates may or may not be measurably affected.

Microscopic examination. — The intracellular developmen¬
tal stages of the microsporidia are readily stained with
the Giemsa technique (appearing globose, with blue cyto¬
plasm and red, compact-to-vesiculate nuclei; often with
paired nuclei in meront stages), but mature spores will re¬
main unstained. To stain the sporoplasm within the
spore, the smear preparations must first be hydrolyzed
for 10 minutes by treatment with IN HC1 at 60°-70° C,
washed several times with distilled water, and then
stained. The spore will then show a central or partly
polar cytoplasmic blue element within but will not take
the stain uniformly throughout. Oil-immersion observa¬
tion with white light is required because of the small size
of most spores. Consult the illustrated references (Weiser
1961, 1969 and others) for the appearance of the many
types known. The most recent key was written by
Hazard, Ellis, and Joslyn in Burges (1981, chapter 9).

Coccidia

Disease cycle.— Commonly, several nearly spherical cocci-
dian spores (or sporocysts) are released from a globose-
walled oocyst into the environment and are then ingested
by susceptible hosts. The spores burst in the gut, re¬
leasing many sporozoites that penetrate and multiply in
the gut epithelium or pass through and multiply in the
fat body. The many asexually formed, wormlike meront
stages eventually differentiate, pair sexually, and finally
give rise to sporoblasts that develop into the final oocyst
stage. Spore stages are released into the environment

100

through the ruptured cadaver or are spread by can¬
nibalism in some species (Brooks 1974), but they are
most commonly distributed with the feces in gut-specific
species (Weiser 1963).

Symptoms and signs. — Heavily infected larvae may move
more slowly than normal larvae, and their tissues are
more frail (for example, in Adelina tribolii infections). In
Adelina sericesthis infections, intermediate stage de¬
velopment induces an externally visible translucent clari¬
ty (in larval hosts with a transparent cuticle) in portions
of the body where the fat-body tissue has been heavily in¬
fected (fig. II). Later infections in the same insects (near-
moribund hosts) color the integument a mottle-brown
(Brooks 1974), making them look as if they were filled
with vermiculite (fig 2A); in late A. sericesthis infections,
both the fat-body and the subcuticular connective tissue
are infected and turn brownish as the mature oocysts are
formed. One coccidian species that infects the malpighian
tubules causes a clarification of that tissue. Reproductive
rates may be affected in populations having sublethal in¬
fections. Insect-infecting coccidia are not host specific,
but the host ranges of the known species have not been
fully described.

Microscopic examination. — The intracellular developmen¬
tal stages are easily stained with Giemsa, but the fresh
spores are very stain resistant. Fortunately, the
characteristic bag-of-balls oocyst structure is readily ap¬
parent in wet-mount smears of gut or fat-body tissue
viewed with phase optics (see Weiser 1969, figs. 315-317).

Neogregarinida (Schizogregarinida)

Disease cycle.— The ovoid-to-spindle-shaped neogregarine
spores are usually ingested in contaminated food. The
spore wall ruptures in the gut, releasing many elongate,
wormlike sporozoites that penetrate the gut epithelium
where they either multiply or pass through to multiply in
the fat body or malpighian tubules. One species is known
to multiply in the hypodermis. First, a large multinu-
cleate plasmodium is formed; from this, nuclei bud off
and form elongate epicellular merozoites or globular intra¬
cellular merozoites. These further divide to fill the in¬
fected tissue with prespore sexual stages that unite to
form the final spores. The spore stages are released from
the host via the feces in gut or malpighian-tubule-specific
species and via ruptured cadavers and cannibalism in
fatbody- or hypodermal-cell-specific species.

Symptoms and signs.— Larvae infected with the neogre¬
garine Mattesia grandis are often unable to molt or
pupate (as in figs. 1F-1H). Emerging adults are often
deformed or die half emerged. Larvae may be chalky in
appearance because of the production of many spores

that replace the fat-body tissue. Insects infected as
adults show a decreased reproductive rate and longevity.
With other neogregarine species, only the more general
debilitating effects common to protozoan infections give
any indication that disease is present in reared stock. In
these cases, periodic samples of susceptible insect tissues
must be taken to monitor for this pathogen. Neogregarine
species are known with both narrow and wide host
ranges. Some genera of the Eugregarinida ( Stictospora
spp. and others) form spores that are contained in large
hemocoelic cysts (figs. 2B and 2C); these spores are
similar to neogregarine spores. See also Weiser (1969,
figs. 273-276).

Microscopic examination. — As with the coccidia, the de¬
velopmental stages are easily Giemsa-stained, but the
spores are stain resistant (unless pretreated with acid).
Again, phase optics will allow one to distinguish the
ovoid-to-spindle-shaped spores in wet mounts from the
coccidian spore type (Weiser 1969, figs. 277-312).

Viruses

Viruses are submicroscopic, obligate, intracellular patho¬
gens that influence the host cell to replicate their DNA or
RNA (depending on the virus type) rather than the
cellular nucleic acids of the host; they require the living
host cell for their reproduction. At the most primitive
level of life, these microbes cannot grow, since they con¬
tain only their nucleic acid, a protein coat, and a small
number of enzymes. They are more resistant than several
other microbial groups to some environmental conditions,
including freezing; and, once present in an insectary, they
may spread widely in the air as aerosols, on dust par¬
ticles, and commonly on adult insect scales, hairs, and
bristles where cleanliness is not scrupulously maintained.
The question of their spread in populations of insects by
transovarial transmission and "latency” is still not com¬
pletely resolved after years of study (Longworth 1973);
many earlier reports have been contradicted by later
work, particularly in Japan, showing no such occult (or
host-genome-connected benign) virus presence in insects
reared under germ-free conditions.

Viruses pathogenic to insects are classified according to
the criteria established for other animal and plant
viruses. These include the type of nucleic acid in the
virion or mature virus particle, the particle morphology,
the subunit symmetry of the protein coat, the presence or
absence of an envelope surrounding the particle, and the
particle size and its degree of resistance to certain
chemicals (Fenner 1976, Matthews 1979).

The natural classification of insect viruses divides them
firstly into groups containing RNA and DNA and second-

101

ly into those that either are embedded in protein-matrix
bodies when mature or are naked when mature. Of the
several viral families, groups, and genera present in in¬
vertebrates (Fenner 1976, p. 100), I will consider only the
RNA-matrix -embedded Reoviridae (cytoplasmic
polyhedrosis viruses), the RNA-naked Picornaviridae and
similar viruses, the DNA-matrix -embedded Baculoviridae
(granulosis viruses and nuclear polyhedrosis viruses), and
the DNA-matrix -embedded Poxviridae (the genus En-
tomopoxvirus). These four virus groups are the most like¬
ly to cause insectary difficulties or are known to persist
in insect populations over a long period at a low level (en¬
zootic). Most can be expected to cause insidious continual
losses and to debilitate stock insects as do the micro-
sporidia. A few, such as the Antheraea virus and certain
polyhedrosis viruses, can be epizootic or catastrophic in
insectaries. Although individual viruses cannot be seen
with a light microscope, the protein-matrix bodies that
some of them are embedded in can be seen and are here
treated as diagnostic indicators.

Reoviridae (cytoplasmic

polyhedrosis viruses)

Disease cycle. — Sharp-cornered, distinctively shaped cyto¬
plasmic polyhedra (or virus-containing matrix bodies)
from contaminated feces or ruptured cadavers infect lepi-
dopteran, neuropteran, dipteran, or hymenopteran larvae
when ingested. They dissolve in the midgut, releasing
virus particles that invade and replicate in the midgut
epithelium and later often in the hindgut and foregut
cells. Restriction of these viruses to the cytoplasm of the
gut cells of infected larvae is the prime diagnostic
characteristic of this virus group. As the infected midgut
cells lyse, polyhedra may be regurgitated or voided in the
feces.

Symptoms and signs.— The growth of infected larvae is
slowed because of reduced feeding. Infected small larvae
can eventually be recognized by their extreme retardation
compared with normal larvae. The head may become dis-
proportionally large, and an abnormal white or yellowish
coloration may be seen ventrally in some larvae where
the cuticle is more transparent. This coloration is due en¬
tirely to the whitened or yellowish, obviously infected
midgut, which can be seen in open dissection; transparent
infected specimens such as mosquito larvae can be
recognized without dissection (figs. 2D and 2E). Adults
can transmit the virus to their offspring through surface
contamination of the eggs, and infected laboratory col¬
onies may show reduced reproduction rates. These
viruses are widely cross infectious throughout the Lepi-
doptera. Only one each is known from the Neuroptera
and Hymenoptera and only a few from the Diptera.

Microscopic examination. — Wet smears or Giemsa-stained

smears of infected gut tissue should reveal many small
(1-15 /j.m), clear, crystalline matrix bodies (polyhedra)
when viewed by light microscopy. The polyhedra usually
appear rounded since they are generally too small for
their sharp-cornered, characteristic, polyhedral shape to
be discerned except by electron microscopy (Maramo-
rosch 1977). The polyhedra remain clear and unstained in
Giemsa-stained smears, fat globules stain purple to red,
and whole cells become differentially colored in nuclei
(pink) and cytoplasm (purple). Some other crystals, such
as ureate concentrations, will also be stain negative but
are usually much larger than the polyhedra. Use of a IN
HC1 acid hydrolysis step before staining allows differ¬
entiation between cytoplasmic and nuclear polyhedrosis
polyhedra; both may occur in the midguts of hyme-
nopterans and dipterans (see the Baculovirus “Micro¬
scopic Examination” section).

Picornaviridae and similar viruses

The Picornaviridae virus group is here taken to include
the known insect picomaviruses that are nominally placed
in Enterovirus “genus 2” (Longworth 1978). This genus
includes the cricket paralysis viruses and Drosophila C
virus and probably acute bee paralysis virus; sacbrood
virus of honey bees, Apis mellifera Linnaeus; and the
Mansonia uniformis (Theobald), a mosquito, virus. There
are also some similar but incompletely characterized
RNA-virus groups near the Picornaviridae: (1) the
Nudaurelia B group, including the (lepidopteran) viruses
from Nudaurelia cytherea capensis Stoll, Antheraea
eucalypti Scott, Philosamia cynthia Drury, Hyalophora
cecropia Linnaeus, and others; (2) the “Group 5” viruses,
including Drosophila P and A viruses, Kashmir bee virus,
flacherie (silkworm) virus, Gonometa podocarpi
Aurivillius (lasiocampid) virus, and others; (3) the ovoid
viruses, including chronic bee paralysis virus and
Drosophila RS virus. Longworth (1978) characterized
these newer RNA-virus groups and others that are prob¬
ably less important to insect culturists. He emphasized
that we have only discovered a few examples of what will
probably become a quite large and varied series of
invertebrate-virus groups.

Disease cycle. — Since the picornaviruses are not a
discrete group, it is not surprising that they have dif¬
ferent pathologies. Most are dysentery viruses that infect
the guts of larvae; they are usually spread through the
contaminated regurgitate and often copious anal dis¬
charges. Sacbrood virus is usually found in larval fat
body and musculature, but it has recently been found to
infect young adult worker bees and drones through con¬
taminated food. It is now known to be dispersed through
many tissues, including the salivary glands in the adult
bees and in the larvae. Adults, therefore, may feed the
virus to larvae in contaminated pollen (Gochnauer 1978).

102

The chronic and acute bee paralysis viruses have been
found only in adult honey bees or in bumblebees. Acute
paralysis virus is carried benignly by adults in nerve
tissues (brains) but causes a lethal fat-body infection
when transferred between bees by injection. The chronic
paralysis virus is a more obvious debilitating and lethal
nervous-tissue disease of adults that is transmissible by
feeding (Gochnauer 1978).

Symptoms and signs. — The largely lepidopteran dysen¬
tery viruses may be highly lethal (the A. eucalypti virus
kills infected larvae in 12 hours) or debilitating and slow¬
ly lethal. Silkworms, Bombyx mori (Linnaeus), infected
with flacherie virus are noted as weak, flabby, feeble, and
sluggish larvae that become soft and putrified in the later
disease stages (Vaughn 1974). Kellen and Hoffmann
(1980) found that the slower chronic stunt virus (fig. 2F)
replicates only in the granular hemocyte cells (adipocytes)
of the navel orangeworm, Amyelois transitella (Walker).
Bee larvae infected with sacbrood virus appear normal
until the final molt, when they fail to shed their last skin.
This becomes a transparent sac around the pupal integu¬
ment (Bailey 1973, fig. 22-1). Such larvae die in their
brood chambers. Likewise, the tetragonal virus of mos¬
quitoes and biting midges (Clark and O’Grady 1975)
causes a similar condition; the larval thorax wall becomes
an inflated sac surrounding the internal tissues (Kellen et
al. 1963, figs. 4 and 5). This virus, originally thought to
be a cytoplasmic polyhedrosis virus (Kellen et al. 1966),
was named for the large tetragonal masses of virus par¬
ticles concreted in the hypodermal and imaginal bud cells
(Kellen et al. 1963, fig. 3). Moribund larvae become slug¬
gish and abnormally curved (fig. 2G). The thoracic cuticle
develops hard, shiny black spots (fig. 2H). The infection
progresses slowly, killing larvae by the fourth instar. In¬
fected insects usually remain attached to the water sur¬
face after death.

Chronic bee paralysis virus (fig. 21) causes distension of
the abdomen, leg and wing trembling, failure to fly, and
leg paralysis; death occurs in about 1 week (reflecting the
nervous-tissue infection). (See also Bailey 1973, fig. 22-1,
and Gochnauer 1978 for further descriptions and photo¬
graphs of this condition). Other paralysis viruses (of
crickets, etc.) may cause general (lack of coordination) or
specific (rigid, extended hind legs) paralytic symptoms
that reflect infections of nervous tissues either alone or
with muscular-tissue involvement (see Scotti et al. 1980).
The host ranges of these other paralysis viruses are
usually fairly narrow; but a few viruses, like the cricket
paralysis group, can infect several hosts, even crossing
ordinal lines (Lepidoptera to Orthoptera).

Microscopic examination. — These small viruses occur na¬
ked in the cytoplasm of those certain tissues often in¬
volved in symptom expression. Their confirmation can be

done only by electron microscopic study of likely tissues
taken from living infected hosts. Representative photo¬
graphs of these viruses have been published by Smith
(1976) and by Maramorosch (1977).

Baculoviridae (nuclear polyhedrosis

viruses and granulosis viruses)

Only one described virus family, the Baculoviridae,
multiplies only in invertebrates. Within this family, the
genus Baculovirus (Fenner 1976) contains both the
nuclear polyhedrosis viruses (subgroup A) and the gran¬
ulosis viruses (subgroup B). The invasive virions of these
subgroups are embedded or occluded either individually
(in granulosis capsules) or in multiples (in polyhedrosis
polyhedra) in paracrystalline, protein-matrix bodies that
protect the virions when they are apart from host tissue.
The capsules of the granulosis viruses are usually ellip¬
soid; the polyhedra of the nuclear polyhedrosis viruses
vary from blunt-cornered or rounded polyhedral to the
rarer crescentic and fusiform, known from only two
viruses infecting certain dipterans.

Another subgroup, C (representing at least one additional
viral genus), has recently been added to accommodate the
nonembedded or nonoccluded baculoviruses. While the
subgroup A- and B-viruses of the genus Baculovirus are
generally lethal viruses having matrix bodies visible
under a light microscope, the subgroup C-viruses have a
broader range of host association and can be seen only
under the electron microscope.

Disease cycle.— Transmission occurs when polyhedra, cap¬
sules, or naked viruses (subgroup C only) from regur¬
gitate or anal discharge (sawflies) or from ruptured
cadavers (all host groups) are ingested by susceptible lar¬
vae; they then dissolve, releasing their virus particles.
These particles penetrate the midgut epithelium and
undergo one replication cycle in the epithelial cell nuclei.
Virus particles are rarely occluded in the few uncommon
polyhedra that are formed in these cells in the Lepi¬
doptera. Most progeny of the gut-cell cycle move out of
the gut cells into hemolymph, where they are transported
to the susceptible polyhedra-producing tissues. In the
Lepidoptera, polyhedra are produced in various combina¬
tions of several tissues, including the hypodermis, trache¬
al matrix, blood, and fat body but also to some extent the
glandular tissues and reproductive organs. In the
Diptera, there are fewer types of polyhedra-producing
tissues. In the Tipulidae, only the fat body produces
polyhedra. In the Culicidae, only the midgut epithelium
produces polyhedra (earlier reports of polyhedra in other
culicid tissues Involved a picomavirus). In the Hy-
menoptera (sawflies), there is the most restriction; poly¬
hedra are produced only in the midgut epithelium.

Among the granulosis viruses, the fat body is the major

103

capsule-producing tissue; in many lepidopteran hosts it is
the only capsule-producing tissue. But granulosis cap¬
sules are sometimes also formed in the tracheal-matrix
and hypodermal tissues. The complete Baculovirus
replication cycle was first delineated accurately by
Adams in 1975 when she discovered the nature of the
hemocoelic virus form (Adams et al. 1977).

Symptoms and signs.— Depending on the dosage and age
of the infected larvae, signs and symptoms may occur
sooner (in young larvae) or quite late (in older larvae). Or,
as in some situations with the slower viruses, there may
be no apparent signs or symptoms, and seemingly
healthy adults may be formed that lay virus-
contaminated eggs. In these cases, signs and symptoms
occur quite soon in the next generation, when many
young larvae die because they have ingested virus from
their own contaminated egg shells as they chewed their
way out. Usual signs and symptoms include a reduction
in feeding and a general sluggishness. The hemolymph
may become milky white rather than clear. If the hypo-
dermal cells become infected, the whole larva may change
color, becoming somewhat whitened or yellowed (figs.
3A-3C). In some cases, the abnormal white coloration
may be subtle (fig. 3D) and may be accompanied by some
localized swelling (fig. 3E). In some cases, an open dissection
may reveal presumptively infected tissues when com¬
pared to a normal insect (figs. 3F and 3G). Ear her infec¬
tions or granulosis infections (fig. 3H) may more closely
resemble the normal insect. In either case, the dissection
should be followed up with a microscopic study of the
presumptively infected tissues. With the sawfly
polyhedroses (restricted to the gut), larvae become
yellowish, their guts becoming an opaque milky white.
When disturbed, they produce a white regurgitate rather
than the normal clear yellow or greenish regurgitate.
Eventually, they exude a brown fluid from the anus, and
this glues them to the substrate where they turn brown
and finally black. Diseased sawfly larvae change their
behavior, becoming solitary rather than gregarious and
wandering randomly rather than together (Bailey 1973).
Infected individuals of some species climb to the highest
point available before dying. Moribund larvae become
flaccid and rupture easily. After death they often hang
head downwards by their posterior prolegs; the integu¬
ment becomes quite fragile as the contents liquify (fig.

31). Granulosis cadavers are less fragile than polyhedrosis
cadavers, particularly when only the fat body is produc¬
ing capsules. Granuloses may progress as fast as poly¬
hedroses; but, in general they are more prolonged, exten¬
ding from a few days to a month or more if death occurs
before pupation.

While the granuloses may be generally restricted to the
Lepidoptera (there is only one reported from a sawfly),
the nuclear polyhedroses have been reported from the

Coleoptera, Diptera, Hymenoptera, and Lepidoptera.
Nonoccluded (subgroup C) baculoviruses have been
reported as lethal agents in the Coleoptera (spreading
venereally between infected adults but lethal to larvae)
and in two acarine mites. Other possible members of
subgroup C apparently do not affect their hosts in the
Coleoptera, Homoptera, Hymenoptera, and Araneida. Cer¬
tain members of subgroup C, infecting the calyx epi¬
thelium of adult braconid and ichneumonid wasps (Stoltz
and Vinson 1979, Vinson et al. 1979) seem to be mu-
tualistic with their parasitoid hosts, either suppres¬
sing the encapsulation reaction of the lepidopteran host
or otherwise influencing the host response.

Within each virus taxonomic group, the experimentally
investigated host range may be quite narrow to fairly
broad. For example, the granuloses are generally re¬
stricted to a single host genus or a single family. In the
Lepidoptera, some polyhedroses are apparently as nar¬
rowly restricted as the granuloses, while others can easily
cross family lines. Nonlepidopteran polyhedroses seem to
be at least family restricted.

Microscopic examination. — Wet smears or Giemsa-stained
smears of infected tissue (fat body, tracheal matrix, hypo-
dermis, or midgut) should reveal 1-15 /um, blunt-cornered
or rounded polyhedra in the nuclei of infected cells (see
Steinhaus 1949, chapter 11, for wet-mount photographs
of polyhedra and granulosis capsules). These will be
Giemsa-stain negative (as are the cytoplasmic poly¬
hedrosis polyhedra). An acid pretreatment, however, (1.0
N HC1 for 10-20 minutes) will render nuclear
polyhedrosis polyhedra stainable (gray to blue-purple
with a 60-90 minute stain time); cytoplasmic poly¬
hedrosis polyhedra will remain clear under these condi¬
tions since the stain is more readily removed by the final
rinse in buffer (see the “Disease Recognition” section).
The gut-restricted, culicid nuclear polyhedrosis virus will
concrete fusiform polyhedra (which are confined to the
nuclei). Similar bodies are formed in the cytoplasm of
other tissues in some entomopoxvirus infections. Tipulid
polyhedrosis polyhedra are crescent-shaped. Granulosis
capsules are quite small, being just at the limit of resolu¬
tion of the light microscope (up to 0.5 by 0.2 urn). When
an infected cell is ruptured while being viewed in wet
mount, the tiny capsules will stream out in a dense cloud.

Poxviridae (Entomopoxvirus)

The genus Entomopoxvirus is apparently restricted to
the invertebrates (Fenner 1976); all isolates known so far
are from the Insecta. It has been divided into three
subgenera on the basis of virus morphology (and coin¬
cidentally by host range): subgenus A has unilateral con¬
cave virion cores and infects nine known species of
Scarabaeoidea; subgenus B has cylindrical-to-rectangular

104

virion cores and infects the Lepidoptera and Orthoptera;
and subgenus C has biconcave cushion-shaped (dumbbell-
shaped in cross section) virion cores and infects
chironomid and culicid Diptera (see Kurstak and Garzon
1977).

Disease cycle.— In the Lepidoptera and the Orthoptera,
the virus replication cycle usually takes about 10-30
days; but, in the Scarebaeid Coleoptera, where the host
life cycle may take 1-2 years, the virus replication cycle
may take 5-10 months or longer depending on the age of
the larva when infected. Since these viruses are largely
confined to the fat body, usually only ruptured cadavers,
cannibalism, or predators can furnish the matrix bodies
containing infectious viruses (or free viruses from the
hemocoel if parasitoids are involved), which usually ini¬
tiate the infection. In the Lepidoptera, however, mori¬
bund infected larvae often regurgitate or defecate fluid
containing virus matrix bodies. When ingested, these
bodies (here called spheroids) dissolve in the midgut, and
the viruses penetrate the midgut epithelial cell cy¬
toplasm. It is not yet known what mechanisms are in¬
volved in the passage of the entomopoxviruses through
the gut cell. The virus particles eventually reach the
target tissues, where final replication and occlusion (in
spheroidal matrix bodies) completes the replication cycle.
The cycle has been described by several authors, in¬
cluding Granados (1973), Kurstak and Garzon (1977), and
Vaughn (1974).

Symptoms and signs.— Infected insects exhibit a general
sluggishness, high mortality, and prolonged developmen¬
tal stages. In the Lepidoptera, death is often preceded by
paralysis of the gut and by regurgitation or defecation of
fluid containing infectious spheroids. In the Scarabaeoi-
dia (larvae having transparent cuticles), an externally
visible white-spotted or mottled appearance often de¬
velops in the dorsoposterior area of heavily infected indi¬
viduals; this appearance results from the enlargement of
infected fat-body tissue and possibly from infection of the
hypodermis (fig. 5 A; see also Godwin and Roberts 1975,
figs. 1-5). In some transparent dipteran larvae, the infec¬
tion of the fat body can be readily seen externally (fig.
5B), which greatly aids in distinguishing infections. In
the Orthoptera {Acrididae ), heavily infected insects be¬
come quite distended with obviously protruding cervical
membranes due to swelling of the spheroid-packed fat
body. Some lepidopterans also show this effect, becoming
larger and longer than normal larvae before succumbing
(fig. 5C). Seen in open dissection, a fat body infected with
Entomopoxvirus (fig. 5D) may be a flat chalky white or
grayish in infections lacking spindles. Where spindles are
present, the fat body takes on a reflective, foamy appear¬
ance (Goodwin and Roberts 1975, fig. 6). The entomopox¬
viruses are restricted to cross infectivity within the family
of origin so far as is known (Kurstak and Garzon 1977).

Of interest to insect culturists is a recent report describ¬
ing the density-dependent action of an entomopoxvirus in
a laboratory colony of chironomid midges (Harkrider and
Hall 1979).

Microscopic examination.— Of the several entomopox¬
viruses isolated from four insect orders, almost all pro¬
duce spheroidal virus matrix bodies only in the fat body
or in the fat body and hemocytes. Only two are more
general in the tissues attacked. One is restricted to the
hemocytes alone. The virus-containing spheroid is parallel
to the polyhedron of the nuclear and cytoplasmic polyhe-
drosis viruses; it is a paracrystalline protein matrix
dissolvable at certain high pH ranges with a combination
of dilute alkali plus a reducing compound such as dimer-
captopropanol, sodium thioglycolate, or cystein.
(Polyhedra from the other virus groups do not require the
addition of reducing agents for dissolution.) The size,
form, and distribution of the spheroids depend on the
virus species. They range in shape from nearly spherical
(coleopteran hosts) to nearly regular ellipsoidal (lepidop-
teran hosts). Some unusual spheroid shapes have also
been noted, as in the case of the Demodema bonariensis
Bruch virus (rounded subcubical) and the Melanoplus
sanguinipes (Fabricius) virus (ellipsoidal in saggital sec¬
tion but almost square in cross section). The Figulus
sublaevis Beauvois (Lucanidae), Chironomus luridus
Strencke, and Camptochironomus tentans (Fabricius)
virus spheroids may be more irregularly globose to round¬
ed polyhedral (Goodwin and Filshie 1975). So each virus
type usually has a strong degree of uniformity of
spheroid shape, while there is a wide range of shapes
among the different virus types or species.

The diagnostic utility of the spheroids (which are easily
overlooked in wet-mount preparations of lightly infected
insects— Moore and Milner 1973), is supplemented in
some cases by the presence of more distinctive virus-
related fusiform or spindle-shaped accessory or inclusion
bodies. The diseases of the Entomopoxvirus group were
in fact originally called the spindle viruses until it was
found that the spindles never contain the viruses and are
not always present. They are absent in the known
entomopoxviruses infecting the Diptera and the Or¬
thoptera but are present in some but not all of the
viruses infecting the Scarabaeoidea and the Lepidoptera.
Spindles range in size from the tiny microspindles, oc¬
cluded with virus particles in some lepidopteran
spheroids, to 25-/xm macrospindles that are larger than
many of the known spheroids. The pointed spindles are
readily distinguishable by shape in wet-mount smears
when viewed with phase optics (Goodwin and Filshie
1969). They stain readily with Giemsa (usually light to
dark blue); the mature spheroids remain unstained. Partly
formed “immature” spheroids of some of these viruses
stain gray or gray-brown with Giemsa alone (Goodwin

105

and Roberts 1975). Prolonged (1 hour or more) Giemsa
staining of certain unresponsive types [Aphodius tas-
maniae Hope virus) results in a green staining reaction of
both spindles and spheroids. But, if this long Giemsa
staining is preceded by a 2-hour acid hydrolysis treat¬
ment with IN HC1, the spindles will stain a pale gray-
brown; the spheroids will stain medium to dark blue and
reveal inner reddish spots that may relate to the often-
seen retractile fissures or the contained virus particles
(Goodwin and Roberts 1975).

A diagnostic stain for the differentiation of both Entom-
opox virus spheroids and spindles was described by Moore
and Milner (1973). It involves the use of concentrated lac-
tophenol cotton blue for staining fresh wet smears; the
rapid result is that both spheroids and spindles are col¬
ored intensely blue, which distinguishes them from fat
globules, uric acid crystals, and other types of occluded
viruses. To prepare this stain (which is also used as a
mounting medium and stain for fungi), combine the fol¬
lowing: phenol crystals (100 g), lactic acid (USP 85%; 80
ml), glycerol (159 ml), and distilled water (100 ml); add
0.5% cotton blue (anilin blue, C.I. 42755, or methyl blue).
This preparation will keep at room temperature indefinite¬
ly (Poinar and Thomas 1978). Add the stain directly to
the dried smear and cap with a cover slip. The stain is
also a mounting medium and will intensify with time.

Rickettsiales and Chlamydiales (see p. 117)

The rickettsiae are a group of lower micro-organisms that
includes both obligate parasites and commensals of ar¬
thropods, though the group has some features common to
the bacteria. Rickettsiae have a cellular structure in¬
cluding a cell wall, they contain both RNA and DNA, and
they contain active metabolic enzyme systems. The insect
pathogens in the rickettsial tribe Wolbachieae were
thought to be restricted to arthropods, but some have
more recently been shown to be infectious to warm¬
blooded vertebrates. Of the four known genera in the
Wolbachieae, one, the intracellularly occurring Wolbachia,
contains types that show no particular tissue tropisms
and that may be transmitted through the eggs from in¬
fected females. These have been isolated from mos¬
quitoes, lice, ticks, and mites. It is possible that some
rickettsiae in this genus may be symbiotes. A second
genus, Eickettsoides, grows (epicellularly) on the gut
epithelium of the host (some lice, fleas, and parasitic
flies), but they are apparently harmless to their hosts.

Two genera, Enterella and Rickettsiella, contain species
pathogenic to insects (see Krieg 1963 and 1971b).

A series of chlamydiae showing affinities to both Enter-
ella and Rickettsiella infections, but that manifest them¬
selves only as stunting agents for a wide variety of hosts,
has been found by J. R. Adams (unpublished data). These

chlamydiae have been isolated from hypodermis tissue
(sometimes causing black spotting of the integument) and
occasionally also from the midgut. One chlamydia can
also be transovarially transmitted in Trichoplusia ni
(Hiibner)— embryonal hypodermal cells contain
chlamydiae. Laboratory-reared species found infected
include the German cockroach, Blattella germanica
(Linnaeus), and the following lepidopterans: the range
caterpillar, Hemileuca oliviae Cockerell; the redbacked
cutworm, Euxoa orchogaster (Guenee); the southwestern
com borer, Diatraea grandiosella (Dyar); the tobacco bud-
worm, Heliothis virescens (Fabricius); the bollworm,
Heliothis zea (Boddie); the cabbage looper, Trichoplusia
ni (Hiibner); the European corn borer, Ostrinia nubilalis
(Hiibner); the almond moth, Ephestia cautella (Walker);
the pink bollworm; the saltmarsh caterpillar, Estigmene
acrea (Drury); and the tobacco homworm, Manduca sexta
(Linnaeus). Chlamydiae were also found in nerve tissue of
infected field-collected species including the satin moth,
Leucoma salicis (Linnaeus); the melon aphid, Aphis
gossypii Glover; the cereal leaf beetle, Oulema melanopus
(Linnaeus); the beetle Coccinella septumpunctata (Lin¬
naeus); the sugarbeet root maggot, Tetanops myopaefor-
mis (Roder); and the alfalfa weevil parasite, Bathyplectes
sp.

Enterella

Disease cycle. — The rickettsiae of this genus are usually
associated with the host gut epithelium, but they grow
intracellularly rather than epicellularly. This genus has
few described members. Some are apparently insignifi¬
cant in their pathogenic effects on hosts. Others are
lethal through their destruction of the midgut epithelium
(as, for example, Enterella stethorae ). Transmission is
through ingestion. The rickettsiae are probably spread
through regurgitate and fecal contamination of the en¬
vironment and by rupture of cadavers. Such rickettsiae
are known so far only from the coccinellid coleopteran
Stethorus and the culicid dipterans Culex, Anopheles, and
Aedes and from one lepidopteran.

Symptoms and signs.— The Enterella isolated from
Hyalophora cecropia (Linnaeus) and Samia cynthia
(Drury) (Lepidoptera : Satumiidae) causes a lethal,
perhaps toxic, dysentery. Infected larvae show a de¬
creased rate of growth and eventually stop feeding entire¬
ly. The body weight decreases 1-2 weeks before death.
Diarrhea precedes regurgitation, and both symptoms are
accompanied by violent muscular contractions of the
body. Moribund larvae are generally flaccid, but some die
in a contracted position. The course of the disease is
about 14 days in larvae (Entwistle and Robertson 1968).

Microscopic examination.— The lepidopteran Enterella
described above was found primarily in the midgut but

106

was also isolated from the fat body and the hypodermis
of infected insects. Normal staining procedures were un¬
satisfactory in revealing the rickettsiae; but, if smears or
sections were pretreated for 3-5 minutes with IN HC1 at
60° C and then stained overnight in dilute Giemsa solu¬
tion (pH 7.5-7. 8), results were excellent. The cell cyto¬
plasm stained light blue, the nuclei, dense violet, and the
intracellular rickettsial organisms and gut bacteria, red.
Both Enterella and Rickettsiella are often visible in
chains (Entwistle and Robertson 1968), which distin¬
guishes them from granulosis virus capsules that are also
quite tiny but never occur in chains.

Rickettsiella

Disease cycle.— Rickettsiella are known to survive for
more than 3 years in the soil. They also show some
resistance to heat, chemicals, antibiotics, and radiation
(Krieg 1971b). They are spread through ingestion from
ruptured cadavers and cannibalism. Long-lived coleop-
teran larvae of the Scarabaeidae show signs or symptoms
after about 2-3 months, but they may not die until 6
months after infection (at 20° C). The rickettsiae
penetrate the midgut epithelium and chiefly infect the fat
body, but there may be some involvement of the ovaries,
ganglia, tracheal matrix, malpighian tubules, muscula¬
ture, and hypodermis, depending on the rickettsial
species.

Symptoms and signs. — Infected larvae become sluggish
as the disease develops, probably due to the involvement
of nervous tissue later in the infection. Acrididae, Blatti-
dae (fig. 4A), Gryllidae, Tenebrionidae, and Carabidae (fig.
4B) infected by Rickettsiella often show a swelling of the
abdomen as the fat body, packed with rickettsiae, be¬
comes enlarged within. This abdominal swelling is usually
apparent because of the exposure of the intersegmental
membranes that are normally folded beneath the segmen¬
tal scelerites. Discolorations of infected larvae are com¬
mon in these infections, particularly among hosts with
transparent cuticles. Infected tipulids and chironomids
become abnormally whitened (as in fig. 1A) as the pro¬
liferating rickettsiae are released from the fat-body cells
and blood cells into the hemolymph. A similar chalky
whiteness due to the same cause is apparent in certain in¬
fected scarabaeid larvae ( Melolontha spp., Europe; Ano-
plognathus spp., Australia; and Costelytra spp. and Odon-
tria spp., New Zealand). Rickettsiella popilliae will color
infected Japanese beetle, Popillia japonica Newman, lar¬
vae bluish ventrally so that externally it resembles an
Iridovirus infection. I have observed several Australian
Rickettsiella infections in scarabaeid larvae and am repor¬
ting them here for the first time: (1) an infection in larvae
of the blackheaded pasture cockchafer, Aphodius
tasmaniae Hope, that causes a scattered black spotting
of the integument probably caused by hypodermal cell

death (fig. 4C); (2) an infection in larvae of the tableland
pasture scarab, Antitrogus (formerly Rhopaea) mor-
billosus (Blackburn), that colors the legs, head, anal area,
and spiracles a deep blue black (figs. 4D-4F); (3) an infec¬
tion in larvae of the cocksfoot grub, Rhopaea verreauxi
Blanchard, resulting in a green and brown mottling of the
entire integumental surface and a progressive darkening
as the insects become moribund; and (4) an infection in
several species of melolonthine larvae in which lobes of
infected fat body swell due to rickettsial growth within
and then break free and float around in the hemolymph
(figs. 4G-4H). In all these infections, typical rickettsiae
were isolated from the fat body, stained with Giemsa as
smears, and then morphologically confirmed as rickett¬
siae by electron microscopy. All infected larvae showed a
progressive sluggishness but no other notable symptoms.

The host range of the known Rickettsiella species is wide
among those few that have been so investigated (Krieg
1971b). Rickettsiella are widely infectious between host
families and perhaps also between host orders. Rickett¬
siae may take a long time to kill their hosts, and infected
hosts may not have very obvious symptoms (these may
be similar to the stunting syndrome in Enterella ).

Because of these factors and since they are quite resist¬
ant to destruction in the field apart from their hosts,
Rickettsiella species may be expected to cause con¬
siderable losses before being discovered in insectaries
that use frequently gathered field stock (fig. 41).

Microscopic examination.— The Rickettsiella are stainable
with Giemsa in smears as are other rickettsiae. Although
the typical chains of mature rickettsiae (also typical of
mature Enterella ) may not be seen until quite late in the
infection, there are more obvious earlier microscopic signs
of this genus. Variously sized intracellular rickettsiae-
filled vacuoles (called RFV or NR bodies in the earlier lit¬
erature) retain their structure and stain violet to deep
purple with Giemsa in fat-body squash or smear prepara¬
tions (see Krieg 1963, fig. 4). Eventually, these vacuoles
release mature rickettsial progeny. Usually, variously
shaped, large, stain-negative crystals are also associated
(inside and outside) with the rickettsiae-filled vacuoles in
such smears. The rickettsiae-filled vacuoles are more use¬
ful as presumptive diagnostic indicators of Rickettsiella
than the rickettsiae themselves since they are more readi¬
ly stained and are present earlier and over a longer period
during such infections.

Bacteria

The bacteria are minute (0. 2-5.0 yum), unicellular plantlike
organisms that differ from higher plants in that they lack
chlorophyll and do not contain organelles. They are classi¬
fied by shape into four main groups: rod-shaped bacilli,
spherical cocci, comma-shaped spirilla, and branched or-

107

ganisms of the actinomycetes. Many are motile by means
of flagella.

Insect relationships of bacteria

Bacteria are widely distributed throughout the environ¬
ment and so, like the fungi, are common contaminants of
artificial insect diets and insectary stock insects. In the
insectary environment, and particularly where artificial
diets are used, several bacterial genera may colonize the
diet and then the guts of stock insects, resulting in
continuous stock losses (McLaughlin and Sikorowski
1978) that may be attributed to other causes. Some of the
bacterial genera and species implicated in such insectary
mortality have been listed among the normal microflora
of a number of insects, particularly Streptococcus spp.
(Jarosz 1979), but also Aerobacter ( Enterobacter ) spp.
(Nunez et al. 1968), Escherichia spp., Erwinia spp., Pro¬
teus spp., Micrococcus spp., Alcalignees spp., and
Flavobacterium spp. (Pristavko 1966). Many in¬
vestigators have listed these and other bacteria “from
diseased and dead larvae” and demonstrated larval mor¬
tality by feeding them to laboratory insects (see several
studies cited by Lipa and Wiland 1972). It is apparent
from these and other conflicting studies that many of the
listed “microflora” are only fortuitous contaminants.
Those that cause diseases in insectaries reflect as much
the ability of some bacteria to outcompete others as the
ability of some to actively invade insects.

There are many complex interactions between bacteria
and other microbes as well as between insects and
bacteria (Brooks 1963). Some “established” symbiotic
relationships, such as those occurring in fruit flies,
especially those reported by Boush and Coppel (1974) for
the olive fruit fly, Dacus oleae (Gmelin), have come into
question when studied more closely (Yamvrias et al.

1970). Recent work with the walnut husk fly, Rhagoletis
completa Cresson, has demonstrated a loose bacterial
symbiotic relationship that may vary between fly strains
or species and that may change as the fly diet changes,
either in the field or in the insectary (Tsiropoulos 1976).
The fact that D. oleae strains adapted to artificial diets
cannot survive on olives (Fytizas and Mazomenos 1971)
may indicate either a genetically deficient fly strain or
the loss of a very specific and morphologically distinctive
bacterial symbiote that has not yet been cultured apart
from the fly (Poinar et al. 1975).

Although some bacterial genera (Serratia marcescens,
Pseudomonas spp., and others) have repeatedly caused
field epizootics in sawflies and are recorded as true in¬
vasive pathogens in certain insects (Grimont et al. 1979a,
Poinar et al. 1979), such bacteria are more often
classifiable in other insect hosts as noninvasive
facultative pathogens. To cause disease, these depend on

factors such as high humidity or temperature stress
(Goodwin 1968, Greany et al. 1977, Habib 1978), wound¬
ing, or tearing of the hindgut or foregut during molting
or pupation (Goodwin 1968). Some bacteria related to the
facultative pathogens— for example, Serratia ficaria in the
fig wasp, Blastophaga psenes (Linnaeus), and others
(Grimont et al. 1979a, 1979b)— may be either insect com¬
mensals or loosely associated symbiotes. For example,
Streptococcus spp., while apparently beneficial to some
insect species (Jarosz 1979), may be primary invasive
pathogens in others (Doane 1971); in this case, the spe¬
cies Streptococcus faecalis was responsible for both
effects.

Management of insectary bacteria

Depending on the insect species, specific strains of S.
faecalis may be considered for use in prophylactic gut
floration, perhaps in association with other bacterial
species (Goodwin 1968) such as the Erwinias and other
plant-rotting bacteria, particularly in the insectary rear¬
ing of phytophagous insects (Martin and Mundt 1972).
Also, such action may be simpler and preferable to the
maintenance of aseptic rearing conditions in preventing
the activities of the facultatively pathogenic bacteria.
While germicide additions to artificial insect diets are
now commonplace, the use of antibiotics is to be
discouraged in general. Such use encourages further un¬
necessary selection of resistant strains of bacteria and
often merely masks an unfavorable insectary environmen¬
tal condition that should be corrected. Instead, further
study should be given to the addition of leaf extracts
from usual host plants (Goodwin 1968) that may contain
bacteriostatic or bactericidal flavonoids or terpenes. Such
extracts may contain factors that interact with the insect
gut to produce protective compounds, such as active caf-
feic acid, which has been isolated from the guts of leaf-
fed silkworms; caffeic acid has been shown to protect
silkworms against pathogenic S. faecalis isolates and
other bacteria (Koike et al. 1979). Again, more naturally
microbe-florated gut microenvironments may be related
to improved survival and competitiveness in colonized
parasitoids or genetically altered stock that is to be used
for field release.

Symptoms and signs

Since some bacteria will be present even within insects
reared in aseptic or clean environments, they must be ex¬
pected in decomposed insects, whatever the cause of
death. Therefore, careful evaluation must follow the
detection and isolation of bacteria from insectary stock.
Dead insects killed by facultative bacterial pathogens in
the insectary often reflect transient unfavorable condi¬
tions such as adverse humidity and/or temperature. More¬
over, large numbers of insects, even among those reared

108

individually but in the same location, will often contain
similar bacteria, these being local ambient air or surface
contaminants. Insects infected by S. marcescens are
usually pinkish to red when infected by pigmented
strains; but many nonpigmented strains exist that are
also pathogenic (Grimont et al. 1979a). Cadavers of in¬
sects infected with Pseudomonas spp. may appear green¬
ish and exude a heavy, sweet odor. Other bacteria also
give off characteristic odors when present in cadavers in
near monocultures.

Insects infected with bacteria become progressively more
sluggish before death, but other more definitive symp¬
toms rarely occur. At death or shortly thereafter, the
usually septicemic growth of bacteria in the hemocoel will
have largely decomposed most of the internal tissues.
Further discussion of symptoms and signs of the spore¬
forming and nonsporeforming primary bacterial
pathogens can be found in Dutky (1963, milky diseases),
Bucher (1963, nonsporeformers), Heimpel and Angus
(1963, sporeformers), Faust (1974, bacterial diseases,
general), and Shimanuki (1978, bacterial diseases of bees).

Microscopic examination

A wet-mount smear may reveal a uniform type of bac¬
teria and its motility, if it is present as a septicemic
agent in the hemocoel. If a Giemsa smear from the in¬
fected insect confirms that only bacteria of a relatively or
completely uniform morphology are present (rods of a cer¬
tain size, cocci, or spirillae), then a Gram stain (Poinar
and Thomas 1978, pp. 187-188) may confirm the presence
of a uniform type (Gram positive or negative). If further
identification of the bacterium is warranted, it may be
isolated by use of the techniques described by Poinar and
Thomas (1978, pp. 60-77 and 167-180). Precise identifica¬
tions can be obtained from contract laboratories. If mixed
types are present in the cadavers, one should attempt to
isolate bacteria either from the moribund insects or from

those that are living but diseased so as to segregate
possible pathogens from the many saprophytic forms
that usually luxuriate in cadavers.

Fungi

The fungi are heterotrophic micro-organisms with chi-
tinized cell walls; they are typically nonmotile, though
motile stages (zoospores) may be present. Most of the en-
tomogenous fungi contain hyphae (fungal hairlike strands
developing from a germinating spore) that, grouped to¬
gether, constitute a mycelium (fungal mat or mass).
Reproduction is mainly by sexual or asexual spores.
Asexual spores are borne in taxonomically characteristic
sporangia (as sporangiospores) or on characteristically
shaped special hyphae or conidiophores (as conidiospores).

Many fungi, like bacteria, are opportunists; that is they
are facultative parasites or saprophytes rather than pri¬
mary pathogens. Often in the insectary, saprophytic
fungi will be found growing on the surface of dead insects
as well as on the food and frass in rearing chambers.

Such fungi usually do not solidify, or mummify, host in¬
sects as many of the entomogenous pathogens do. Ento-
mogenous fungi occur among four classes: the
Zygomycetes, including Entomophthora, Massospora, and
others; the Chytridiomycetes, including Coelomomyces; *
the Ascomycetes (sacfungi), including Ascosphaera, Cor-
dyceps, and others; the Basidiomycetes (club fungi), in¬
cluding Septobasidium; and the Deuteromycetes (Fungi
imprefecti, sexual stages unknown), including Asper¬
gillus, Beauveria, Hirsutella, Isaria, Metarrhizium,
Nomuraea ( Spicaria ), Paecilomyces, and others. For a
more complete generic listing, see Roberts and Yendol
(1971, table 1); and, for the associations between genera
of fungi and insect host taxons (including references), see
the listing by Bell (1974, table 1) and key 2, which
follows.

109

Key 2. — Key to common insect fungi:

1.

1'.

2(1).

2(1).

3(1').

3 ' (1 ' ).
4(3').

4(3).

5(4).

5(4).

6(5').

6 ' (5 ' ).
7(6').

7(6).

8(4').

8'(4').

9(8).

9(8).

10(9).

10 '(9).

11(9').
11 (9).
12(11').

12(11 ').

Thallus attached to chitinous gut lining or exoskeleton by a secreted holdfast or a specialized

holdfast cell . 2

Thallus without a holdfast, intercellular or intracellular . 3

Thallus mycelial, not organized into a sporocarp, spores one-celled, mostly commensals or

symbionts of aquatic Diptera, Ephemeroptera, and certain Coleoptera . Trichomycetes

(Lichtwardt 1973, illus.,4 keys;5 Moss 1979, illus., descrip.6)

Thallus reduced, forming a cellular, perithecioid ascocarp; often hairlike or setose; ascospores

two-celled; obligate ectoparasites . Laboulbeniomycetes

(fig. 6A; Benjamin 1973, illus., keys; Tavares 1979, illus., keys)

Hyphae lacking or scarce, septate, reproduction by yeastlike cells .

. Candida and related genera

(Lodder 1970, illus., keys, descrip.)

Hyphae abundant, septate or unseptate . 4

Mycelium unseptate (septa formed only during reproduction) and fragmenting into

uninucleate or multinucleate hyphal bodies (Mastigomycota) . 5

Mycelium septate throughout the life cycle (Ascomycotina and Deuteromycotina) . 8

Hosts aquatic, usually in larvae of Culicidae; zoospores present . Coelomomyces Keilin

(Couch 1945, illus., keys, descrip.; McNitt and Couch 1977; Whisler 1979; Burges 1981,
illus., descrip., see keys to Coelomomyces and Lagenidiales)

Hosts terrestrial, zoospores absent . 6

Spores produced outside the host, often between intersegmental folds .

. Entomophthora Fresenius

(fig. 6B; Macleod and Mviller-Kogler 1973, illus., keys, descrip.; Macleod et al. 1976, illus.,
keys, descrip.; Waterhouse 1973)

Spores produced in the host . 7

Spores aggregated on a palisade of conidiophores and discharged through an abdominal hole

on muscoid diptera . Strongwellsea Batko & Weiser

(Humber 1976, illus., keys, descrip.)

Spores surrounded by hyphae and released on disintegration of the host abdomen, restricted
to Cicadidae . Massospora Peck

(Soper 1974, illus., keys, descrip.)

Spores within asci (ascospores) formed in perithecia (subclass Ascomycotina) . 9

Spores on conidiophores that are solitary, in pycnidia, acervuli or on coremia (subclass

Deuteromycotina) . 13

Asci scattered throughout the ascocarp . 10

Asci arranged in a hymenium . 11

Ascocarps formed within a stroma, ascospores septate, usually restricted to scale insects . . .

. Myriangium Montagne & Berkeley

(Miller 1940, illus., keys, descrip.; von Arx 1963)

Ascocarps (erronously referred to as cysts by some) formed superficially on a white, cottony

mycelium; ascospores unseptate and grouped as spore balls; on Apoidea .

. Ascosphaera Olive & Spiltoir

(fig. 6C; Gilliam 1978; Skou 1972, illus., descrip.)

Ascospores two- to five-celled, ellipsoid or fusoid . Nectria (Fries) Fries

Ascospores many -celled, filiform or elongate fusiform . 12

Hosts mummified; perithecia borne within a stalked, light-colored, stromatic head;

ascospores fragmenting at septa . Cordyceps Link

(Mains 1958, illus., keys, descrip.; McEwen 1963)

Hosts not mummified, perithecia borne on a superficial, effused-to-pulvinate, dark-colored

stroma; ascospores not fragmenting . Hypocrella Saccardo and Podonectria Peck

(Petch 1921)

3By L. R. Batra, Mycology Laboratory, Plant Protection Institute, Agricultural Research Service, U.S.
Department of Agriculture, Beltsville, Md. 20705.

‘illus.— The reference has illustrations of the taxa below the rank discussed here.

“keys— The reference has keys to the taxa below the rank discussed here.

6descrip.— The reference has descriptions of the taxa below the rank discussed here.

110

13(8'). Conidiophores in flask-shaped pycnidia (Sphaeropsidales), pycnidia immersed in bright-
colored stromata, conidia one-celled, hyaline, sexual stages in the genera Hypocrella and

Stereocrea H. & P. Sydow . Aschersonia Montagne

(Mains 1959)

13 ' (8 ' ). Conidiophores not as above . 14

14(13'). Conidia of two kinds: (1) Macroconidia— slightly bowed and often with chlamydospores; (2)

Microconidia— one-celled . Fusarium Link

(fig. 6F; Booth 1971, illus., keys, descrip.; Madelin 1963)

14 '(13'). Conidia of one kind, neither bent nor septate . 15

15(14 ' ). Sporogenous cells formed on globose-to-clavate heads or swellings of the conidiophore . . . . 16

15 '(14'). Sporogenous cells not formed on a swelling of the conidiophore . 17

16(15). Coremia absent, conidia globose to subglobose, in definite chains .

. Aspergillus Micheli ex Fries

(fig. 6E; Madelin 1966; Raper and Fennell 1965, illus., keys, descrip.)

16 '(15). Coremia usually present, conidia fusoid to ellipsoid, solitary or in chains, confined to spiders

(ascigerous stage Torrubiella ) . Gibe llula Cavara

(Mains 1950a)

17(15 ' ). Sporogenous cells formed on synnemata . 18

(figs. 6L, 6M)

17 '(15'). Sporogenous cell usually formed on solitary hyphae . 23

18(17). Sporogenous cells arranged in a hymenium . 19

18 ' (17). Sporogenous cells usually not forming a hymenium . 20

19(18). Sporogenous cells obtuse, conidia solitary and without mucous . Hymenostilbe Petch

(fig. 6H; Mains 1950b, illus., descrip.)

19' (18). Sporogenous cells usually pointed, sometimes sharply so; conidia catenulate, with or without

mucous . Insecticola Mains and Akanthomyces Lebert

(figs. 61 and 6 J ; Mains 1960b, illus., descrip.)

20(18 ' ). Conidiogenous cells not elongated, conidia dry . Isaria Persoon ex Fries

(DeHoog 1972, illus., keys, descrip.)

20 ' (18 ' ). Conidiogenous cells elongated and usually tapering to a point, conidia mucilaginous . 21

21(20 ' ). Conidiogenous cells usually enlarged at the base, conidia alone or in groups of two or more in

droplets . Hirsutella Patouillard

(fig. 6K; Mains 1951, illus., descrip.)

21 ' (20 ' ). Conidiogenous cells not enlarged at the base, conidia held in a conspicuous mucilaginous ball

. 22

22(21 '). Sclerotia present, spore balls not disintegrating readily . Synnematium Speare

(fig. 6M; Mains 1951, illus., descrip.)

22 ' (21 ' ). Sclerotia absent, spore balls disintegrating readily . Tilachlidium Preuss

(fig. 6L; Mains 1951, illus., descrip.)

23(17 ' ). Conidia aggregated in slime at the tips of conidiophores .

. Acrostalagmus Corda and Cephalosporium Corda

(fig. 6N; Ganhao 1956)

23 '(17'). Conidia dry or powdery . 24

24(23 ' ). Conidia borne in chains . 25

24 ' (23 ' ). Conidia borne singly . 28

25(24). Hosts mummified; conidiophores formed in an olive-green velvety mound; conidia basipetal,

cylindrical . Metarrhizium Sorokin

(fig. 6P; Gams and Rozsypal 1973, illus., descrip.)

25 ' (24). Host usually not mummified, conidia ellipsoid or globose . 26

26(25 ' ). Conidiogenous cells clustered on penicillate, nondivergent conidiophores Penicillium Link

(fig. 6G; Raper and Thom, 1949, illus., keys, descrip.)

26 ' (25 ' ). Conidiogenous cells clustered or solitary along divergent, often verticillate conidiophores . 27
27(26 ' ). Conidia pale green or light purple (green or purple in mass), broad— ellipsoid or cylindrical . . .

. Nomuraea Maublanc

(fig. 6D; Getzin 1961; Sampson 1974, illus., descrip.)

27 ' (26 ' ). Conidia hyaline, ellipsoid to fusiform . Paecilomyces Bainer

(For P. farinosus Gray, an entomogenous sp., see Sampson 1974)

28(24'). Conidia borne on minute sterigmata, conidiogenous area often zigzag . Beauveria Vuilleman
(fig. 60; DeHoog 1972, illus., keys, descrip.)

28 ' (24 ' ). Conidia borne on depressions on the conidiophores . Sporotrichum Link ex Fries

(see Petch 1938 for entomogenous species)

Disease cycle

Most entomogenous fungi initiate infection with a ger¬
minating spore (conidium) that adheres to and penetrates
the cuticle of the insect. The invasive hypha grows,
enters the host tissues, and ramifies through the hemo-
coel. As a result, the fungus becomes distributed
throughout the host’s hemocoel, filling it with hyphae
(mycelial mass). After death, the solidified host may dry
and become hard (mummification). On incubation of the
mummified cadaver in a moist environment, emergence
hyphae grow out through the insect’s integument and
produce spores (usually conidia) on the external surface of
the host (see Steinhaus 1949, chapter 10; Weiser 1969,
figs. 180, 181, 190C, 203, 213, 216-219, 222, 242-250 and
Poinar and Thomas 1978, figs. 3-41). These spores are
dispersed by forcible ejection (as in fig. 5E) or by wind to
contact further hosts. In certain aquatic forms (Coelomo-
myces spp.), rupture of the cadaver releases asexual spor¬
angia (Weiser 1969, figs. 159-166) that later release
zoospores infectious to alternate hosts (Copepoda). A
similar (sexual) cycle occurs in the alternate host, and the
zoospores produced there are infectious to the original
mosquito or other dipteran host species (Whisler 1979).
Both Coeiomomyces and similar aquatic fungi in the
Lagenidiales and Saprolegniales were described more re¬
cently by Bland, Couch, and Newell in chapter 8 of
Burges (1981). The general fungus disease cycle is des¬
cribed in more detail by Roberts and Yendol (1971) and
by MacLeod (1963), Madelin (1963), and Bell (1974), who
also discuss the environmental conditions favoring fun¬
gus infections.

Symptoms and signs

Larvae may show black spots at the sites of spore
penetration as an early disease sign. There may be nerv¬
ous activity and restlessness followed by sluggishness
and a cessation of feeding before death. Often, color
changes (to brown, fig. 5F, or to pink, red, purple, or
yellow, fig. 5G) are apparent before or after death. When
the emergence hyphae grow out of the solidified cadaver
and form conidia, the resulting macroscopic appearance
may allow a presumptive diagnosis (figs. 5E, 5F, 51). Cor-
dyceps infections are particularly characteristic (McEwen
1963) but are unlikely to occur in insectaries. Various
fungus infections that are characteristic in appearance
have been demonstrated in illustrations by Steinhaus
(1949), Weiser (1969), and Poinar and Thomas (1978).

More extensively described symptoms and signs appear
in Steinhaus (1949), Couch and Umphlett (1963), McEwen
(1963), MacLeod (1963), Madelin (1963), Bell (1974), and
Lipa (1975). Fungi in bees are described by Gilliam
(1978). Earlier generic keys to some entomogenous fungi
were presented by Weiser and Briggs (1971) and Poinar
and Thomas (1978). More recent keys to the Deuteromy-

cetes; Entomophthorales; and aquatic fungi in the
Lagenidiales, Saprolegniales, and Coeiomomyces are
given in Burges (1981).

Microscopic examination

Incubation of the suspect solidified cadaver (terrestrial,
not aquatic, hosts) in a moist chamber, will cause out¬
growth of the characteristic conidiophores or spore-
bearing emergence hyphae (figs. 5G-5I). These can be
pulled off with forceps and mounted on a slide with
Guegen’s solution (Poinar and Thomas 1978) or lacto-
phenol cotton blue (see above in the section on micro¬
scopic examination of Entomopoxvirus ) to render the
characteristic morphology more easily visible. Then, us¬
ing key 2 and the microscopic illustrations in figure 6 (as
well as the photographs presented in Weiser 1969, Poinar
and Thomas 1978, and Burges 1981), one can make a
presumptive generic identification. Photographs or draw¬
ings of unusual morphological types and cultures pro¬
vided on suitable agars (Poinar and Thomas 1978) would
aid specialists in determining unusual types accurately if
a species identification is necessary.

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Figures 1-6 follow on pages 118-129.

Note Added in Proof

A recent serological analysis by J. R. Adams has placed the stunting “chlamidiae”
described here closer to the Rickettsiales than to the Chlamydiales. Further work
may place them in a new rickettsial tribe or other subgrouping.

117

118

Figure 1

A. Chironomid larvae. Fat body infected with a microsporidian, Telomyxa sp. See fig. 5B
for appearance of normal larva. (Photograph courtesy of the Gulf Coast Mosquito Re¬
search Unit, U.S. Agricultural Research Service, Lake Charles, La.)

B. Anopheles sp. (Culicidae) larvae. Fat body infected with a microsporidian, Nosema sp.;
arrows indicate swollen, diseased abdominal segments. (Photograph courtesy of the Gulf
Coast Mosquito Research Unit, U.S. Agricultural Research Service, Lake Charles, La.)

C. Aedes sollicitans (Culicidae) larvae. Left larva infected with a microsporidian, Thelo-
hania sp.; right larva normal. (Photograph courtesy of the Gulf Coast Mosquito Research
Unit, U.S. Agricultural Research Service, Lake Charles, La.)

D. Culiseta incidens (Culicidae) larvae. Left larva, with oenocytes infected by a microspor¬
idian, Thelohania sp.; right larva normal. (Photograph courtesy of W. R. Kellen, Stored-
Product Insects Research Laboratory, U.S. Agricultural Research Service, Fresno, Calif.)

E. Amyelois transitella (Pyralidae) larvae infected with the microsporidian Nosema in-
vadens, showing characteristic black spotting of the integument. (Photograph courtesy of
W. R. Kellen, Stored-Product Insects Research Laboratory, U.S. Agricultural Research
Service, Fresno, Calif.)

F. Amyelois transitella (Pyralidae) larvae. Normal-sized larvae above; smaller larvae below
stunted by the microsporidian Nosema invadens. (Photograph courtesy of W. R. Kellen,
Stored-Product Insects Research Laboratory, U.S. Agricultural Research Service, Fresno,
Calif.)

G. Amyelois transitella (Pyralidae) pupae, normal. (Photograph courtesy of W. R. Kellen,
Stored-Product Insects Research Laboratory, U.S. Agricultural Research Service, Fresno,
Calif.)

H. Amyelois transitella (Pyralidae) pupae, malformed by the microsporidian Pleistophora
sp. (Photograph courtesy of W. R. Kellen, Stored-Product Insects Research Laboratory,
U.S. Agricultural Research Service, Fresno, Calif.)

I. Sericesthis geminata (Scarabaeidae) larvae. Normal larva below; larva above infected
with the coccidian Adelina sericesthis early stages, causing translucent clearing of dorsal
tissues indicated by arrow.

119

120

Figure 2.

A. Sericesthis geminata (Scarabaeidae) larva showing the characteristic brown mottling
caused by oocyst formation in a late infection with the coccidian Adelina sericesthis.

B. Rhopaea verreauxi (Scarabaeidae) larva containing hemocoelic cysts of a gregarine
Stictospora sp.; external view.

C. Rhopaea verreauxi (Scarabaeidae) larva containing hemocoelic cysts of a gregarine
Stictospora sp.; open dissection.

D. Culex salinarius (Culicidae) larva. External view by transmitted light showing a
cytoplasmic polyhedrosis virus (Reovirus) infection of the midgut. Arrows denote swollen
midgut portions containing polyhedra. (Photograph courtesy of T. B. Clark, Insect
Pathology Laboratory, U.S. Agricultural Research Service, Beltsville, Md.)

E. Aedes sollicitans (Culicidae) larvae. Left insect with a cytoplasmic polyhedrosis virus
(Reovirus) infection of the midgut; right insect with a nuclear polyhedrosis virus
(Baculovirus) infection of the midgut. (Photograph courtesy of T. B. Clark, Insect
Pathology Laboratory, U.S. Agricultural Research Service, Beltsville, Md.)

F. Amyelois transitella (Pyralidae) larvae. Top two larvae normal; small larvae below
showing stunting due to infection with chronic stunt virus (a picornavirus). (Photograph
courtesy of W. R. Kellen, Stored-Product Insects Research Laboratory, U.S. Agricultural
Research Service, Fresno, Calif.)

G. Culex tarsalis (Culicidae) larvae. Bottom insect apparently normal; middle insect mori¬
bund, showing abnormal curvature and distended thorax characteristic of the tetragonal
virus (picornavirus) infection. (Photograph courtesy of W. R. Kellen, Stored-Product In¬
sects Research Laboratory, U.S. Agricultural Research Service, Fresno, Calif.)

H. Culex salinarius (Culicidae) larva showing cuticular black spotting characteristic of in¬
fection with the tetragonal virus (picornavirus). (Photograph courtesy of the Gulf Coast
Mosquito Research Laboratory, U.S. Agricultural Research Service, Lake Charles, La.)

I. Apis mellifera (Apidae) adult showing the swollen abdomen characteristic of infection
with the chronic bee paralysis virus (picornavirus). (Photograph courtesy of A. J. Gibbs,
Department of Developmental Biology, Australian National University, Canberra, A.C.T.,
Australia.)

121

122

Figure 3

A. Pieris rapae (Pieridae) larvae. Top two larvae are dark green and normal; larva below
shows pale yellow-green coloration caused by a granulosis virus (Baculovirus) infection.
(Photograph courtesy of R. E. Teakle, Department of Primary Industries, Indooropilly
Brisbane, Queensland, Australia.)

B. Euxoa auxiliaris (Noctuidae) larvae. Left larva retarded and with abnormal pale colora¬
tion due to a granulosis virus (Baculovirus) infection; right larva normal. (Photograph
courtesy of G. R. Sutter, Northern Grain Insects Research Laboratory, U.S. Agricultural
Research Service, Brookings, S.D.)

C. Plodia interpunctella (Pyralidae) larvae. Lower six whitened larvae infected with a
granulosis virus (Baculovirus); upper six brown larvae normal. (Photograph courtesy of W.
R. Kellen, Stored-Product Insects Research Laboratory, U.S. Agricultural Research Ser¬
vice, Fresno, Calif.)

D. Wiseana sp. (Hepialidae) larvae, ventral aspect. Left larva normal; right larva shows
swollen and whitened intersegmental areas caused by a nuclear polyhedrosis virus
(Baculovirus) infection. (Photograph courtesy of S. G. Moore, Microbiology Department,
Medical School, Dunedin, New Zealand.)

E. Wiseana sp. larvae, ventral aspect close up. Normal larva on the left; larva on the
right infected with nuclear polyhedrosis virus and shows localized swelling due to distend¬
ed internal tissues. (Photograph courtesy of S. G. Moore, Microbiology Department,
Medical School, Dunedin, New Zealand.)

F. Wiseana sp. (Hepialidae) normal larva in open dissection. (Photograph courtesy of S.

G. Moore, Microbiology Department, Medical School, Dunedin, New Zealand.)

G. Wiseana sp. (Hepialidae) larva infected with nuclear polyhedrosis virus (Baculovirus);
open dissection; note disruption of fat body. (Photograph courtesy of S. G. Moore,
Microbiology Department, Medical School, Dunedin, New Zealand.)

H. Wiseana sp. (Hepialidae) larva infected with granulosis virus (Baculovirus); open
dissection; with fat body somewhat more proliferated than in normal larva of figure 3F.
(Photograph courtesy of S. G. Moore, Dunedin, New Zealand.)

I. Trichoplusia ni (Noctuidae) post mortem. Larvae infected with nuclear polyhedrosis
viruses (Baculoviruses) characteristically assume this posture, hanging by their posterior
prolegs when they die. (Photograph courtesy of A. M. Heimpel, Insect Pathology
Laboratory, U.S. Agricultural Research Service, Beltsville, Md.)

123

124

Figure 4

A. Blatta orientalis (Blattidae) cockroaches. Middle insect normal; top and bottom insects
have swollen abdomens revealing paler intersegmental membranes caused by infection
with a Rickettsiella sp. (Photograph courtesy of A. M. Huger, Institut fur biologische
Schadlingsbekampfung, Darmstadt, West Germany.)

B. Carabid beetle showing a swollen abdomen caused by infection with a Rickettsiella sp.
(Photograph courtesy of A. M. Huger, Institut fur biologische Schadlingsbekampfung,
Darmstadt, West Germany.)

C. Aphodius tasmaniae (Scarabaeidae) larva. Integumental black spotting caused by an
infection with a Rickettsiella sp.

D. Antitrogus (Rhopaea) morbillosus (Scarabeidae) larva showing abnormal blue-black
spiracles caused by an early infection with a Rickettsiella sp.

E. Antitrogus [Rhopaea ) morbillosus (Scarabaeidae) larva showing a later infection than
that shown in figure 4D. The blue-black discoloration caused by Rickettsiella sp. has pro¬
gressed to include the head, legs, and anal portions.

F. Antitrogus (Rhopaea) morbillosus (Scarabaeidae) larva. Same insect as that in figure
4E but post mortem, showing that the characteristic discoloration caused by a Rickett¬
siella sp. infection is retained for some time after death.

G. Antitrogus (Rhopaea ) morbillosus (Scarabaeidae) larva showing an external view of a
Rickettsiella sp. infection that causes swollen infected lobes of fat body to break off from
the normal tissue and float around loosely in the hemolymph; the arrow indicates the
most prominent loose floating lobes.

H. Antitrogus Rhopaea ) morbillosus (Scarabaeidae) larva, in open dissection, showing the
same insect as that in figure 4G. The fat-body lobes infected with Rickettsiella sp.
disperse in the dissection water when the diseased larva is opened.

I. Diabrotica speciosa vigans (Chrysomelidae) adults. Normal dark green insect below.

Top insect pale green with a yellow-green abdomen caused by infection with a Rickett¬
siella sp. (Photograph courtesy of J. R. Adams, Insect Pathology Laboratory, U.S.
Agricultural Research Service, Beltsville, Md.)

126

Figure 5

A. Othnonius batesi (Scarabaeidae) larva showing a late infection with an Entomopox-
virus. Arrow indicates abnormally whitened hypodermal tissue that is transparent in the
normal larva; infected larvae turn progressively dull white throughout (compare with the
generally normal appearance of the closely related larva in fig. 4G).

B. Chironomid larvae. Left larva shows abnormally opaque-white segmental fat body
caused by an Entomopoxvirus infection; right larva shows normal transparent aspect.
(Photograph courtesy of the Gulf Coast Mosquito Research Unit, U.S. Agricultural
Research Service, Lake Charles, La.)

C. Choristoneura fumiferana (Tortricidae) larvae. Left two larvae normal; right two larvae
show the extreme distension and lightening characteristic of this Entomopoxvirus infec¬
tion in late instar larvae. (Photograph courtesy of F. T. Bird, Forest Pest Management In¬
stitute, Sault Ste. Marie, Ontario, Canada.)

D. Wiseana sp. (Hepialidae) larva in open dissection, showing some increase in fat-body
tissue caused by an Entomopoxvirus infection but also by the spherical cysts formed by a
parallel infection of the fat body with a coccidian. (Photograph courtesy of S. G. Moore,
Microbiology Department, Medical School, Dunedin, New Zealand.)

E. Plutella maculipennis (Yponomeutidae) larvae. Left larval cadaver showing outgrowth
of an Entomophthora sp. fungus through the integument and forcible ejection of sticky,
infective spores, which form a ring around the cadaver at the limits of their range. Normal
larva on the right. (Photograph courtesy of R. E. Teakle, Department of Primary In¬
dustries, Indooropilly Brisbane, Queensland, Australia.)

F. Rhopaea verreauxi (Scarabaeidae) mummified, brown larval cadaver showing early
outgrowth of the infecting Hirsutella sp. fungus on incubation in a moist chamber for 5-7
days.

G. Anoplognathus sp. (Scarabaeidae) mummified (solidified), yellow larval cadaver as
recovered from the field. Compare this fungus infection with the same insect in figures 5H
and 51.

H. Anoplognathus sp. (Scarabaeidae) mummified larval cadaver after 3 days incubation in
a moist chamber; note early outgrowth of white hyphae of infecting fungus and early
(green) fruiting bodies (conidiospores) in small patches laterally on each segmental spiracle
of the host.

I. Anoplognathus sp. (Scarabaeidae). Same larval cadaver as in 5G and 5H but after 7
days incubation in a moist chamber, surface of larva covered with the characteristic
(green) conidiospores of the infecting green muscardine fungus Metarrhizium anisopliae.

127

128

Figure 6— Camera Lucida Drawings
of Fungi Viewed With a Microscope

A, Laboulhenia. Left, 2 one-septate ascospores with mucilaginous sheath; right, thallus
with appendages and a basal holdfast. B, Entomophthora. Conidiophores and conidia. C,
Ascosphaera. An ascocarp with several globose asci and ellipsoid ascospores. D,

Nomuraea. Conidiogenous cells and conidia. E, Aspergillus. A conidiophore with globose
head, sporogenous cells, and conidia. (Adapted from Raper and Fennel 1965.) F, Fusarium.
Macroconidia and microconidia. G, Penicillium. A branched conidiophore, conidiogenous
cells, and conidia. H-J, Hymenostilbe, Akanthomyces, and Insecticola, respectively. A
hymenium of sporogenous cells and conidia (adapted from Mains 1950a, 1951). K, Hir-
sutella. Sporogenous cells and conidia with mucilage. L and M, Tilachlidium and Syn-
nematium, respectively. Synnemata with conidiogenous cells and mucilaginous spore
balls. N, Cephalosporium. Mucilaginous spore balls on solitary conidiophores. O,

Beauveria. Conidiogenous cell, zigzag deticules, and conidia. P, Metarrhizium. Clustered
conidiophores and conidia. (Figure prepared by L. R. Batra, Mycology Laboratory, Plant
Protection Institute, Agricultural Research Service, U.S. Department of Agriculture,
Beltsville, Md. 20705.)

Micro-organisms as Contaminants
and Pathogens in Insect Rearing

By Martin Shapiro1

Introduction

Although historically insect pathology has concentrated
on the honey bee, Apis mellifera Linnaeus, and the
silkworm, Bombyx mori (Linnaeus), because of their
economic importance (see, for example, Bassi 1835,
Pasteur 1870, Steinhaus 1956), this paper will discuss
how micro-organisms may affect the rearing of many
species and what measures may be used to prevent or
minimize their impact (see also Steinhaus 1953, Helms
and Raun 1971, and McLaughlin 1971).

Micro-organisms may have little effect on insects, or they
may destroy an entire colony (Steinhaus 1953, 1968),
depending on the micro-organism involved and the mea¬
sures used to minimize or eliminate it. For example, dis¬
ease was a major obstacle to successful rearing of the
corn earworm, Heliothis zea (Boddie), in 1967-69 (Sparks
and Harrell 1976). And Raulston and Lingren (1972)
stated that viral and protozoan diseases and diet con¬
tamination by bacteria and fungi were often responsible
for insect-rearing failures. They concluded that failure to
control microbial contaminants and pathogens could off¬
set the value of new rearing techniques.

The most common microbial contaminants encountered in
insect cultures are Aspergillus spp. fungi (see, for exam¬
ple, Leonard and Doane 1966, Chawla et al. 1967,

Hensley and Hammond 1968, Kishaba et al. 1968, Fidet
1972, Griffin and Lindig 1973, and Ludemann et al.

1979). These fungi affect insect-rearing programs in
several ways. For example, Aspergillus growth covering
the rearing medium has prevented young pink bollworm,
Pectinophora gossypiella (Saunders), larvae from feeding
(Ouye 1962). But Aspergillus flavus Link and A. niger
van Tieghem growing as saprophytes on the feces of the
cabbage looper, Trichoplusia ni (Hiibner), do not harm it,
because contamination occurs mainly during prepupation
and pupation (Ignoffo 1964, 1966a). In one program,
which was rearing the codling moth, Laspeyresia
pomonella (Linnaeus), A. niger contamination occurred
mainly at the end of the larval feeding period (Howell
1971). The larvae left the areas where, in some instances,
the fungal hyphae had penetrated the diet. Then some

‘Research entomologist, Otis Methods Development Center,
Agricultural Research Service, U.S. Department of Agriculture,
Otis Air National Guard Base, Mass. 02542.

larvae died from starvation, and those crawling through
the fungal growth became covered with conidia; some of
these insects died, presumably from asphyxiation. The in¬
cidence of Aspergillus contamination increased in the
rearing area from year to year and reduced moth yield
from 125.5 moths per tray to 64.6 per tray.

Many other microbes can harm laboratory-reared insects.
For example, Doane (1969) found that hatching gypsy
moth, Lymantria dispar (Linnaeus), larvae are often
contaminated by bacteria and fungi. In general, these
micro-organisms are not pathogenic, but their growth on
artificial diet adversely affects larval development.
Baumhover et al. (1977) reported that, in one case,
microbial contamination of the diet caused losses of
80%-85% of a tobacco hornworm, Manduca sexta (Lin¬
naeus), culture despite preventive measures. Gast (1966)
reported that contamination from Pseudomonas spp. in
boll weevil, Anthonomus grandis grandis Boheman, cul¬
tures reduces adult yield and increases adult mortality.

Viruses are another group of micro-organisms that can
harm laboratory-reared insects. For example, the only lar¬
val disease observed in codling moth rearing is a
granulosis virus. In one program, this virus disease was
present initially in only a few insects, but two epizootics
occurred within a year and severely reduced insect pro¬
duction (Howell 1971). Similarly, examination of diseased
and dead codling moth larvae in a Russian rearing pro¬
gram showed that 55.3% were infected with a granulosis
virus (Pristavko et al. 1971). In another program, the in¬
cidence of granulosis increased among Indian meal moth,
Plodia interpunctella (Hiibner), larvae in successive
generations and caused many deaths (Spitler 1970). Early
attempts to rear the cabbage looper failed because of
repeated virus epizootics (McEwen and Hervey 1960),
and Henneberry and Kishaba (1966) concluded that
nuclear polyhedrosis virus was a major impediment in
large-scale rearing of the cabbage looper.

Protozoa, mainly microsporidia, can present serious pro¬
blems in the rearing of many insects, for example, the
honey bee; the European com borer, Ostrinia nubilalis
Hiibner; and com earworm. Laboratory cultures of the
alfalfa weevil, Hypera postica (Gyllenhal), often suffer
from high levels of infection caused by Nosema spp.
(Hsiao and Hsiao 1973). Nosemosis has also affected rear¬
ing of the boll weevil; Gast (1966) reported a dramatic in¬
crease in occurrence of Nosema spp. over eight genera¬
tions in the laboratory; this was correlated with reduced

130

egg production and eventual destruction of the colony.

He also reported a similar sequence of events when the
boll weevil colony was infected by another protozoan,
Mattesia grandis McLaughlin. A third protozoan, similar
to Glugea gasti McLaughlin, occurred in 0%-100% of the
weevils examined, without causing apparent symptoms.
During epizootics, some reduction in oviposition occurred
among infected females (Flint et al. 1972). In one pro¬
gram, adult European corn borers infected by the pro¬
tozoan Perezia pyraustae Paillot laid fewer eggs, had a
lower percentage of hatch, and had a shorter longevity
than controls (Lewis et al. 1971). In another program, the
cinnabar moth, Tyria jacobaeae (Linnaeus), which is used
for biological control of the tansy ragwort, Senecio
jacobaea L., was heavily infected by a microsporidian.

The disease caused high mortality among the larvae,
leading Bucher and Harris (1961) to conclude that it may
play a major role in the failure of mass-reared
lepidopterans to control weeds. Nosema infections have
been reported in colonized anopheline mosquitoes by Fox
and Weiser (1959), Vavra and Undeen (1970), and Hazard
and Lofgren (1971). But, Hazard (1971) concluded that
few epizootics occurred in colonies of anopheline species
native to the United States. In fact, he traced the origin
of the disease to colonies of Anopheles at the London
School of Hygiene and Tropical Medicine that were
originally collected from Africa and Asia.

Effect of the Rearing Environment
on Microbial Contaminants

Movement of insects from the field to the laboratory
often stimulates disease and microbial contaminants. Ser-
ratia marcescens (Bizzio), for example, is a common
organism that often becomes pathogenic in the laboratory
(Doane 1960, Wood 1961). This bacterium is typically
unable to invade healthy, unwounded insects but is trans¬
mitted with other bacteria when the insects bite each
other because they are crowded.

Often, bacteria contaminating insect colonies are those
that are more commonly associated with man (Sikorowski
1975, Hedin et al. 1978). As early as the 1830’s, Bassi
(1835) recognized that silkworm muscardine, caused by
the fungus Beauveria bassianna (Balsamo), spreads by
contamination of food and of insectary personnel. And
concentration of the insects into a confined space in¬
creases the chances of pathogen transmission (Gast
1966). Also, incidental fungal contamination may increase
with increase of larval populations in containers (Hensley
and Hammond 1968).

Steinhaus (1953) stated that controlling disease in
laboratory insects requires rearing them in the best possi¬
ble environmental conditions. High humidity and
temperature have often been associated with increased

occurrence of diseases and contaminants. First to
associate high humidity with fungal germination was
Bassi (1835); more recently, Howell (1970) reported that
A. niger develops rapidly on codling moth diet exposed to
warm, stagnant air. Steinhaus (1953) observed that
holding insects on diet in closed containers results in con¬
densation caused by a temperature gradient and often in¬
creases disease occurrence. Stephens (1962) reported that
mortality of greater wax moth, Galleria mellonella (Lin¬
naeus), larvae caused by a bacterium, Streptococcus
faecalis Andrewes and Harder, increases when
temperature and relative humidity are high. Likewise,
McLaughlin (1962) reported that mortality caused by the
bacteria Pseudomonas aeruginosa (Schroeter), Aerobacter
aerogenes (Kruse), and S. marcescens increases among
armyworm, Pseudaletia unipuncta (Haworth), larvae with
increase in temperature. And Steinhaus (1945) reported
that he controlled Serratia and Aerobacter infections of
the potato tuberworm, Phthorimaea operculella (Zeller),
by regulating environmental temperature. So, proper
design of the insect-rearing facility to allow control of en¬
vironmental conditions is important in preventing or
minimizing the occurrence of diseases and contaminants.

Effect of the Insect
on Microbial Contamination

Field-collected insects are often used to start and aug¬
ment colonies in the laboratory because of the availability
and the need for genetic variability. They are also a ma¬
jor source of parasites, pathogens, and microbial con¬
taminants. For example, Sutter et al. (1971) reported
field-collected eggs of the southern corn rootworm.
Diabrotica undecimpunctata howardi Barber, to be a ma¬
jor source of bacteria and fungi. Likewise, Dunbar et al.
(1972) reported that an entomogenous fungus,
Pascilomyces farinosus Brown and Smith, was observed
growing on gypsy moths in 88% of the collection sites.
Lynch and Lewis (1978) isolated four genera of fungi and
an unidentified yeast from European com borer egg
masses. Also, Lynch et al. (1976) reported isolating
bacteria of six different families, and several that were
unidentified, from European corn borer. Some of these
bacteria reduce egg hatch, and the Bacillaceae also reduce
larval establishment.

Viruses and microsporidia often occur at high rates in
field populations. Collection of wild insects for starting
laboratory colonies may result in introduction of these or¬
ganisms. Chauthani and Claussen (1968) reported a 40%
incidence of a natural virosis in Douglas-fir tussock moth,
Orgyia pseudotsugata (McDonnough), larvae from field-
collected egg masses; and Doane (1975) reported up to
95% mortality from viruses in field-collected gypsy moth
larvae. Likewise, the microsporidian Perezia pyraustae is

131

often reported from wild European corn borer populations
(see, for example, Raun 1966). Spruce budworm, Choris-
toneura fumiferana (Clemens), is often naturally infected
with a Nosema (Wilson 1974). Collecting insects from
areas where they are not dense should decrease chances
of colonizing diseased insects (Magnoler 1970a, 1970b), at
least in the case of the gypsy moth and the Douglas-fir
tussock moth (Chauthani and Claussen 1968).

Transmission of pathogens

Many pathogens are transmitted from one generation to
the next either on the egg surface (transovum) or in the
egg (transovarial). Hatching larvae get nuclear and cyto¬
plasmic polyhedrosis viruses from the egg surface (Bul¬
lock et al. 1969, Sikorowski et al. 1973, Doane 1975).
Gypsy moth larvae acquire virus from the egg surface or
from contaminated hairs as they hatch from the egg
mass (Doane 1975). Transmission through the egg was
observed over 100 years ago when Pasteur (1870) noted
that the protozoan N. bombycis could be transmitted
through the egg of the silkworm. Since then, it has been
demonstrated that other microsporidian parasites are
passed transovarially from one generation to another (see,
for example, Raun 1961; Ishihara and Fujiwara 1965;

Gast 1966; Gast and Davich 1966; McLaughlin 1966,

1971; Jenkins et al. 1970; Lewis et al. 1971; and Hsiao
and Hsiao 1973). Gast (1966) reported that the percent¬
age of boll weevil eggs containing M. grandis varies with
the percentage of diseased adult females; and, as the dis¬
ease spreads from adult to adult, the number of diseased
eggs increases.

Parasites are also transmitted when the spores are in¬
gested by healthy insects (Gast 1966, Hsiao and Hsiao
1973). After the gut epithelium is infected, transmission
probably occurs through contamination of feces (Bucher
and Harris 1961, Lewis et al. 1971). For example, in one
program, the rate of N. bombycis infection in silkworms
was high in the first, second, and fifth larval stages, cor¬
responding with a change in the number of larvae ex¬
creting Nosema spores (Ishihara and Fujiwara 1965).
Another mode of transmission may be cannibalism (Gast
1966).

It should be assumed that survivors from infected cul¬
tures may have an infection; for example, Bucher and
Harris (1961) found in one study that two-thirds of the
apparently healthy pupae of the cinnabar moth were in¬
fected with the microsporidian Nosema cerasivoranae
Thomson. Insect-to-insect transmission of micro-organisms
also occurs with virus diseases (Jacques 1962, Henneberry
and Kishaba 1966, Stewart et al. 1976) and fungus dis¬
eases (Bassi 1835, Hensley and Hammond 1968).

Diet Contamination
by Micro-organisms

Contamination of the diet with fungi (Ignoffo 1966a,
Leonard and Doane 1966, Chawla et al. 1967, Kishaba et
al. 1968) and bacteria (Afrikian 1960, Sutter et al. 1971,
Childress and Williams 1973) may be a minor inconven¬
ience or a serious impediment to laboratory or insectary
rearing. Dietary ingredients may be one source of micro¬
bial contaminants. Ignoffo (1965) isolated yeasts, molds,
and bacteria from vitamins, wheat germ, casein, and
water; and no microbial contaminants were isolated from
potassium hydroxide, methylparaben (methyl p-hydroxy-
benzoate), choline chloride, Formalin (formaldehyde),
Alphacel, agar, sucrose, or Wesson Salts (Ignoffo 1966a).

I have isolated contaminants from wheat germ, casein,
torula yeast, and from tap water used in gypsy moth
rearing; more than 95% of the total bacterial population
was isolated from the raw wheat germ (unpublished data).

Semisynthetic, artificial diets provide the nutrients essen¬
tial not only for insect growth and development but also
for growth and development of microbial contaminants.

In many instances, contamination of the diet, mainly by
fungi belonging to the genus Aspergillus (see, for exam¬
ple, Ouye 1962, Ignoffo 1966a, Chawla et al. 1967, and
Kishaba et al. 1968), is the most common contamination
problem. For example, Griffin et al. (1974) isolated two
bacterial species and one fungus, A. niger, from unsteri¬
lized boll weevil diets. McLaughlin and Sikorowski (1978)
isolated 16 bacterial species from insectary-reared boll
weevils and tested their growability on artificial diet; on¬
ly one bacterium, a Flavobacterium, could not grow on
the diet. They further tested 35 bacterial cultures, in¬
cluding human pathogens. Twelve of these grew on the
diet, including 5 of 15 human pathogens and 7 of 20 non¬
pathogens. In our laboratory at the U.S. Agricultural Re¬
search Service’s Otis Methods Development Center, Otis
Air National Guard Base, Mass., bacteria (Bacillus cereus
Frankland and Frankland, Staphylococcus aureus Rosen-
bach, Escherichia coli (Migula) Castellani and Chalmers,
Enterobacter aerogenes Hormaeche and Edwards, Pro¬
teus vulgaris Hauser) and fungi (Aspergillus niger van
Tieghem, Penicillium notatum Westling, Rhizopus spp.,
Saccharomyces cerevisiae Meyen) have been tested for
their ability to grow on a diet high in wheat germ (Bell et
al. 1981) as part of a study on antimicrobials. All of the
micro-organisms tested grew and developed on the insect
diet (M. Shapiro, unpublished data).

The diet’s pH is another factor that might favor or in¬
hibit microbial growth and development. In one test,
when the diet was adjusted to pH 5.5, the bacterial pop¬
ulation was 1.6 million/g (at day 7); at 6.5 pH, the
bacterial population increased to 290 million/g (Gifawesen
et al. 1975). At the Otis Methods Development Center.

132

elimination of antimicrobials from the gypsy moth diet
allowed contamination at pH’s 4-8; antimicrobials could
not inhibit microbial development at a dietary pH greater
than 6.5. And larval growth was retarded at pH 7 or
higher regardless of the insect diet used. Dietary pH’s of
4. 5-6. 5 (unadjusted pH’s: 5. 3-5. 8) were satisfactory for
rearing the insect, and contamination was minimal at the
acid end of the pH range (Bell et al. 1981).

Counteracting Micro-organisms
in the Insect and the Diet

Sanitation and decontamination

Strict sanitation measures are needed to minimize or
eliminate micro-organisms (see, for example, McEwen and
Hervey 1960, Gast 1966, Henneberry and Kishaba 1966,
Ignoffo 1966a, Lyon and Flake 1966, Raun 1966, Mag-
noler 1970a, Helms and Raun 1971, McLaughlin 1971,
Sutter and Miller 1972, and Sikorowski 1975). Both
rooms and equipment should be sanitized, and equipment
should be autoclaved where possible. Items that cannot
be autoclaved should be soaked in disinfectants such as
Formalin (see, for example, Steinhaus 1953, Gast 1966,
and Stewart et al. 1976) or sodium hypochlorite (see, for
example, Henneberry and Kishaba 1966, Martin 1966,
Odell and Rollinson 1966, and Baumhover et al. 1977).
Recently, Shimanuki et al. (1969) and Tompkins and
Cantwell (1975) have shown that ethylene oxide can be
used to decontaminate equipment and facilities.

Chemical disinfection of the insect

Whether insects are collected from the field or reared in
the laboratory, they are a source of contaminants and
pathogens. To minimize or eliminate the micro-organisms,
the insect should be disinfected. Any insect life stage
may be disinfected, but the egg stage is most often
selected.

Chemical antimicrobials. — No chemical antimicrobial is
perfect, and compromises must be made with each to fit a
rearing program’s particular needs. The two chemicals
most commonly used are sodium hypochlorite (NaOCl)
and formaldehyde (in Formalin). Ignoffo and Dutky
(1963) suggested that NaOCl was an ideal general disin¬
fectant because of its broad microbicidal spectrum of ac¬
tivity, good solubility in water, stability in aqueous
solutions, nontoxicity to humans and insects, availability,
and low price. Odor and skin irritation after prolonged
contact are disadvantages; and treatment of eggs with
NaOCl causes partial dechorionation, resulting in sus¬
ceptibility to desiccation and mechanical injury. In
general, two treatments, with variations, have been used
for disinfection. In the first, eggs are soaked in NaOCl
and rinsed in water (see, for example, Getzin 1962). In the

second, eggs are soaked in NaOCl, treated with sodium
thiosulfate to neutralize the chlorine, and rinsed in water
(see, for example, Ignoffo and Dutky 1963). These
methods have been used for egg disinfection (by Getzin
1962, Ignoffo and Dutky 1963, Grisdale 1968, Vasiljevic
and Injac 1971, Sikorowski et al. 1975, Beckwith and
Stelzer 1979, and many others), larval disinfection (see,
for example, King et al. 1979), pupal disinfection (see, for
example, Stone 1969), and adult disinfection (see, for ex¬
ample, Harein and de Las Casas 1968).

Formaldehyde is often used as an egg disinfectant (see,
for example, Golanski 1961, Lyon and Flake 1966, Chau-
thani and Claussen 1968, Magnoler 1970a, Sikorowski
1975, Bell et al. 1981, and many others). Other chemicals
that have been used as disinfectants include mercuric
chloride (see, for example, Barras 1972, Singh and Fowler
1973, and Kamano 1971); quaternary ammonium salts
(see, for example, Martignoni and Milstead 1960); cupric
sulfate (see, for example, Nettles and Betz 1966); para-
hydroxymethyl-benzoate (see, for example, Cothran and
Gyrisco 1966); and sorbic acid (see, for example, Karpel
and Hagmann 1968). (For detailed description of these
disinfectants, refer to Steinhaus 1953 and to “Microbial
Contamination in Insectaries. Occurrence, Prevention,
and Control,” by Peter P. Sikorowski, below.)

Effects of chemical disinfectants on micro-organisms. —
Protozoans are among the most difficult pathogens to re¬
move by disinfection. In these instances, the insect is most
often disinfected in the egg stage. Tyler (1962) successfully
removed spores of the sporozoan Triboliocystis gamhami
Dissanike from Tribolium spp. eggs by washing them in a
detergent solution. And a microsporidian ( Nosema algerae
Vavra) infection in Anopheles stephensi Liston was reduced
from over 90% to between 1% and 15% when eggs were
treated with water rinses alone (Alger and Undeen 1970).

But washing boll weevil eggs in a detergent (Triton X-100;
0.1%) for 5 minutes, mercuric chloride (0.06%) and NaOCl
(0.8%) for 10 minutes, or Formalin (4%) for 30 minutes,
failed to reduce an infection of M. grandis (Gast 1966).

Formalin and NaOCl have been highly effective in
eliminating or suppressing fungi. Ignoffo and Dutky (1963)
reduced the viability and infectivity of B. bassiana spores
by treating them with NaOCl. Sikorowski (1975) reported
that treatment of boll weevil eggs with 0.1%-0.2%

NaOCl produced contamination-free eggs, and treatment
of the eggs with 10% Formalin caused a 91% reduction in
fungal contamination.

Formalin and NaOCl are also highly effective in reducing
bacterial contamination (see, for example, Ignoffo and
Dutky 1963, Sikorowski 1975, Stewart et al. 1976, and
King et al. 1979). Other compounds have been used as
bactericides. For example, Martignoni and Milstead

133

(I960) tested two quaternary compounds, Zephiran Chlor¬
ide2 and Hyamine 10-X,3 4 and reported that they had low
mammalian toxicity, good bactericidal activity, and good
wetting action; however, only Hyamine 10-X adequately
removed bacteria from variegated cutworm, Peridroma
saucia (Hiibner), eggs. Barras (1972) tested Hyamine
10-X, mercuric chloride, White’s solution/ and a
modified White’s solution for pupal disinfection of
southern pine beetle, Dendroctonus frontalis Zimmerman,
and reported that only the modified White’s solution and
Hyamine 10-X effectively reduced bacterial contamina¬
tion. Nettles and Betz (1966) reported controlling bacteria
on boll weevil eggs by treating them with 18% cupric sul¬
fate solution (for 35 minutes) followed by treatment in a
25% ethyl alcohol solution with 0.04% mercuric chloride.

Formaldehyde and NaOCl are used alone and in combina¬
tion for control of viruses. Ignoffo (1964) reported effec¬
tively sterilizing cabbage looper eggs with NaOCl, and
Vail et al. (1968) demonstrated NaOCl and formaldehyde
to be effective in eliminating nuclear polyhedrosis virus.
Grisdale (1968) controlled virus infection in the forest
tent caterpillar, Malacosoma dis stria Hiibner, with a
0.5% NaOCl wash. Several studies report using NaOCl to
disinfect field eggs of the gypsy moth (see, for example,
Leonard and Doane 1966; ODell and Rollinson 1966;
Doane 1967, 1969, 1975; and Smith et al. 1976) to
minimize nuclear polyhedrosis virus levels.

Formaldehyde has often proven to be more effective than
NaOCl as a viricidal egg disinfectant. Thompson and
Steinhaus (1950) recommended the use of a 10% formal¬
dehyde solution (for 90 minutes) as an egg disinfectant.
Later, Golanski (1961) tested many chemical solutions for
controlling nuclear polyhedrosis virus in silkworm cul¬
tures and reported formaldehyde as most effective. Form¬
aldehyde was also found most effective for controlling
nuclear polyhedrosis virus in Douglas-fir tussock moth by
Lyon and Flake (1966) and Chauthani and Claussen
(1968); cytoplasmic polyhedrosis virus in pink bollworm,
by Bullock et al. (1969), Mangum et al. (1972), and
Stewart et al. (1976); granulosis virus in the Indian meal
moth, by Spitler (1970); and granulosis virus in Pieris
brassicae, by David et al. (1972).

Effects of chemical disinfectants on insects. — Chemical
disinfectants can harm the insect. For example, treat¬
ment of eggs with NaOCl may cause dechorionation that
allows the eggs to dry out and hatch early (see for ex¬

2Benzalkonium chloride.

3Methylbenzethonum chloride.

40.25 g Mercuric chloride, 6.5 g sodium chloride, 1.25 ml hydrogen
chloride, 250 ml 95% ethanol, and 750 ml sterile distilled water.

ample, Gast 1966, Vail et al. 1968, Sutter et al. 1971, and
Beckwith and Stelzer 1979). In many of these cases,
NaOCl was replaced with Formalin, which causes less
severe dechorionation (Bullock et al. 1969). But reduced
hatch after formaldehyde treatment of eggs has been re¬
ported (by Howell 1970, David et al. 1972, and 1975).
Reduced egg hatch has also been reported as a side effect
of treatment with other materials, such as methylparaben
and sorbic acid (Greene 1970), Hyamine 10-X, and
Zephiran Chloride (Sutter et al. 1971).

Heat treatment to disinfect insects

The egg stage for several species can tolerate higher
temperatures than the pathogen will. Finney et al. (1947)
and Allen and Brunson (1947) reported using this dif¬
ference to control Nosema in a potato tuberworm by
heating the eggs to 47° C. Likewise, immersing eggs of
European corn borer in a 43.4° C water bath for 30 min¬
utes to control P. pyraustae was a successful treatment
for several years (Raun 1961); this treatment eventually
lost its effectiveness, apparently because of selection of a
heat-resistant Nosema (Lewis and Lynch 1970). Heat has
also been used to reduce the incidence of cytoplasmic
polyhedrosis virus in the tobacco budworm, Heliothis
viriscens (Fabricius), by Roberson and Noble (1968),
Bullock et al. (1969), and Bullock (1972).

Thompson (1959) demonstrated that nuclear polyhedrosis
viruses of the cabbage looper and the bollworm (H. zea)
are not infective at rearing temperatures of 39° C or
higher. Ignoffo (1966b) confirmed that temperatures
above 55° C inhibit nuclear polyhedrosis virus infectivity
in Heliothis spp.; but nuclear polyhedrosis virus of the
yellowstriped army worm, Spodoptera omithogalli
(Guenee), is infective at temperatures as high as 46° C.
Similar results have been obtained for alfalfa caterpillar,
Colias eury theme Boisduval, infected by cytoplasmic
polyhedrosis virus (Tanada and Chang 1968); for Indian
meal moth infected by granulosis virus (Hunter and
Hartsell 1971); for codling moth infected by granulosis
virus (Pristavko et al. 1971); and for tobacco budworm in¬
fected by cytoplasmic polyhedrosis virus (Bullock 1972).
Unfortunately for disease control, rearing insects at high
temperatures can produce sterility and abnormal growth
(Bullock 1972).

Obtaining healthy insects

for rearing facilities

Obtaining healthy insects, when available, from other
rearing facilities may be the simplest, most effective way
to establish a disease-free culture (see, for example, Stein¬
haus 1953, Gast 1966, Brooks 1968, Helms and Raun
1971, and Tompkins and Cantwell 1975). When insects
are brought in from the field, it should be assumed that

134

they may be contaminated, diseased, or parasitized. Us¬
ing egg sources from areas free from obvious disease is
desirable. But the past history of the insect population
may not be known. Magnoler (1970b) collected gypsy
moth egg masses from light infestations, which he pre¬
sumed to be disease free, but disease might still have
been present. Grisdale (1968) collected forest tent cater¬
pillar egg masses in the fall, especially from areas of new
infestation where the level of disease and parasitism was
low; even so, eggs were often contaminated by
polyhedrosis viruses.

Field-collected insects should be kept isolated or quaran¬
tined from the main colony or rearing stock until their
state of health is ascertained. The incidence of disease
can be determined by not sterilizing some of the field-
collected material (Beckwith and Stelzer 1979). But gener¬
ally, the field-collected insects (egg stage, preferably)
should be disinfected to minimize or eliminate con¬
taminants. Spatial isolation or quarantine may not be
practical because of space limitations. In that case, other
measures must be taken, such as rearing the insects in¬
dividually or in small groups (Steinhaus 1953, Hen-
neberry and Kishaba 1966). This method may be time
consuming and cumbersome, but it will certainly
minimize transmission of pathogens between insects.

Pasteur (1870) helped to save the silk industry from the
ravages of pebrine, a protozoan disease caused by N.
hombycis, because, like Osimo (in an 1859 publication
cited by Steinhaus 1956), he recognized that the pathogen
could be transmitted from one generation to the next
within the egg. So he examined the adult females as a
way of insuring healthy eggs; if microscopic examination
showed spores in the moth, the moth and eggs were
destroyed. If no spores were found in the moth's tissues,
the eggs were saved and used for clean stock. This
method is still used successfully among commercial
breeders in Japan (Ishihara and Fujiwara 1965). And re¬
cently, variations of the Pasteur method have been suc¬
cessful in reducing or eliminating other protozoa (see, for
example, Bucher and Harris 1961; Gast 1966;

McLaughlin 1966, 1971; Jenkins et al. 1970; and Hamm
et al. 1971) and viruses (see, for example, Henneberry and
Kishaba 1966).

Diet treatments

Heat sterilization. — Little information exists on steriliza¬
tion of insect diets. Ignoffo (1965) demonstrated that
temperatures of 70°-75° C used in diet preparation reduce
the total contaminant level. This reduction may not be
enough, however, to eliminate the effects of dietary con¬
taminants. At the Otis Methods Development Center, we
boil the diet ingredients during processing; doing so in¬
creases the shelf life of the diet and reduces the level of

microbial contaminants (Bell et al. 1981) without hinder¬
ing growth and development of the gypsy moth. I have
found no differences in growth and virus yields between
gypsy moth larvae fed this standard diet and those fed
autoclaved diet (unpublished data). Similarly, Griffin et
al. (1974) found no statistical difference between boll
weevils reared on autoclaved (120°-125° C for 15-20
minutes) diet and those reared on diet that had been flash-
sterilized (130°-144° C for 30 seconds). A flash-sterilization
temperature of 151° C results in fewer and smaller wee¬
vils. At the U.S. Agricultural Research Service’s Boll
Weevil Research Laboratory at Mississippi State, Miss.,
where Griffin and his associates did this work, flash-
sterilization was adopted because it requires less labor
than autoclaving and saves time. Vanderzant (1975)
demonstrated that growth of the tobacco budworm was
not affected when its wheat germ diet was autoclaved;
but she recommended flash sterilization because of its
practicality.

Antimicrobial treatment. — Chemical preservatives, anti¬
microbials, are routinely added during diet preparation to
insect diets susceptible to contamination and spoilage.
These chemicals are the same as those accepted as food
additives by regulatory agencies of the United States and
most other Western countries (Chicester and Tanner
1968). Of these, the parabens, sorbates, benzoates, ace¬
tates, and proprionates are most commonly used in insect
diets (Singh 1977). Antibiotics are also used, especially
the tetracyclines and streptomycin, for their bactericidal
and bacteriostatic activities (Singh 1977).

Selection of antimicrobials will depend on the insect be¬
ing reared and on the contaminant. Sodium benzoate is
most active against yeasts and bacteria, and the para¬
bens are most active against yeasts and molds. Sorbic
acid and its salts have broad-spectrum activity against
yeasts and molds. Several acetates (vinegar and purified
acetic acid, sodium acetate, calcium acetate, potassium
acetate, and sodium diacetate) are most effective against
yeasts and bacteria (Chicester and Tanner 1968). For¬
malin is often used in diets to help control viruses (see,
for example, Ignoffo and Garcia 1968, Vail et al 1968,
and David et al. 1969). In tests of several antimicrobials
for control of bacteria and molds in gypsy moth artificial
diet, sodium omadine was the most effective antibacterial
compound, and sorbic acid was more effective than meth-
ylparaben, Formalin, or acetic acid. Methylparaben and
sorbic acid were the most effective antifungal com¬
pounds; when used together, they prevented fungal
growth (M. Shapiro, unpublished data).

Methylparaben is often used in combination with sorbic
acid and/or formaldehyde for control of several microbes,
particularly molds (see, for example, Gast and Davich
1966, Kishaba et al. 1968, and Howell 1971). Some re-

135

searchers (Nettles and Betz 1966, Burton and Perkins
1972) have reported these combinations to be ineffective
at the concentrations used. Similarly, Shorey and Hale
(1965) reported some resistance by molds to methylpara-
ben and sorbic acid used alone. And Gifawesen et al.
(1975), in assessing 37 fungicides for control of A. niger
in a wheat germ and casein diet, found that one A. niger
strain was resistant to methylparaben.

Seven of the fungicides studied by Gifawesen et al. (1975)
were effective. And Ludemann et al. (1979), in testing
other fungicides against resistant Aspergillus, found
some effective up to 14 days.

Fumagillin has been found effective in controlling various
protozoans: Nosema apis (Zander), by Katznelson and
Jamieson (1952), Bailey (1953), Moffett et al. (1969), and
Hartig and Przelecka (1971); Nosema spores, but not M.
grandis, in boll weevils, by Gast (1966); and P. pyraustae
in the European com borer, by Lewis and Lynch (1969,
1970), Lewis et al. (1971), and Lynch and Lewis (1971).
Fumagillin has also reduced incidence of N. fumiferanae
in spruce budworm larvae (Wilson 1972). And it has been
used in combination with other antimicrobials to control
Nosema spp. and other microbes (Hsiao and Hsiao 1973,
Gilliam and Morton 1974). Another agent effective in
suppressing Nosema spp. is benomyl6 (Shinholster 1974,
Armstrong 1976).

Of the antibiotics, chlortetracycline is the one most often
used. It controls yeasts and bacteria (Ignoffo 1963). The
antibiotics that have been successful in screening tests
against various micro-organisms include: streptomycin
sulfate for boll weevil diet (Gast 1966); methenamine
mandelate and nalidixic acid against S. marcescens (King
et al. 1975); erythromycin against the bacterium
Leuconostic mesenteroides van Tiegham on boll weevil
diet (Childress and Williams 1973); erythromycin thio¬
cyanate, erythromycin sulfate (Hitchcock 1964), and
sugar solutions containing tylosin actage, tylosin tar¬
trate, or sulfathiozole sodium (Hitchcock et al. 1970)
against European foulbrood in honey bee colonies;
Thipyrameth6 against the ameba Melameba locustae King
and Taylor in grasshoppers (Henry 1968); and maramycin
against Nosema apis in honey bees (Moffett et al. 1969). I
bioassayed B. cereus, P. vulgaris, E. coli, E. aerogenes,
and S. aureus against streptomycin, tetracycline, and
chlortetracycline on gypsy moth diet; all three antibiotics
were active against the five bacterial species at concen¬
trations of 0.05%-0.1%, except chlortetracycline,
which was ineffective against B. cereus.

‘Methyl l-(butylcarbamoyl)-2-benzimidazolecarbamate.

86% Sulfamethazine, 12% sulfathiozole sodium, and 8% sulfapyri-
dine sodium.

To be usable in insect diets, antimicrobials must work
over a wide pH range and resist degradation during
heating. The most effective pH range for sodium ben¬
zoate is 2.5-4.0; for the parabens, 3-9; for sorbic acid, up
to 6.5; for propionic acid, 3-5; and for the acetates, up to
5.2 (Chicester and Tanner 1968). The parabens and sorbic
acid are stable during autoclaving and flash sterilization
(Hedin et al. 1974).

Insects have been, and are still being, reared on natural
plant material. Often this plant material must be decon¬
taminated. Afrikian (1960) reported soaking of mulberry
leaves in bacteriostatic concentrations of streptomycin,
Aureomycin (chlortetracycline), and tetracycline to reduce
disease in the silkworm. Likewise, Zlotin (1965) sup¬
pressed disease in the gypsy moth by soaking acorns in
0.1% potassium permanganate.

Little use has been made of antimicrobial treatment to
destroy or inhibit microbial contaminants already grow¬
ing on the diet (spot treatment). At the U.S. Agricultural
Research Service’s Fruit Insects and Agricultural Engi¬
neering Research Unit at Yakima, Wash., Chawla et al.
(1967) tested sodium hypochlorite, methylparaben, and
sorbic acid both before codling moth eggs were implanted
on the diet and after fungi developed. Only sorbic acid
(1% solution) worked for the preimplantation treatment;
but, once the fungi had developed, each antimicrobial con¬
trolled fungal growth on the diet. Sorbic acid was then
used routinely in rearing the codling moth without harm¬
ing the insects. The spot treatment must have been effec¬
tive for the control of fungi, as it was still being used 4
years later (Howell 1971). Spot treatment was also used
successfully in the rearing of the European pine shoot
moth, Rhyaciona buoliana (Schiffermiiller), at Yakima.
Although this method is useful in some instances, it is
more prudent to discard the contaminated containers.

Antimicrobials have biological activity against the insect
as well as the microbe (Singh and Bucher 1971). Singh
and House (1970) and Singh and Fowler (1973) defined a
safe level of the antimicrobial agent as the concentration
that does not make the insect take more than 25% longer
than the control to grow and develop. The acceptable
level will probably have to be determined for each insect.
For example, Ouye (1962) recommended concentrations of
0.15% methylparaben and 0.05% Formalin in pink
bollworm diet because higher concentrations increase
duration of the larval and pupal stages. Moore et al.

(1967) reported that use of methylparaben and potassium
sorbate in boll weevil diet at concentrations greater than
0.5 ml inhibits egg hatch and larval growth. Henneberry
and Kishaba (1966) reported that concentrations of
methylparaben up to 8,000 p/m (parts per million) do not
harm cabbage loopers, but sorbic acid concentrations
greater than 4,000 p/m kill eggs. Other adverse effects on

136

cabbage looper growth and development include lack of
silk formation in cocooning caused by butylparaben
(re-butyl p-hydroxybenzoate) and increase of the larval
period caused by high concentrations of sorbic acid,
methylparaben, and butylparaben. Boush et al. (1968)
reported death of progeny of black carpet beetle, At-
tagenus megatoms (Fabricius), and the beetle Trogoderma
parabile Beal fed sorbic acid in the larval stage. Neilson
(1973) reported that methylparaben, butylparaben, and
sorbic acid hinder egg hatch of the apple maggot,
Rhagoletis pomonella (Walsh). Gifawesen et al. (1975)
reported thimerosal (merthiolate) as an effective mold in¬
hibitor but toxic to tobacco budworm larvae; and
Ludemann et al. (1979) reported that agricultural
fungicides incorporated into the diet prolong larval
development and decrease pupal weights. Ignoffo (1963a)
reported that chlortetracycline (0.5 mg/ml) inhibits cab¬
bage looper larval development, and, at 1.0 mg/ml, larvae
fail to molt. Gast (1966) and Flint et al. (1972) found that
boll weevils fed various levels of fumagillin develop ab¬
normally. Fumagillin used to control Nosema also harms
European corn borer (Lynch and Lewis 1971), alfalfa
weevil (Hsiao and Hsiao 1973), and spruce budworm
(Wilson 1974). Antimicrobial effects on insects may be
immediate (toxic) or long term (ovicidal). These effects
must be considered in any rearing system.

Conclusions

Steinhaus (1953) suggested the following principles that
could be used to suppress or eradicate a disease and pre¬
vent its recurrence:

1. Whenever possible, diagnose the disease. Identify the
contaminant or microbe.

2. If a disease is present in at least 25% of the insects, it
is best to destroy the stock and decontaminate the
facility and equipment. Reestablish the stock from a
healthy culture if possible. In the case of chronic infec¬
tions (for example, protozoan infections), selection of
healthy insects may be feasible.

3. Cleanliness should be maintained constantly for the
insect, the rearing environment, the equipment, and
the food.

And Steinhaus (1968) observed that contaminant or
pathogen suppression not only means sterilization and
sanitation but also “new methods of prevention and
therapeutics, more precise studies on the role of stressors
(e.g., adverse temperature, humidity, food and space) in
bringing about outbreaks of disease, and a greater under¬
standing and application of what has been learned by
those who for so long a time have been concerned about
suppressing disease in the silkworm and the honey bee.”

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1973. Improved method for rearing apple maggot
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142

Microbial Contamination in Insectaries

Occurrence, Prevention, and Control

By Peter P. Sikorowski1

Introduction

Steinhaus and Martignoni (1970) defined contamination
as the harboring of, or having contact with, micro¬
organisms without a relationship that is commensalistic,
mutualistic, or parasitic. Microbial contaminants are
usually composed of seemingly innocuous microbes. But
animal pathology has shown that micro-organisms pre¬
viously considered as innocuous, commensals, or contam¬
inants may multiply extensively in the tissue of weak¬
ened hosts and cause disease that is often severe (Neter
1974).

About 450 species of phytophagous insects have been
reared on synthetic diets (1977). Synthetic and semi¬
synthetic insect diets are usually complex media subject
to spoilage by many species of bacteria and fungi (see, for
example, Ignoffo 1966, Singh and House 1970a, Sikorow¬
ski 1975, McLaughlin and Sikorowski 1978, and others).
Spoilage is the result of metabolic activity associated
with microbial growth, causing catabolism of the media
and release of products of digestion. The biochemical
changes produced by microbes alter the nutritional value
of the diets. Also, some bacteria and fungi produce toxins
that may harm insects.

Humans are also affected by microbial contamination.
Several airborne micro-organisms such as Aspergillus,
Pseudomonas, and Streptococcus spp., which grow on
almost any organic matter, are also human pathogens
and present some hazard to employees in insect-rearing
programs.

The cost of rearing insects can be greatly reduced by
establishing an environmental sanitation program (Sikor¬
owski 1975). Much information pertinent to laboratory
rearing of insects has been obtained only recently, and
most areas covered in this report merit further study.

But research results to date suggest that sanitation will
play an important part in mass rearing of most insects
and that more attention wiU have to be given to enforce¬
ment of basic sanitary measures to assure volume pro¬
duction of healthy insects.

Occurrence

of Microbial Contamination
Sources of contamination

Humans are the primary source of microbial contamina¬
tion, and the level of contamination relates directly to ac¬
tivity and density of personnel (Favero et al. 1966, 1968;
Runkle and Phillips 1969). The healthy human body har¬
bors millions of micro-organisms on the skin in the
mouth, respiratory tract, genitourinary tract and intes-

Staphylococcus aureus
Escherichio coli
Streptococcus viridans

Staphylococcus spp
Corynebacterium spp
Bacillus spp

Streptococcus spp
Staphylococcus spp.
Neisseria spp.
Corynebacterium spp.

Staphylococcus

aureus

£2°'ngt>(,c' Ve'Hwt

Streptococcus spp

Staphylococcus spp.
Bacillus spp.
Lactobacillus spp.
Corynebacterium spp

‘Professor, Mississippi Agricultural and Forestry Experiment
Station, Department of Entomology, Drawer EM, Mississippi
State University, Mississippi State, Miss. 39759.

Figure 1.— Prevalent microbial flora of man (based
on National Aeronautics and Space Ad¬
ministration 1969).

143

Figure 2.— Relative composition of microbes on (A) clean, freshly laundered uniform and (B) worn
uniform.

tines (fig. 1). Jawetz et al. (1978) arranged the normal
microbial flora of the human body into two groups: The
resident flora consist of fixed types of micro-organisms
regularly found in a given area and in persons of a given
age. The transient flora consist of nonpathogenic or po¬
tentially pathogenic micro-organisms that inhabit the
skin and mucous membranes for hours, days, or weeks; it
is derived from the environment, does not produce dis¬
ease, and does not establish itself permanently on the
surface. Jawetz et al. (1978) reported that constant
exposure to and contact with the environment
results in contamination of the skin with transient
micro-organisms.

Smith and Bruch (1969) monitored healthy individuals
who exercised naked for 30 minutes and found that each
dispersed into the air 2-6 million viable micro-organisms.
Sikorowski (1975) showed that freshly laundered and
sterilized cotton uniforms do not provide an efficient bar¬

rier for movement of bacteria through the cloth into the
environment (fig. 2). The hair of 50 individuals (1 hair per
person) had from several to many bacteria per hair.
Cultures from hands almost always produced bacterial
colonies even shortly after normal washing.

It is generally agreed that properly cared for equipment,
instruments, walls, floors, etc. are minor sources of
microbial contamination. The microbial content of the air
in an area usually reflects the total microbial contamina¬
tion of the surrounding area (Loughhead and Moffett
1971).

Normal flora of wild insects

Understanding the microflora of insects reared in insec-
taries is based on a knowledge of the normal flora of wild
insects. The most commonly occurring, internally har¬
bored micro-organisms in insects are bacteria or bacteria-

144

like forms that are found in Blattodea, Isoptera, Homop-
tera, Heteroptera, Phthiraptera, Coleoptera, Hymenop-
tera, and Diptera. Also, flagellates are found in wood¬
eating insects and yeast and yeast-like organisms in
Homoptera and Coleoptera (Steinhaus 1949 and Chapman
1971). Steinhaus (1941) studied the bacterial flora of 30
species of insects and isolated 83 strains of bacteria, 2
strains of yeasts, and 2 molds from the insects. And, in
most cases, the species of bacteria found in the several
specimens of any given insect species were surprisingly
constant. A study of the internal microbial flora of 2,016
unfed specimens of the Rocky Mountain wood tick, Der-
macentor andersoni Stiles, showed that only 1.6% of the
adults harbored bacteria (Steinhaus 1942). Over 75% of
field-collected adult boll weevils, Anthonomus grandis
grandis Boheman, of various ages examined by Sikor-
owski et al. (1977) and McLaughlin and Sikorowski (1978)
had 100 or fewer bacteria per insect. Antibacterial con¬
stituents (such as gossypol, caryophyllene, gallic acid,
and tannins) found in host cotton plants are responsible
for the low bacterial content of wild weevils (Hedin et al.
1978). Greenberg (1962) reported that 73% of house flies,
Musca domestica Linnaeus, and 54% of stable flies,
Stomoxys calcitrans (Linnaeus), are germ free at
emergence from the puparium. He concluded that ster¬
ilization of the digestive tract is the usual consequence of
metamorphosis.

Insects that feed on diets deficient in certain elements
usually have micro-organisms associated with them. This
suggests that the micro-organisms make good the diet de¬
ficiencies. But, in many cases, the precise nature of the
micro-organisms is not known (Chapman 1971; for
reviews of the literature about micro-organisms of
healthy insects, see Steinhaus 1940 and Brooks 1963).

Bacterial flora

of insectary-reared insects

Insects reared in an insectary are often contaminated
with different species of aerobic and anaerobic bacteria
(see, for example, Ignoffo 1966; Sikorowski 1975; Davis
1976; McLaughlin and Sikorowski 1978; Smalley and
Ourth 1979; Bell et al. 1981; and others); relative com¬
position of contaminants often changes from day to day.

I have isolated, from apparently normal insectary-reared
boll weevils, many taxonomically unrelated species: from
the family Micrococcaceae, Micrococcus luteus (Schroeter)
Cohn and Staphylococcus aureus Rosenbach; from Strep-
tococcaceae, a Streptococcus sp. and Leuconostoc
mesenteroides (Tsenkovskii) van Tieghem; from
Bacillaceae, Bacillus sphaericus Meyer and Neide; from
Lactobacillaceae, Lactobacillus plantarum (Orla-Jensen)
Bergey et al.; from Pseudomonadaceae, Pseudomonas
aeruginosa (Schroeter) Migula; from Enterobacteriaceae,
Enterobacter aerogenes Hormaeche and Edwards; and

from the Coryneform group, Corynebacterium hurniferum
Seliskar (unpublished data). Pseudomonas aeruginosa, M.
luteus, a Streptococcus sp., Serratia marcescens Bizio, S.
rubidaea (Stapp) Ewing et al., Aspergillus niger van
Tieghem, A. flauus Link, Rhizopus nigricans Ehrenberg,
a Cladosporium sp., a Fusarium sp., and two yeast
species were isolated from laboratory-reared larvae of
Heliothis spp. by Bell et al. (1981). McLaughlin and
Sikorowski (1978) tested 35 bacterial cultures, mostly
from the American Type Culture Collection, for their
ability to grow on the boll weevil diet. Five of 15 human
pathogens and 7 of 20 saprophytes developed on the diet.

Effects of bacterial
contamination on insects

The number of insects that emerge from contaminated
diets is influenced by species of micro-organisms present,
stage of the insect at the time of contamination, tem¬
perature, pH of the medium, and perhaps by other fac¬
tors. For example, in one study, I found that boll weevil
diet contaminated with S. aureus at the time of egg im¬
planting produced 57% as many weevils per petri dish as
did uncontaminated diet, 57% for diet contaminated 3
days after larval hatch, and 71% for diet contaminated 6
days after larval hatch (unpublished data). Likewise, con¬
tamination of the diet at the same time intervals with a
Streptococcus sp. provided only 76%, 64%, and 80% as
many weevils per petri dish as did uncontaminated diet.
The normal developmental time of 13 days from egg to
adult weevil extended to 14 or 15 days on diets con¬
taminated with some species of bacteria. Fast-growing
bacteria such as Leuconostoc spp. may overgrow an area
of diet of about 700 mm2 in 24 hours and smother all the
larvae on the dish. The staphylococci (enterotoxin pro¬
ducers) may harm larvae.

Maeda et al. (1953) reported that, under aseptic condi¬
tions, larvae of the oriental fruit fly, Dacus dorsalis
Hendel, develop equally well in diets of pH 4.5, 5.5, 7.0,
and 8.0. But, when aseptic techniques are not followed,
any medium with a pH higher than 5.5 has poor larval
growth because bacterial contamination is heavy. At
room temperature, the pH of diet for phytophagous in¬
sects decisively influences the microbe’s ability to flourish
on it. For most bacteria, a minimum pH is 4. 5-5.0, max¬
imum 8.0-8. 5, and optimum 6.5-7. 5 (Oginsky and Um-
breit 1955). Approximate minimum and maximum pH
values for molds are 1.5-11.0, and for yeasts 1.5-8. 5 (Jay
1978). But the metabolic activity of a bacterium may pro¬
duce several acidic and basic products from the compo¬
nents of the medium. The release of such substances
during bacterial growth would shift the pH of the
medium. In general, insect media with pH 4 and above
are subject to bacterial spoilage. Media with pH 4 and
below may undergo mold and yeast spoilage.

145

G. A. Virginio (personal communication) found that
infection of lepidopterous larvae with microbial contami¬
nants causes high mortality and delay in larval develop¬
ment. He also observed that, when infections do not kill
the larvae, pupae and adults are abnormally small.

McLaughlin and Sikorowski (1978), Sikorowski and
Thompson (1979), and MacGown and Sikorowski (1980)
showed destructive effects of bacterial contamination on
the brush border of the midgut epithelium of the boll
weevil. In contaminated weevils, the brush border is fre¬
quently ulcerated and overgrown by bacteria. In testing
how bacteria affect pheromone production in the boll
weevil, Gueldner et al. (1977) and G. Wiygul and P. P.
Sikorowski (unpublished data) measured the amount of
pheromone of the male boll weevil in the feces; and P. A.
Hedin and P. P. Sikorowski (unpublished data) measured
the amount in body homogenate. More pheromone was
always isolated from the frass and body homogenate of
uncontaminated weevils than from contaminated weevils.
Pheromone production, and therefore attractiveness of
the sexually sterile weevil, is a desirable characteristic in
mass-reared weevils; the quality of weevils that are
bacterially contaminated is reduced to significantly below
normal, and these weevils would be expected to have a
greatly diminished competitive value. Thompson and
Sikorowski (1978) and Thompson et al. (1977) studied
how a Streptococcus sp., Micrococcus varians Migula,
and E. aerogenes contamination affect amino and fatty
acid contents of boll weevils. They reported that, except
for tyrosine and glutamine in the females, all amino acids
analyzed are found in greatest amounts in weevils free of
bacterial contamination. The average reduction of amino
acids due to bacterial contamination is 33% in males and
52% in females. The high level of tyrosine and glutamine
in contaminated female weevils was not explained. Com¬
parison of individual fatty acids between the groups
shows a decrease of up to 76% in highly contaminated in¬
sects ( _> 210,000 bacteria per insect). Hurej et al. (in
press) reported that bacterially contaminated boll weevils
react differently to insecticide treatment than do those
free of contamination. The bacteria tested decreased mor¬
tality in all groups of boll weevils treated with methyl
parathion.2 The effects of contamination on the toxicity
of mirex3 varied with the species of bacteria. So bacter¬
ially contaminated boll weevils have differed from uncon¬
taminated insects in all variables tested. This, to my
knowledge, is the first attempt to evaluate the effects of
bacterial contamination on the quality of artifically
reared weevils or any other insectary-reared insect.

20, 0-Dimethyl 0-(p-nitrophenyl) phosphorothioate.

sDodecachlorooctahydro-l,3,4-metheno-l//-cyclobuta[c<i]

pentalene.

Prevention

To a rearing program, contamination can mean increased
expense, poor quality of insects, high mortality of all
stages, additional workload, and loss of confidence in the
work. To prevent contamination, insect diets must be of
good sanitary quality, so they must be free of hazardous
micro-organisms, and saprophytes should be at accept¬
able levels. To overcome the problems caused by contam¬
ination and to prevent its recurrence, insectary personnel
must be trained in the need for sanitation, and regular
sanitary measures must be established and maintained.
Monitoring techniques are used to demonstrate that the
system can reliably keep microbial contamination below
undesirable levels.

Training of insectary personnel

Personnel must be trained to understand rearing opera¬
tions and that their own potential for contaminating the
area is high. Training programs for insectary workers
should begin with a discussion of the quality of insect re¬
quired. Most people without basic training or experience
in sanitary microbiology have difficulty comprehending
the presence of microbes on apparently clean hands, gar¬
ments, and equipment. Sikorowski (1975) described
inservice training conducted in the U.S. Animal and
Plant Health Inspection Service’s Robert T. Gast Boll
Weevil Rearing Facility at Mississippi State, Miss., to
familiarize all employees with the basic concepts of
sanitary biology. A major part of the training was the
conducting of experiments that showed the presence of
microbes on unsanitized floors, clothing, shoes, and hair,
and in the breath.

Sanitary measures

The objective of establishing an environmental sanitation
program is to eliminate contamination that interferes
with the rearing of healthy insects. The level of cleanli¬
ness required must be decided on before the needed tech¬
niques are determined. Many researchers have described
different sanitation programs used in their insectaries
(see, for example, Steinhaus 1953; Sikorowski 1975; Davis
1976; Martignoni and Iwai 1977; and Bell et al. 1981).
Most of these include in their sanitary measures personal
hygiene; maintenance of a clean, sanitary environment by
proper housekeeping; and various methods of sterilizing
and sanitizing insectary equipment and environment.

Personal hygiene.— Micro-organisms are usually shed
with the human skin scales they are attached to. Taking
a shower degerms the skin by causing about a five-fold
increase in the rate of shedding micro-organisms. The
high population of normal flora returns in 1-2 hours after
thorough showering (National Aeronautics and Space Ad-

146

ministration 1969). The major purpose of clean uniforms
is to limit the passage of skin microbes to the environ¬
ment; some microbes escape when the uniform is not pro¬
perly closed. Frobisher et al. (1969) reported that the
human mouth and throat constantly contain many kinds
of micro-organisms. The micro-organisms multiply in the
secretions of the nose and oral cavity and between the
teeth. Commonly found are streptococci, staphylococci,
pneumococci, and spirochetes. Sikorowski (1975) reported
that personnel concerned with handling insect diets and
adult boll weevils and with planting eggs were required
to wear caps and wash their hands frequently with Surgi-
Bac (Economics Laboratory, St. Paul, Minn.). Workers
handling weevil diets were also required to wear hair caps
and face masks.

Housekeeping.— Litsky and Litsky (1968) and many
other workers in the field of microbial contamination
have demonstrated that flooding and wet-vacuuming is
the most effective method for cleaning and disinfecting
floors. Portable vacuum cleaners may be used only if the
vacuum cleaner exhaust is filtered to the same extent
that the air is filtered through the air-filtration system in
the room. Sweeping and dry-mopping might stir up
micro-organisms and should be avoided (Nagasawa et al.
1970). Runkle and Phillips (1969) recommend elimination
of sweeping through the use of dry or wet pickup vacuum
cleaners with high-efficiency exhaust air filters. The
general housekeeping routines are given by Sikorowski
(1975), Davis (1976), and Martignoni and Iwai (1977).

Sterilization of equipment.— “Sterilization" means the
destruction of all life. Heat is the most reliable and
universally applicable method of sterilization and should
be the method of choice whenever possible. All living
organisms can be rapidly destroyed by steam under
pressure. The greatest problems for steam sterilization
are moisture penetration, air removal and entrapment,
heat and moisture damage, and thorough wetting (Ernst
1977). (See Martignoni 1978 for the principles and proper
use of the autoclave and Hedin et al. 1974 for how auto¬
claving and flash sterilizing affect potassium sorbate and
methyl p-hydroxybenzoate (methylparaben) in boll weevil
diets.)

Ultraviolet light.— The most effective antibacterial wave¬
length of ultraviolet light is between 240 nm and 280 nm;
the optimum is about 260 nm. Ultraviolet light kills bac¬
teria because it is absorbed by and damages the DNA
(Willett 1976). Ultraviolet light has poor penetrability; it
does not penetrate solids, and it penetrates liquids very
slightly. Micro-organisms are protected by dust in the air
and by dust on the ultraviolet-light bulbs; and clumping
of microbial cells (microbial cells attached to dust) allows
those in the central part of the clump to escape the effect
of ultraviolet light. At the R. T. Gast Rearing Facility,

ultraviolet light failed to produce uniform results in the
control of airborne micro-organisms in ultra-violet light
passthrough box units used for passage of materials from
one area of the laboratory to another.

Sanitizing equipment.— Large equipment, such as a diet¬
processing apparatus that is permanently attached to the
walls, should be sanitized immediately before and just
after use to prevent entry of microbes into the apparatus.
Small instruments that cannot be autoclaved may be
sanitized by immersion in sanitizing solution. Procedures
used to sanitize equipment are given by Sikorowski
(1975), Martignoni and Iwai (1977), and Griffin et al.
(1979).

Air disinfection and air filtration. — Air filtration is one of
the most important tools against microbial contamina¬
tions. Many different types of air-cleaning equipment are
available— gravitational, inertial, filtration, washing, and
electrostatic precipitation. Filtration is usually recom¬
mended when maximum removal of bacterial particles
from the air is required.

Particles carrying bacteria are usually greater than 4//m
in diameter, with a median of 10-20 /xm, although single
spores of bacteria are often not associated with dust car¬
riers (National Aeronautics and Space Administration
1968). HEPA (high-efficiency particulate air) filters
remove 99.97% of particles that are 0.3 umin diameter
and larger. This laminar airflow prevents microbial ac¬
cumulation in a room atmosphere. Laminar-airflow
devices can be classified as follows: horizontal-airflow
units, vertical-airflow units, and curvilinear-airflow
(change in the direction of the air during its flow) units.

In the R. T. Gast Rearing Facility, air in the egg-
implanting area has been filtered with considerable suc¬
cess through HEPA filters having a 0.3-/nm pore size
(Sikorowski 1975). After several years’ experience in the
use of laminated airflow, I find that this method is one of
the most useful tools in controlling microbial con¬
taminants. (See Phillips and Runkle 1973 for a general
discussion of principles of laminar airflow and its possible
biomedical applications.)

Fogging a room.— Greene (1970), in discussing the advan¬
tages and disadvantages of fogging a room with ger¬
micides, reported that heavy airborne particles settle by
gravity while the room is left undisturbed; so the objec¬
tive changes from air decontamination to surface decon¬
tamination. He also found that the lighter microbial
particles have only a remote chance of contacting a ger¬
micidal droplet and of staying with it in a moist state
long enough to be killed. Friedman et al. (1968)
demonstrated that spray-fogging hospital rooms with a
quaternary ammonium disinfectant effectively reduces
the number of detectable airborne and surface bacteria.

147

They also observed that fogging with quaternary am¬
monium disinfectant does not produce local or generalized
irritation in housekeeping personnel and technicians. F.

M. Davis’ (unpublished data) and Sikorowski’s (1975)
studies also show that spray-fog disinfection with
quaternary disinfectant usefully decreases airborne
micro-organisms .

Antimicrobial compounds. — Below is a list of the major
groups of antimicrobial compounds used for cleaning and
sanitizing.

1. Iodophors.— Iodophors are combinations of iodine and
a carrier. Their activity is based on the slow release of
iodine. Iodine is effective against a wide variety of
micro-organisms; it is lethal to viruses, bacteria (in¬
cluding spore formers), fungi, protozoa, and algae. But
recent works suggest that iodine is less sporicidal than
hypochlorite (Trueman 1971). Mikroklene (Economics
Laboratory, St. Paul, Minn.) is an iodophor detergent-
disinfectant claimed by the manufacturer to be effec¬
tive against the spores of Bacillus subtilis (Ehrenberg)
Cohn and Clostridium sporogenes (Heller) Bergey et al.
at a concentration of 2.0% with a temperature of 80° C
and an exposure time of 30 minutes.

2. Quaternary ammonium compounds.— The bactericidal
power of the quaternaries is high against Gram¬
positive bacteria, but they are less effective against
Gram-negative bacteria. They are not sporicidal. They
also have bacteriostatic characteristics far beyond
their bactericidal concentration. They are also
fungicidal and can destroy certain sanitizing agents.
Other features are low mammalian toxicity, low cor¬
rosiveness, and combined properties of germicidal and
detergent activity. They are stable, odorless, do not
stain, dissolve easily in water, and are inexpensive.
Commerical quaternary ammonium compounds include
Bradophen4 * (Ciba-Geigy Ltd., Basel, Switzerland),
Mikro-quat6 (Economics Laboratory, St. Paul, Minn.),
and Roccal6 and Quatsyl 2567 (National Laboratories,
Montvale, N.J.). (See Lawrence 1950 and Petrocci 1977

4Benzyl-dodecyl-bis-(2-hydroxyethyl)-ammonium chloride.

‘Active ingredients: alkyl (50% C14, 40% C12, 10% Cl6) dimethyl

benzyl ammonium chlorides, 9.0%; trisodium ethylenidiamine-

tetraacetate, 0.4%.

‘Benzalkonium chloride.

’Active ingredients: octyl decyl dimethyl ammonium chloride,
3.750%; dioctyl dimethyl ammonium chloride, 1.875%; didecyl
dimethyl ammonium chlorides, 1.875%; alkyl (50% C14, 40% Ci2,
10% C16) benzyl dimethyl ammonium chloride, 5.000%; tetra-
sodium ethylenediamine tetraacetate, 3.420%; isopropyl alcohol,
3.000%; ethyl alcohol, 1.000%.

for reviews of quaternary ammonium compounds and
Martignoni and Milstead 1960 for discussion of their
use in surface sterilization of insects.)

3. Phenolic compounds.— Phenol in high concentrations is
a protoplasmic poison that penetrates and disrupts the
cell wall and precipitates the cell proteins. At lower
concentrations, it inactivates the essential enzyme
systems. Phenolic compounds are effective against
vegetative cells of bacteria, and, at high temperatures,
against fungal and bacterial spores. Commercial
phenolic compounds include Amphyl8 (National
Laboratories, Montvale, N.J.) and Con-O-Syl9 (Johnson
and Son, Racine, Wis.).

4. Sodium hypochlorite.— Hypochlorites (calcium and
sodium) are the oldest and most widely used chlorine
compounds in the field of chemical disinfection
(Dychdala 1977). They are effective against a wide
spectrum of micro-organisms, including spore formers.
The sodium hypochlorite solutions range in concentra¬
tions from 1% to 15% with l%-5% available chlorine
products used domestically and stronger solutions in¬
dustrially. Today, hypochlorites are used for microbial
control in households, hospitals, schools, restaurants,
food-processing plants, dairies, canneries, wineries, and
beverage bottling plants (Dychdala 1977). Hypochlor¬
ites are also extensively used as disinfectants in
rearing facilities. Sodium hypochlorite solution is
available in most grocery stores as a laundry bleach
(for example, Clorox and Purex).

5. Formalin.— Formalin is commonly a 37%-40% solution
of formaldehyde in water. This antimicrobial com¬
pound is sporicidal, bactericidal, and, at lower concen¬
trations, bacteriostatic. Formalin contains certain
stabilizers, 8%-15% methanol being the one most com¬
monly used, to prevent the formation of solid polymers
if the solution is chilled or is kept for a long time
(Phillips 1977). Formalin is most effective at high
relative humidity. It penetrates very slowly; therefore,
its application should be limited to surface sterili¬
zation. Although Formalin has been widely used for at
least 80 years, it'should be used only when no other
method is available. (For more information about For¬
malin, see Hoffman 1971 and Trujillo and David 1972.)
Recently, a New York University study has provided

‘Active ingredients: (o-phenylphenol, 10.5%; o-benzyl-p-chloro-
phenol, 5.0%.

‘Active ingredients: o-phenylphenol, 11.75%; o-benzyl-p-chloro-
phenol, 7.82%; sodium dodecylbenzenesulfonate, 3.17%; tetra-
sodium ethylene diamine tetraacetate, 2.96%; isopropyl alcohol,
2.60%; essential oils, 0.30%.

148

decisive confirmation of industry findings that form¬
aldehyde is an animal carcinogen (Sun 1981).

6. Soaps.— Many different soaps have bacteriostatic or
bactericidal properties. The germicidal soaps are or¬
dinary toilet soaps with various antimicrobial agents
added to a maximum concentration of 2%. Phenol deriv¬
atives, especially creosols and bis-phenols are often
mixed with soaps. (For more information on soaps, see
Frobisher et al. 1969 and Hamilton 1971.)

Contamination-monitoring program

Contamination control manages microbial contamination
at desired levels. The monitoring program evaluates the
effectiveness of the contamination-control program. One
of the earliest systems for monitoring microbial con¬
tamination is the program, described in Sikorowski
(1975), that was used for several years in the R. T. Gast
Boll Weevil Rearing Facility. It included microbiological
sampling of air; larval diet; weevil eggs; equipment, walls,
floors, and surfaces; and adult weevils. What follows is a
description of this program.

Air microbial content.— Most of the important bacteria in
the food industry range from 2 to 4 /um long and 0.5 to
1.5 yum in diameter. So 100 liters of air were drawn
through a 0.45-/xm membrane filter supported in a filter
assembly. The filter was then placed in a 12- by 54-mm
plastic petri dish with an absorbent pad saturated with
mycological or plate-count broth and incubated at 35° C
for 24 hours. Afterward, the filters were removed from
the dishes, air-dried, stained with methylene blue solution
(0.5% methylene blue in 100% ethyl alcohol) for 30
seconds and redried. A 1- by 2-cm strip was cut out from
the central portion of the filter, placed on the slide, and
examined with a microscope at X 100. (Runkle and
Phillips 1969, Dark and Harper 1972, and others have
described techniques used in monitoring air microbial
content.)

Sterility of larval diet.— Twenty-ml samples of diet were
collected aseptically in sterile petri dishes every 15 min¬
utes during the operation time. The dishes were examined
after 24-48 hours incubation at 37° C.

Sterility of eggs.— From every 2,000-ml bottle of eggs
(150 ml of eggs +l,750-ml of egg-implanting solution), 5
samples of 2,000 eggs were implanted in each petri dish
with sterile trypticase soy agar, then incubated for 72
hours at 37° C. A Gas Pak Anaerobic System (Baltimore
Biological Laboratory, Cockeysville, Md.) was used for
isolation of anaerobic bacteria. A dish with one or more
colonies indicated egg contamination.

Micro-organisms on surfaces. — Two methods were used
for detection of bacteria on surfaces: the swab-rinse
method and the agar-contact method. The swab-rinse
method was originally developed by Manheimer and
Ysanez in 1917. It has had many modifications and is
now widely used. A cotton-tipped sterile swab is rubbed
over the surface of the object to be sampled. The tip of
the swab is broken into a tube containing sterile dilutant,
the tube is shaken, and the rinse fluid is plated on an ap¬
propriate culture medium. Care is taken to avoid con¬
taminating the swab. Patterson (1971) summarized the
major disadvantages of this method: there is often poor
recovery of bacteria from the surfaces sampled, results
are not reproducible between different workers or labora¬
tories, and cotton retains micro-organisms.

The agar-contact method uses a Rodac plate, a modified
petri dish that contains a raised nutrient-agar bed. The
plate is placed in contact with the tested surface, and the
samples are usually incubated for 48 hours at 37° C be¬
fore the colonies are counted. This method monitors sur¬
face cleanliness. Trypticase soy agar with lecithin and
polysorbate 80 medium is prepared for detection of sur¬
face contamination. Lecithin and polysorbate 80 inact¬
ivate residual disinfectant collected with the specimen.

The microbial contamination in the R. T. Gast Rearing
Facility varied with the area tested. In the egg-implanting
and larvae holding rooms, where air was filtered through
the HEPA filters, the bacterial count was less than 10
colonies per plate. The tests were made immediately after
the floor had dried following wet cleaning and before traf¬
fic was permitted.

Adult examination.— The criteria used by Sikorowski
(1975) to evaluate quality of adult boll weevils were
amounts of bacteria, protozoa, viruses, and yeast.

1. Bacteria.— A sample of 20 adult weevils was frozen
(—20° C) for 15-20 minutes, surface-sterilized with
0.5% sodium hypochlorite solution for 5 minutes, and
washed in 2 changes of sterile water (5 minutes each);
each weevil was blended in 40 ml of sterile water for
30 seconds. Forty ml of the homogenate was mixed
with 30 ml of melted agar (45° C) and poured into petri
dishes. After incubation, the colonies were counted.

2. Protozoa (Microsporidia).— A sample of 25 adult
weevils was blended in 25 ml of water. The suspension
was then filtered through cheesecloth and centrifuged
at 3,000 revolutions per minute for 10 minutes. The
resulting pellet was resuspended in a small quantity of
water, smeared on slides, and examined under a phase-
contrast microscope. (For more information about

149

diagnostic techniques used in identification of insect-
pathogenic protozoa see Hazard 1975 and Poinar and
Thomas 1978.)

3. Viruses.— If a virus disease was suspected, the host
tissue was examined with a light or electron
microscope as required. Insect viruses can be divided
into two broad groups: occluded viruses (nuclear and
cytoplasmic polyhedroses, the granuloses, and the in¬
sect pox diseases); and free viruses, which are not
embedded in a crystalline matrix. Inclusion bodies of
the occluded viruses can be easily detected with light
or phase-contrast microscopes. For detection of nonoc-
cluded viruses, an electron microscope must be used.
M. E. Martignoni (personal communication) has sug¬
gested that an infectivity test would be a more general
and probably cheaper method for detection of nonoc-
cluded viruses.

4. Yeasts.— If a yeast contamination was suspected, the
methods given by Poinar and Thomas (1978) were used.

Control

of Microbial Contamination
Use of antimicrobials

Singh and House (1970b) reported that, to prevent mi¬
crobial contamination in synthetic diets of 50 different
phytophagous insects, chlortetracycline has been used in
19 diets, formaldehyde in 18, methyl p-hydroxybenzoate
in 58, sodium benzoate in 6, and sorbic acid and its salts
in 42. Singh and House (1970a, 1970b, 1970c) and Singh
and Bucher (1971) studied how 21 antimicrobials mixed
with synthetic diets (meridic, or chemically defined) af¬
fected development of Agria housei Shewell. Singh and
House (1970b) concluded from their work that the safe
level for each antimicrobial compound and insect should
be determined for each diet-insect combination. To min¬
imize decreases in insect quality, the smallest effective
amounts of an antimicrobial compound should be used in
the diet. Bell et al. (1981) incorporated Aureomycin
(chlortetracycline), benomyl,10 sorbic acid, and methyl
p-hydroxybenzoate, in various combinations, into
lepidopterous larval diets to inhibit bacteria and fungi.
They concluded that benomyl and sorbic acid combina¬
tions provide the best protection against microbial con¬
taminants in the specific diet tested. Bell et al. (1959)
studied how sorbic acid affects 66 molds, 32 yeasts, and
6 species of lactic acid bacteria and found that all grow in
the presence of 0.1% sorbic acid at pH 7.0. The molds
and yeasts are inhibited when pH is lowered to 4.5, but
not all bacteria are inhibited until it is lowered to 3.5.

'“Methyl l-(butylcarbamoyl)-2-benzimidazolecarbamate.

Greany et al. (1977) reported that high pupal mortality
experienced during laboratory rearing of Opius longi-
caudatus (Ashmead), a parasitoid of the Caribbean fruit
fly, Anastrepha suspensa (Loew), was mainly caused by
opportunistic pathogens, S. marcescens and P. aeru¬
ginosa. Their study showed that stress caused by
parasitism and high rearing temperatures contributed to
deaths caused by bacteria. They also presented evidence
that methenamine mandelate chemotherapy had no pro¬
phylactic effect; instead, potentially harmful side effects
(aberrant premating sounds from the fly), were caused by
incorporation of this antibiotic in A. suspensa larval
medium. Control was accomplished by upgrading the cul¬
tural conditions rather than by the use of antibiotics.

Chawla et al. (1967) described a surface treatment with
antimicrobials to control fungi on wheat germ diets used
for mass rearing the European pine shoot moth, Rhya-
cionia buoliana (Schiffermiiller), and the codling moth,
Carpocapsa pomonella (Linnaeus). Sikorowski et al. (1980)
described a method that called for mixing antimicrobials
with sand or corncob grit and depositing the mixture on
the surface of the growth medium to shield boll weevil
larvae from microbial contamination. The results of the
study showed that: antibiotics can be applied to the
medium to shield the larvae from microbial contamina¬
tion without detectably hindering boll weevil develop¬
ment, egg production, hatch, and pheromone production;
and the standard diet overlain with medicated sand
yields 86%-97% newly emerged weevils with less than
500 bacteria per weevil, while diet with unmedicated
sterile sand yields only 18%. Sikorowski et al. (1980)
believe that antibiotics should be used only under certain
unusual circumstances when sanitary measures are tem¬
porarily interrupted. Prolonged use of antibiotics may
lead to the selection of resistant strains of bacteria.

Upgrading existing

sanitary measures

When enforcement of sanitation programs was relaxed in
the R. T. Gast Rearing Facility, the cleanup procedure
deteriorated because personnel overlooked the basic sani¬
tary measures (Sikorowski 1975). Clearly, then, in selec¬
tion of insectary workers, one should be assured of the
ability of the individual to perform the operation. The
worker must also be able to do this work so that the
essential cleanliness is not compromised. The worker
must know contamination-control principles and be aware
of the consequences of each action or inaction.

The furniture and equipment selected for the insectary
should be chosen to minimize maintenance and cleaning.

A large selection of cleanroom furniture, used in food in¬
dustries, hospitals, etc., is being marketed today. Work
surfaces should be noncorrosive and resistant to heat,

150

moisture, and abrasion. Stainless steel, because of its
durability and resistance to most corrosive cleaning and
sanitizing agents, is most frequently used for surfaces
that contact diet. Consideration of sanitary requirements
in the design of the equipment results in easier cleaning
and sanitizing, thereby saving labor and materials.

Conclusions

Microbial contamination could be described as the occur¬
rence and persistence of microbes in an environment
where they are not wanted. Synthetic and semisynthetic
insect diets may provide growth media for an almost
infinite number of microbe species. The microbial con¬
taminants may have diversified effects on insects, de¬
pending on vigor, age, composition of medium, etc. For
example, a change in the host resistance to microbial in¬
fection, caused by stress, may raise a nonvirulent
bacterial contaminant to the level of a disease-producing
microbe. Among the more common problems associated
with microbial contamination are: high mortality of
young instars, prolonged developmental time, diminutive
pupae and adults, reduced pheromone production, re¬
duced amino- and fatty-acid synthesis, wide fluctuations in
the quality of insects, and direct pathological effects. An
environmental sanitation program, then, should eliminate
contamination that interferes with the rearing of healthy
insects. I see a trend toward more involvement of insect
pathologists in the field of insect rearing. The control of
microbial contaminants in insectaries often presents
problems for which the field of insect pathology offers
solutions.

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1955. An introduction to bacterial physiology. 404
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1971. Microbiological assessment of surfaces. J.

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1977. Quaternary ammonium compounds. In S. S.
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Phillips, G. B., and Runkle, R. S.

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180 pp. CRC Press, Cleveland, Ohio.

Poinar, G. O., Jr., and Thomas, G. M.

1978. Diagnostic manual for the identification of in¬
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1969. Microbial contamination control facilities. 198
pp. Van Nostrand Reinhold Co., New York.

Sikorowski, P. P.

1975. Microbiological monitoring in the boll weevil
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Sikorowski, P. P.; Kent, A. D.; Lindig. O. H.; Wiygul, G.;
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1980. Laboratory and insectary studies on the use
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Sikorowski, P. P., and Thompson, A. C.

1979. Effects of bacterial load on mass reared boll
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153

Section 5

Production, Use, and Quality
Testing in Insect Rearing

The intent of this section is to convey the state of the
science of insect rearing. The papers describe representa¬
tive rearing systems for production of insects to be used
for research, development, and biological control. We be¬
come aware of how complex it is to integrate the modern
technologies of genetics, engineering, microbiology, and
dietetics and to coordinate the principles of quality con¬
trol and management with basic insect biology. We bene¬
fit from scientists who have many years of experience in
rearing insects and who have firsthand knowledge of the
advances made in the systems they describe. Each paper
sets the priorities for maintaining the integrity of each
rearing program. And some authors suggest ways to im¬
prove rearing capability. The following recommendations
summarize the major problems and challenges addressed
by the authors of this section.

The control of bacteria and fungi, like a persistent hypha,
is common to every rearing system described in this sec¬
tion. Efforts to prevent, mitigate, eliminate, and manage
microbial contaminants require much of the resources
available to any rearing program. Contamination can
eradicate an insect colony and years of work. And, even
in less extreme cases, the subtle effects of sublethal
microbial infections on the development, behavior, and re¬
production of an insect may significantly impair its use
for any purpose. Efficient methods are needed to sample
for, and determine the effects of, sublethal infections.
And, just as important, management guidelines for pre¬
venting contamination in the first place must be de¬
veloped and documented.

Insects reared in the laboratory can differ significantly in
quality from those in the wild parent population. But
little is known about population dynamics and behavioral
responses of most laboratory-reared species; indeed, there
are relatively few published tests for deriving such infor¬
mation. Results of studies that use laboratory-reared in¬
sects without comparing their performance with their
wild counterparts must be interpreted narrowly. Because

various laboratory cultures are used worldwide in many
ways, standardized tests must be developed. And studies
must be done to provide reference descriptions of how
established insect colonies that have been reared with
standard techniques respond to those techniques. The
availability of uniform testing procedures and perform¬
ance standards would promote communication and co¬
operation among scientists working with the same
species, increase our understanding of the biology and
behavior of the species, and enhance our ability to moni¬
tor and manage flaws in our rearing systems that lower
the quality of insects produced.

The development of artificial diets has pioneered advance¬
ments in insect colonization. The renewed emphasis on
biological control of agricultural pests insures that dietet¬
ics will continue to be important in the development and
evolution of standardized rearing systems. There are
several crucial areas in dietetics that need support and in¬
vestigation. First, the quality of diet ingredients must be
more closely defined, and tests to monitor ingredient
quality should be available to the insectary manager. Sec¬
ond, the effects of various preparation procedures on diet
quality and stability must be described and documented.
Finally, large-scale rearing and use of many potentially
effective entomophagous arthropods is prohibited by cost
and difficulty of rearing both host and parasite. A con¬
centrated effort is needed to develop artificial diets for
entomophages; investigation of in vitro rearing systems
would be especially appropriate.

The advances in genetic manipulation, in mechanization
of rearing procedures, and in insectary design and man¬
agement testify to the creative ingenuity of scientists to
design and implement complex, cost-effective rearing
systems that safeguard insect quality and quantity and
the health and safety of insectary personnel. Developing
such systems is the challenge for those people involved in
production, use, and quality testing of laboratory-reared
insects.

Thomas M. ODell,
Research entomologist,
U.S. Forest Service

155

Production of the Gypsy Moth,
Lymantria dispar,

for Research and Biological Control

By Thomas M. ODell,1 Robert A. Bell,2 Victor C. Mastro,3
John A. Tanner,3 and Leonard F. Kennedy4

Introduction

The first attempt to rear the gypsy moth, Lymantria
dispar (Linnaeus), in the laboratory in the United States
occurred in 1868 and resulted in its escape and establish¬
ment in Medford, Mass. This notorious European forest
pest was brought to the United States from France by
Professor Leopold Trouvelot, an astronomer and natural¬
ist who was trying to breed a new silkworm (Medford
Mercury Press 1906). His experiment led to the creation
of a pest-control industry that has flourished since the
first major outbreak in 1889.

After escaping from a window of Trouvelot ’s basement
laboratory, the polyphagous gypsy moth found suitable
and ample food; within 10 years, the population was com¬
pletely stripping the leaves from large shade trees in
residential areas surrounding the release site and alarm¬
ing citizens of Medford (Medford Mercury Press 1906).
The outcry and devastation prompted State and local au¬
thorities to investigate the biology of the gypsy moth
and methods of control. At the same time, experimental
applications of insecticides were begun. In 1893, the
desire to test candidate pesticides during the winter led
to the first documented indoor rearing of the gypsy moth
since its introduction. Eggs were hatched “early in the
season by means of artificial heat,” and larvae were main¬
tained on lettuce until leaves of elm, apple, or willow
could be obtained (Forbush and Fernald 1896). Larvae
were confined in open trays by tanglefoot smeared
around the edges. Temperature and humidity were main¬

'Research entomologist, Center for Biological Control of North¬
eastern Forest Insects and Diseases, Northeastern Forest Ex¬
periment Station, Forest Service, U.S. Department of
Agriculture, 51 Mill Pond Rd., Hamden, Conn. 06514.

2Research leader, Otis Methods Development Center, Agricul¬
tural Research Service, U.S. Department of Agriculture, Bldg.
1398, Otis Air National Guard Base, Mass. 02542.

3Entomologist, Otis Methods Development Center, Animal and
Plant Health Inspection Service, U.S. Department of Agricul¬
ture, Bldg. 1398, Otis Air National Guard Base, Mass. 02542.

‘Supervisory biological laboratory technician, Otis Methods
Development Center, Animal and Plant Health Inspection
Service.

tained at “comfortable levels” within the limits of stand¬
ard heating systems. These early attempts to maintain
the gypsy moth in the laboratory for experimental pur¬
poses were hampered by its univoltine life cycle, and by
the lack of appropriate food during the late winter when
eggs could be hatched at room temperature. Never¬
theless, this style of rearing continued until the 1950’s
with little modification.

The gypsy moth is univoltine; eggs are laid in light-
brown seta-covered masses in July and August; embry-
onation is completed in about 6 weeks; after this, the
apparently fully developed larva begins its diapause. Egg
hatch occurs in late April or early May, usually in syn¬
chrony with budbreak of the red oak, Quercus rubra L.,
the gypsy moth’s major host in the northeastern United
States. Males have five larval instars, and females have
six; they range in size from about 3.5 mm in the first
stage to about 30 mm in the sixth. Pupation begins in
June and continues through late July. Adults eclose dur¬
ing July and August. Besides the limitations of the uni¬
voltine life cycle and diet, the large larval size and the
long 35- to 40-day development period have been major
obstacles to the development of a biologically sound and
economical rearing methodology.

Between 1957 and 1960, intensive studies to develop
laboratory techniques for rearing large numbers of gypsy
moths were carried out at the U.S. Forest Service’s
Northeastern Forest Experiment Station, Forest Insect
and Disease Laboratory, New Haven, Conn. (Lewis 1961).
Experiments were directed at determining the physical
and biological factors affecting the growth and survival
of the gypsy moth. The tests were conducted in walk-in
environmental chambers built specifically to meet test re¬
quirements. Nutrition was supplied by plants grown in a
greenhouse. As in the past, intraspecific variation in host-
plant quality placed limits on interpretation of results in¬
volving insect growth and survival. Nevertheless, the
tests provided direction for later studies.

Rachel Carson’s book “Silent Spring,” published in 1962,
significantly affected forest-insect research and increased
the demand for research on biological control, experimen¬
tal diets, and a predictable means for rearing the gypsy
moth. At first, attempts were made to maximize foliage
quality by applying solutions of rice polish, sugars, and

156

vitamins directly to the leaves. Although larval growth
and survival apparently increased, off-season rearing re¬
mained unpredictable.

Rollinson (1964) began the first attempt to develop a
uniform diet by modifying a wheat germ diet originally
developed for the spruce budworm (McMorran and
Grisdale 1963). This research ultimately led to the
development of a diet and rearing technique (ODell and
Rollinson 1966) that, with slight modification, is still be¬
ing used for small-scale laboratory rearing. At the same
time, a diet was developed at the Connecticut Agricul¬
tural Experiment Station, New Haven (Leonard and
Doane 1966), and was used in studies on the development
and behavior of the gypsy moth (reviewed by Leonard
1974). Other diet formulations for rearing the gypsy moth
were later developed by Magnoler (1970), Vasiljevic and
Injac (1971), and Ridet (1972).

In 1965, the U.S. Animal and Plant Health Inspection
Service (APHIS) Gypsy Moth Methods Development
Laboratory (GMMDL) at Otis Air National Guard Base,
Mass., began studying ways to develop large-scale rear¬
ing of the gypsy moth to provide insects for research on
the sterile-male technique, pheromones, the trapping of
male moths for survey and detection, parasite production,
and new insecticides. Several improvements were made
(Secrest and Collier 1967, Tardif 1975), but reducing rear¬
ing costs was emphasized, and little attention was given
to product quality. Although procedures to prevent
microbial contamination were instituted, no internal
standards were developed for measuring colony quality.

In 1975, with support from the U.S. Department of
Agriculture’s (USDA) Expanded Gypsy Moth Program, a
USD A interagency team began investigating how to im¬
prove mass-rearing technology for the gypsy moth. The
main stimulus for developing a large-scale rearing
capability at that time was the potential of a virus
specific to the gypsy moth; this virus is now registered
under the name GYPCHEK. The goal of the team was to
develop the best possible system for mass rearing,
specifically: a low-cost diet and containerization, methods
for handling insects, procedures for control of diseases,
the best possible rearing environment, quality-control
procedures, procedures for controlling the health and
safety of workers, and pilot-scale facilities for insect and
virus production. The accomplishments of this program
through 1977 have been reviewed by Bell et al. (1981).

The rapid improvement of the mass-rearing capability by
1977 was directly responsible for reevaluation of the
sterile-male technique. It has also been largely re¬
sponsible for the success of several other studies that re¬
quire large numbers of uniform, healthy insects. Because
this operation is so important to the pest-management
program for gypsy moths, this paper will concentrate on

describing production, use, and quality testing now done
at the APHIS GMMDL.

Colony Establishment
and Maintenance

Facility

The facility began as several connected mobile homes in a
large brick warehouse; these have been replaced by spe¬
cially designed units built in the same space. The virus-
production facility is next to and connected with the
rearing area; traffic is routed for maximum isolation.
Research laboratories, walk-in environmental chambers, a
well-equipped shop, a large general work area, and offices
are located next to the central rearing area. The rearing
system is a prototype for facilities that mass-rear gypsy
moths; so it functions mainly as a methods-development
laboratory. At peak operation, the system is capable of
producing about 25,000 insects a day.

Colony selection

Colony selection began 112 years ago when the gypsy
moth was first introduced from France. The present
North American strain was derived from the relatively
few individuals that escaped in Medford, Mass. Accord¬
ing to Mayr’s founder effect (Mayr 1942), the initial
laboratory stock would have been derived from a small
amount of field material that was probably not genet¬
ically identical to the parent French population and
contained only a small part of the original genetic
variability.

In 1975, when the intensive research began on rearing of
gypsy moths, several laboratory and wild strains were
evaluated as potential breeding material for a standard
colony. The evaluation considered the rate of population
increase, the incidence of disease, and the rate of develop¬
ment. The laboratory strain selected and currently main¬
tained as the standard colony was originally colonized
from eggs collected in the winter of 1967 in an area near
Blairstown, N.J. At that time, eggs were collected be¬
cause there were many easily collectible egg masses; the
collection was not intended as a starter colony for future
production and was therefore not evaluated as potential
breeding material. The decision to continue the strain
was made on the basis of the question: “Why not?” The
egg masses were relatively small, and many of the
neonates died, apparently from virus. Disease, which was
present from the beginning, continued to destroy part of
each generation despite rigorous attempts to maintain
and culture under sterile conditions. Resistance to disease
was probably the strongest selective pressure on the col¬
ony. Researchers saved about 5% of each generation for
colony production. The largest and most lively pupae

157

were chosen by eye; these characteristics were also used
in selecting adults for mating. Eventually (generation not
known), one could expect viable egg masses from 80% of
the mated colony females; the starter colony had only
25%-40%. The decision to use the NJSS (New Jersey
standard strain) as the standard was dictated by the
evaluation procedure (and the colony’s apparent improve¬
ment) and by the fact that this particular strain had been
used by APHIS cooperators for many years. The decision
was based on the strain’s present performance not its
past poor quality.

Colony maintenance

and quality control

Techniques for colony maintenance and quality control
are still developing as rearing procedures and sampling
techniques are improved. In the system now in use,

10,000 neonates are implanted on the diet each week to
maintain the colony and to accommodate emergency re¬
quests from cooperators. Neonates are transferred to the
diet by hand with a soft camel’s-hair brush. Even though
this part of the rearing operation is not mechanized, there
has been no problem in meeting colony or production re¬
quirements; an average worker can transfer about 1,500
neonates per hour. Rearing containers (XE-6, Sweetheart
Plastics, Wilmington, Mass.), which contain 80 ml of a
diet high in wheat germ (see table 1 for recipe) and eight
neonates, are placed on plastic bread trays on aluminum
carts. Each cart has 14 shelves holding 28 trays and 840
containers. The 5-inch space between the top of the con¬
tainer and the tray provides critical air circulation.

Airflow in the rearing room is distributed downward in a
vertical laminar-flow pattern with a complete clean-air ex¬
change— HEPA (high-efficiency particulate air) filtered,
95% DOP (dioctyl phthalate aerosol penetration)
rating— every 3-5 minutes. At present, colony insects are
reared in the same room as production insects; tempera¬
ture is maintained at 25° C; relative humidity at 55%-
60%; and the photoperiod at 16 hours of light, 8 hours of
dark, with photophase beginning at 0500 hours. Environ¬
mental conditions in the rearing room are monitored con¬
tinually by a wet-dry bulb recorder read from outside the
room and by hygrothermographs located strategically in
the room. Recorders are checked once daily to make sure
that equipment is functioning properly. This monitoring
helps to keep systems running smoothly and to minimize
expensive equipment repair.

On artificial diet, development from newly hatched larva
to pupa takes about 30 days (male, 29-30 days; female,
30-31 days). Containers are sampled at 7, 14, and 21
days postimplant date (PID) for synchrony of develop¬
ment and for survival. These measurements are compared
on graphs to monitor colony performance, and deviations

Table 1.— Composition of diet used for rearing gypsy moths1

Grams

per

Ingredient Source liter

Wheat germ, raw .... Mennel Milling Co. 120

Fostoria, Ohio 44820.

Casein . New Zealand Milk Products 25

Rosemont, Ill. 60013.

Salt mix, ICN Biochemicals, 8

Wesson. Cleveland, Ohio 44128.

Sorbic acid . Dirigo Corporation 2

South Boston, Mass. 02127.

Methylparaben Tenneco Chemical 1

(methyl Piscataway, N.J. 08854.

p-hydroxy-
benzoate).

Vitamin premix Roche Chemicals 10

No. 26862. Division of Hoffman-LaRoche

Nutley, N.J. 07110.

Agar2 . Moorehead and Co . 15

Water . 800

‘In 1980, the cost per liter=$0.33; with HWG=$10.24.

2Gelcarin HWG (Marine Colloids) at 1% (10 g/liter) may be used
as an agar substitute.

from normal patterns are reported to the insectary
management for investigation. Pupae are harvested at
day 35 PIC, when about 80% of the insects have
pupated. Two hundred each of the largest male and
female pupae are selected by eye for colony production.
These are segregated by sex in 16-ounce, unwaxed Dixie
cups (50 males or 25 females per cup) and held in the
rearing chamber until eclosion 10-12 days later.

The newly eclosed males and females (25 of each per car¬
ton) are placed in unwaxed ice-cream cartons lined with a
layer of brown wrapping paper. Mating and oviposition
usually occur within 24 hours; but, to insure completion
of egg deposition, egg masses are not harvested for 72
hours. To allow embryonation, egg masses are held 25
days in the rearing room (25° C) before being transferred
to a walk-in refrigerator. In refrigeration, they are main¬
tained at 6° C, 90% relative humidity, and a 16 : 8 (hour)
light : dark cycle for 120-180 days.

Colony performance for each implantation group (all con¬
tainers in which neonates were implanted on any partic¬
ular day) is measured in survival from neonate to pupa,
sex ratio, mean weight (by sex) of 50 pupae harvested at
day 35 PID, proportion of adults eclosing, and incidence
and degree of adult deformity. Fecundity, embryonation,
and hatchability are primary indexes for determining the
rate of increase as a measure of population performance.
Measurements include mean egg-mass weight, mean eggs

158

per mass, percentage of embryonation, and percentage of
hatch. Standards are being established as sampling
techniques are refined. Current sampling methods are
relatively crude but allow for batch-to-batch com¬
parisons— sustained change from normal fluctuations of
any particular measure indicates a possible breakdown in
the rearing system and requires investigation.

Thirty egg masses (about 700 eggs per mass) are re¬
moved from cold storage each week for colony produc¬
tion. Egg masses are disinfected, intact, in a 10%
Formalin (formaldehyde) solution (see Shapiro 1977).
Compared to a cleaning of dehaired egg masses with
sodium hypochlorite, treatment of intact egg masses with
Formalin significantly reduced egg mortality and the in¬
cidence of virus in young larvae (ODell and Rollinson
1966). After the Formalin treatment, the intact egg
masses are placed in 100- by 25-mm plastic petri dishes,
three masses per dish, and held in a controlled-humidity
cabinet at 80% relative humidity and 25° C. Under these
conditions, eclosion of eggs chilled for 180 days begins
after the second day of incubation and continues for 3
days. Neonates are placed on the artificial diet after most
of the population has hatched. In this procedure, larvae
begin feeding quickly, and a 1 : 1 sex ratio is usually ob¬
tained at pupal harvest.

Recent investigations have focused on identifying critical
factors in the rearing system that affect variation in in¬
sect development, size, and survival: once identified,
these critical factors can be manipulated to stabilize nor¬
mal variation, which hinders program objectives. Critical
factors are also good places to start searching for causes
of unsatisfactory colony performance. Some examples of
critical factors and their treatment follow.

1. Elimination of microbial and fungal contamination sig¬
nificantly reduces mortality and enhances growth and
synchrony of development. Primary treatment is the
disinfection of eggs with Formalin. Additional treat¬
ment includes augmentation of the diet with microbial-
inhibiting additives (as is done in the diet shown in
table 1); provision of shoe covers and lab coats for per¬
sonnel; disinfection of work surfaces, carts, and trays
with Oxine (Biocide Chemical Company, Norman,
Okla.); and daily misting of the main insect-handling
room with Oxine. Airborne contaminants are mon¬
itored by standard agar plates (Trypticase soy agar for
bacteria and Sabouroud dextrose agar for fungus).
These are placed, 1 day per week, in each room of the
rearing facility, and daily in the larval implantation
room while neonates are being placed on the diet.

Plates are incubated for 2 days before being checked.

If bacteria or fungal colonies appear, appropriate steps
are taken to determine the inoculum source. Colony
and production insects that may have been exposed

are monitored more closely; significant increases in
diet contamination or an increase in larval mortality
may require that all insects from that implantation
date be destroyed.

2. Regulation of moisture in the rearing container is
critical to maintaining the best possible environment
for the larvae. And excessive moisture will increase
frass wetness, providing an excellent substrate for
microbial growth and creating a mat that at times in¬
hibits larval feeding. Treatment includes careful weigh¬
ing and measuring of diet ingredients, precision in diet
preparation time and in the volume of diet dispensed,
and implantation of larvae within 2 to 4 hours after
diet preparation. Variation in the volume of diet dis¬
pensed is monitored by randomly selecting and weigh¬
ing five containers per day. Moisture is controlled
mainly by having a container that provides adequate
ventilation though only the lid is permeable. This con¬
tainer regulates respiration, rate of food depletion, and
the amount of accumulated frass by controlling insect
density— ideally eight larvae per container. Having ap¬
propriate humidity control in the holding room and
maintaining an airflow across the top of the container
are also essential.

3. Subtle changes in the quality of diet ingredients may
be expressed by variation in rate and synchrony of lar¬
val development, pupal weight, and insect survival. To
minimize such variation, 52 batches of diet ingredients
are weighed and measured from the same stock of
fresh ingredients and then stored at — 15° C for use in
colony maintenance throughout the year.

Personnel health and safety

The gypsy moth is notorious for its urticating setae and
abundance of scales. Etkind (1976) tested the allergeni¬
city of the gypsy moth and found that the antigens of
cast larval skins and whole larvae caused positive im¬
mune reactions. Egg-mass hairs, though eliciting a signif¬
icantly lower immune reaction, are easily airborne and
cause sneezing and itching. These observations suggest
that egg-mass hairs are a strong primary irritant. The
main reason for installing HE PA filters in all units of the
APHIS GMMDL rearing facility was to provide con¬
tinuous cleaning of the air to remove insect hairs and
scales and so reduce potential health problems associated
with handling large numbers of gypsy moths. Also, work
stations are equipped with exhaust hoods and vacuum
systems to remove airborne hairs and scales and cast
skins. In areas where adult moths and egg masses are
handled, work stations are equipped with high-velocity
exhaust ducts to remove the irritating scales and egg-
mass hairs. Workers routinely wear protective clothing
and gauze masks or respirators to further reduce poten-

159

tial health problems. A centrally located safety station is
equipped with first-aid equipment for emergency and
routine health care. And a regularly checked bulletin
board at the safety station has space for management
and staff safety suggestions.

Production and Use

Although procedures for production and colony mainte¬
nance are similar, production quality necessarily deviates
from colony quality as levels of production increase. For
example, at peak production in 1979, the amount of dis¬
pensed diet per cup varied by as much as ±15%. The
asynchrony in larval development caused by such varia¬
tion was compensated for by increasing production and
changing the time of pupal harvest. How the product will
be used may allow other changes in the rearing system
without significantly reducing product quality. For ex¬
ample, larval density during production for virus can be
maintained at 10 per container (instead of 8) because
deaths occur before the end of the feeding period and
before food and available space are depleted. General pro¬
cedures have been established for monitoring and con¬
trolling quality during production. For each specific use
of the gypsy moth, quality traits are being identified, pro¬
cedures implemented during insect development to
monitor insect performance, and procedures established
for systematically investigating the rearing system to
determine causes for any deviation from standards.

Virus production

The nuclear polyhedrosis virus of the gypsy moth is host
specific, naturally occurring, and a main cause of death in
the gypsy moth during its outbreaks. In the laboratory,
the nuclear polyhedrosis virus is produced from
laboratory-reared larvae that have been hatched from
disinfected eggs and reared in sanitary conditions on an
artificial diet (Lewis 1976). In 1979, after 5 years of inten¬
sive investigation, laboratory-produced nuclear polyhe¬
drosis virus was registered by the U.S. Forest Service
under the trade name GYPCHEK.

Safety guidelines established by the U.S. Environmental
Protection Agency (1975) for virus preparations for con¬
trol of insects directly affect the production and quality
control of the insects to be used for virus production. The
guidelines are: “The fungal and bacterial contamination
of the technical virus due to production procedures must
not exceed the limits which can be maintained under
sanitary manufacturing conditions. More importantly, hu¬
man and mammalian pathogens (e.g. Salmonella, Shigella,
and Vibrio ) and microorganisms producing toxins known
to affect mammals must be absent from the technical
products.”

The degree of infectivity of the nuclear polyhedrosis virus
varies with the number of viable rods per polyhedra. The
activity of the nuclear polyhedrosis virus product is cal¬
culated by a standard bioassay procedure that is part of
the registration package for GYPCHEK (Lewis and
Rollinson 1978) and must be within ±20% of the limit
described in the standard bioassay. The NJSS was first
used for virus production in 1977-78 when about 5,000
acre-equivalents (quantity required to spray 1 acre) were
produced. During the winter of 1978-79, about 9,000
acre-equivalents were produced in the first attempt to
mass-rear gypsy moths for virus production.

Quality control during rearing of insects for virus produc¬
tion.— Rearing of moths for virus production is similar to
rearing for colony maintenance except that 10 larvae are
put in each cup instead of 8. A standard minimum weight
range for female larvae is established for 18 days PID,
the time of virus challenge. The standard is based on pre¬
vious production weights that produced maximum yields
of polyhedral inclusion bodies per gram and low bacterial
counts. Weights below the standard indicate a problem in
the production system and initiate investigation. But pro¬
duction continues unless bacteria or fungal counts exceed
standards or unless the diet quality has changed.

Sanitary conditions are monitored constantly to insure
that fungal and bacterial contamination is minimal.

Quality control during virus production. — At 18 days
PID, larvae are transferred from the main production
facility to the virus production area. They are then han¬
dled by separate personnel to minimize contamination of
the production facility. After the virus is sprayed on the
diet in each rearing container, all insects are transferred
to an environmental chamber maintained at 29° C. Four¬
teen days after inoculation, the diseased larvae are frozen
for 24 hours at —15° C. The frozen larvae are then freeze-
dried, dehaired, and ground. The processed product is
tested for quality by the following measures: (1) the
number of polyhedral inclusion bodies per gram is deter¬
mined for each implantation date; (2) bacterial and fungal
contamination is determined by procedures described by
Podgwaite and Bruen (1978); (3) each batch is bioassayed
and the data evaluated by Berkson’s logit chi-square
method (Paschke et al. 1968); and (4) each batch is intra-
peritoneally injected into 10 white mice. If any of the
mice die, the lot is rejected because the batch is contam¬
inated with organisms not detectable by normal means.
This is the primary quality-control test.

When contamination reduces virus quality, standard
microbial techniques are used to identify and eliminate
the source, and procedures are implemented to reduce
re-establishment. Methods are being developed for inves¬
tigating reduction in quality due to insect size and sus-

%

160

ceptibility to nuclear polyhedrosis virus. Presently, data
derived from production control are analyzed for signs
of breakdown in the system. The analysis determines
whether further investigation is warranted.

Factors affecting virus quality.— Human error, which
enhances microbial and fungal contamination during pro¬
duction, virus challenge, and harvest, is the main cause of
poor virus quality. Airborne contamination caused by
poor sanitary conditions is the result of human error, as
are poor harvesting conditions, which increase microbial
and fungal contamination in dead insects. Deviation from
production procedures is likely to damage virus quality.
For example, increasing insect density per cup or changing
environmental conditions would likely increase variation
in insect development and decrease polyhedral inclusion
bodies per gram; a change in diet composition (for ex¬
ample, a reduction in vitamin C concentration) might in¬
crease insect susceptibility to bacterial contamination,
producing more contamination in the final product. Quali¬
ty of the virus may be measured indirectly by gypsy
moth susceptibility. The degree of susceptibility to NJSS-
produced virus depends on the geographic distribution,
age, and physiological condition of the host, species of
the host’s food plant, and climatic conditions.

Sterile-male production

Use of the sterile-male technique for the gypsy moth was
proposed as early as 1952 by P. A. Godwin. Studies of
how gamma irradiation and chemical sterilants affect the
gypsy moth male began in 1957 and continued sporad¬
ically through 1978 (reviewed by Mastro et al. 1980).
Although gamma irradiation effectively sterilizes the
adult male, implementation of the sterile-male technique
for gypsy moth was inhibited by the inability to rear
large numbers of the insect. At first, Richerson and Cam¬
eron (1974) and Richerson (1976) reported that certain
laboratory-reared gypsy moth males, irradiated and non-
irradiated, did not fly as well and were not as sexually
aggressive as wild males. As these investigations used
adults reared from both field-collected and NJSS eggs
(generation unknown), it is difficult to speculate on the
factors that contributed to the moth’s apparent debility.

In 1976, because the gypsy moth rearing capability at
the APHIS GMMDL had been improved (Bell et al. 1981;
LaChance 1976), the USDA Expanded Gypsy Moth Re¬
search and Development Program supported new investi¬
gations of the sterile-male technique. These studies were
to find out whether sterilized males could be used to
suppress and eliminate newly established or isolated low-
density populations of gypsy moth. First, the competi¬
tiveness of the nonirradiated NJSS adult male was evalu¬
ated. Then, the effects of irradiation and use— packaging,
shipping, and distribution— were to be evaluated to pre¬

vent confusion about the causes of any debility. Several
operations (colonization, production, irradiation, and use)
that might individually or collectively affect performance
have been or are currently being evaluated. To date, only
the effects of colonization and production have been ade¬
quately studied.

LaChance (1976) identified four performance factors rele¬
vant to the mass-reared gypsy moth’s ability to compete
with wild males: the ability to locate females in various
situations and densities, the distance of dispersion,
spatial and temporal activity patterns, and mating suc¬
cess. These factors are being used to assess the competi¬
tiveness of the NJSS male. They are also being used to
establish performance indices based on measurements
that can be done directly, in the field and in the labora¬
tory. (See also Chambers 1977 for a discussion of general
problems of quality control in mass rearing and for a re¬
view of behavioral deficiencies often found in programs
for sterile-insect release.)

Quality control during production of insects for sterile-
male release.— Insects being reared for sterilization are
sampled at 31 days PID to determine percentage of pupa¬
tion and synchrony of larval development. An indication
that normal variation of either is being exceeded initiates
investigation and allows advance warning to field crews
applying the sterile-male technique that the production
schedule may not be met. Abnormal performance also
means that adults should be retested (see below) a‘t the
same time that more sterile males are released.

Quality control of adults in sterile-male production. — To

assess competitiveness, tests requiring field release and
recapture are conducted in forest areas with traps baited
either with the pheromone (+)— disparlure6 or with
females. The traps are arrayed vertically and horizontally
in 70- and 140-m-radius circles around a central release
point. The release point and trap sites are monitored
hourly or continually. This monitoring provides quantita¬
tive data on periodicity of eclosion and dispersal, on
spatial and temporal activity, on the ability to find mates
at various sites, and on the ability and propensity to
disperse. Data on survival and age of captured insects are
obtained by monitoring trap sites for several days after
release.

Onsite meteorological equipment and nearby weather
stations provide data for establishing field-performance
indices based on average temperature, humidity, and
windspeed. Performance indices are also being used to
evaluate laboratory techniques for testing quality; for ex-

5cis-7,8-Epox.y-2-methyloctadecane.

161

ample, demonstrating that a trait behaves the same way
in both natural and controlled environments would in¬
dicate that direct measurements of that trait could be
made in the laboratory.

At the APHIS GMMDL, quality testing is conducted
with an actograph and flight tunnel (modifications of
those described by Leppla and Spangler 1971— acto¬
graph— and Carde and Hagaman 1979— flight tunnel).
Waldvogel (1980) found that the NJSS (F16) preflight and
in-flight response to (+)— disparlure was similar to that
of wild males. The flight tunnel is now being used to pro¬
file preflight behavior in the presence of various concen¬
trations of (+)— disparlure and at several temperatures.
Field observations and measurements of preflight be¬
havior of individual moths (NJSS and wild) and their
mating success will determine whether flight-tunnel
preflight measurements can be used to evaluate competi¬
tiveness as a mark of quality. The flight tunnel is also be¬
ing used to characterize how pheromones affect behavior
of males exposed as pupae or adults to various holding
procedures during or after production. This information
will provide more quality standards for analysis of the
critical factors.

In 1979, researchers began actograph analysis to char¬
acterize the flight periodicity of NJSS (F19) and wild
males. With lights on and off simulating natural timing
of sunrise and sunset and with constant temperature,
humidity, and air movement, preliminary tests with the
actograph indicate the same flight periodicity demon¬
strated in field release-recapture tests. With this success,
the actograph is being used to establish performance in¬
dices for flight periodicity at different temperatures,
humidities, and photoperiods. These performance indices
will allow a laboratory comparison of various laboratory-
reared and wild strains that can then be used to predict
field performance and provide the standards for discern¬
ing critical factors affecting flight periodicity.

Factors affecting adult quality in sterile-male production.
In 1977, 1978, and 1979, field tests of wild and NJSS
(Fie, Fl7, and Flg) males demonstrated comparable
performance in ability to locate females in various places
in low-density populations (a simulation of 5-10 females
per hectare), ability to disperse from the release point,
and in mating success (Mastro and ODell 1978). But peak
capture of NJSS males occurred about 2 hours later in
the day than that of their wild counterparts. Actograph
analysis of flight periodicity showed a similar asynchrony
of activity. Similar asynchronies between laboratory-reared
and wild male moths have inhibited other programs using
the sterile-male technique (Raulston et al. 1976, White et
al. 1977). So, such behavior is a defect in the NJSS male.
Actograph studies are currently being conducted to iden¬

tify colonizing or production factors that hamper peri¬
odicity of activity.

Parasite production

Since 1906, about 37 species of parasites of the gypsy
moth have been introduced into North America, but only
10 species have become established (Hoy 1976). In
evaluating possible reasons for establishment or non¬
establishment, Hoy (1976) discussed briefly the use of
laboratory propagation of parasites for inoculative release
and observed that “the practical problems of preventing
genetic deterioration during laboratory propagation have
. . . not been solved.” Today, it is still true that, though
laboratory research of behavior of gypsy moth parasites
is increasing, the “practical problems” of laboratory prop¬
agation remain relatively untouched and unresolved.

Mackauer (1976) and Chambers (1977) have reviewed how
laboratory production affects parasite performance; they
found that adaptability and reproductive capability are
primary measures of the quality necessary for the success
of programs to release entomophagous insects. But little
information is available on the biology and behavior of
these parasites in their natural environments. Lack of
such information limits characterizations of gypsy moth
parasites.

Quality control in parasite production.— The NJSS is be¬
ing used for production of gypsy moth parasites by the
New Jersey Department of Agriculture in Trenton, the
Connecticut Agricultural Experiment Station in Hamden,
the University of Massachusetts in Amherst, and the
Forest Insect and Disease Laboratory also in Hamden,
Conn. Each of these organizations has a unique rearing
system, so comparing their production methods and prod¬
ucts is difficult. But they all have problems with periodic,
sometimes disastrous contamination by bacteria and
fungi. Gypsy moth eggs are sterilized, and work surfaces
and equipment are routinely cleaned at each facility; but
inadequate environmental control, other facility
peculiarities, and the additional stress of parasitization
make control of diseases and fungi extremely difficult.

Each year, the New Jersey Department of Agriculture
produces about 2.5 million gypsy moths to use at various
life stages for parasite production. The NJSS is used for
about 30% of the production; the rest is produced from
gypsy moths reared from egg masses collected each year
from natural populations. Larval mortality and variation
in the rate of larval development preclude the use of
about 50% of all larvae hatched. Most of this loss is
caused by contamination, poor vigor, and the intrinsic
variation in field-collected populations. Although no
specific data are available, more parasites axe produced

162

from NJSS larvae than from larvae reared from field-
collected eggs.

In New Jersey, the gypsy moth is reared on one of three
diets, depending on which one most increases numbers of
parasites for the least money. For example, Rogas lyman-
triae Watanabe is produced in hosts reared on a modifi¬
cation of the diet described by ODell and Rollinson
(1966). The less expensive but more nutritious (for the
host— R. A. Bell, personal communication) high wheat
germ and modified hornworm diets (both produced by
BioServ, Trenton, N.J.) have been tried. But in both
cases, hosts die soon after oviposition by R. lymantriae,
without producing a parasite. The causes of the apparent
differences in survival of host and parasite are unknown
and are not now being investigated (R. Chianese, personal
communication).

The NJSS are shipped as pupae from the APHIS GM-
MDL to the Connecticut Agricultural Experiment Station
and to the Entomology Department at the University of
Massachusetts for production and study of Brachymeria
spp., pupal parasites of the gypsy moth. The use of hosts
reared in the same system (APHIS GMMDL) reduces
probable variation in performance, size, reproduction, suc¬
cess, survival and behavior, caused by different methods
of host rearing; so the two studies are complementary
(though the Brachymeria populations will have intrinsic
differences).

The Connecticut Agricultural Experiment Station also
uses NJSS larvae, reared from eggs on the modified horn-
worm diet, for producing Apanteles melanoscelus
Ratzeburg (R. Weseloh, personal communication). NJSS
is preferred over wild strains because there is significant¬
ly less mortality of parasitized hosts. But comparisons in
size, reproduction, success, survival, and behavior have
not been made between parasites produced on various
host strains (R. Weseloh, personal communication).

The Forest Insect and Disease Laboratory at New Haven
uses NJSS to produce several species of gypsy -moth
parasites for research. R. lymantriae is being investigated
to determine whether it can be established in natural
habitats. Adult and mummy weights, fecundity, and
mating behavior are being recorded to establish indices of
colony performance. Environmental conditions are being
monitored closely so that changes in behavior and in
population characteristics can be analyzed (W. E.

Wallner, personal communication).

Factors affecting parasite quality.— Specific changes in
behavior and in population characteristics in parasites
and other insects subjected to laboratory colonization
(Boiler 1972, Mackauer 1976, Chambers 1977) have not
been reported for parasites reared on NJSS. ODell and

Godwin (1979) demonstrated that NJSS could be used to
rear large numbers of Blepharipa pratensis Meigen. In a
concurrent study (unpublished), they found that the
puparial weight of two groups of B. pratensis— reared at
the same time, under the same environmental conditions,
and on hosts fed the same diet (ODell and Rollinson
1966), but on different gypsy moth strains, differed
significantly from each other. The difference depended on
the geographic location and the density of the population
that the host eggs were collected from. The puparial
weights of B. pratensis reared from NJSS were similar to
those of parasites reared on hosts from low-density
populations.

A few studies have been done of parasite rearing on gyp¬
sy moths other than the NJSS. For example, Shapiro
(1956) found that B. pratensis (=Sturmia scutellata
Robineau-Desvoidy) was heavier when it emerged from
gypsy moth from low-density populations. Greenblatt and
Barbosa (1980) had similar results with Coccygomimus
turionellae (L.), but they had different results with
Brachymeria intermedia (Nees), which were heavier when
reared in hosts from high-density populations. At the
Pennsylvania Biocontrol Laboratory (Bureau of Forestry,
Division of Forest Pest Management, Harrisburg), the
size and fecundity of adult parasites are monitored
routinely to insure that the parasite colony maintains a
specified degree of quality; generally, gypsy moth
parasites reared on natural hosts are larger and produce
more eggs than those, such as the greater wax moth,
Galleria mellonella (Linnaeus), reared on unnatural hosts
(R. A. Fusco, personal communication). Hoy (1975a,

1975b) pointed out the need for field evaluation of
laboratory parasites, and for more observational studies
on their behavior. Certainly, if standard laboratory
strains such as NJSS are to be used for research and
development, such studies are justified.

Production for research and development

Leppla (1979) summarized the value of laboratory-reared
insects for research, noting that they are the “white rats”
of the entomologist, being “uniform, relatively inexpen¬
sive, continuously available, and comparable to their
counterparts in nature.” At this time, the NJSS is not a
“white rat.” Bioassays and standards for detecting
changes in the NJSS that might occur because of rearing
modifications, microbial contamination, and genetic selec¬
tion are just now being established. Nevertheless, NJSS
is widely used for research and development.

In 1976, NJSS (Fu) larvae were evaluated for use in bio¬
assaying plant extracts for deterrent compounds
(Doskotch et al. 1977). Such use would allow NJSS to be
used during the winter when larvae from field-collected
eggs are not available. In the bioassay, the dry weight of

163

frass was used to estimate the amount of feeding and the
relative deterrency of various plant extracts. Third-stage
larvae were reared from NJSS and from field-collected
eggs by the method described by ODell and Rollinson
(1966). When tested, the NJSS third-stage larvae con¬
sumed twice as much of the control diet (extract from red
oaks) as was consumed by their wild counterparts; but
there was no significant difference in the proportional
decrease in feeding response to an added deterrent com¬
pound (extract from Liriodendron tulipifera L.). A later
bioassay of other leaf extracts produced similar results;
so we were confident that NJSS larvae could be used in
the bioassay without compromising project objectives (T.
M. ODell, unpublished data).

NJSS larvae reared from eggs on a modified ODell-
Rollinson diet, where antibiotics were omitted, are used
to bioassay formulations of Bacillus thuringiensis
(Berliner). The bioassay compares the LD60 of each test
formulation with that of a universal B. thuringiensis
standard (Dulmage et al. 1971). There are indications that
NJSS larvae reared from eggs that have been held under
refrigeration (4°-6° C) for extended periods of time (200
days or more) do not respond normally to the standard;
that is, there is inconsistency in the normal variation of
the standard LD60. To avoid this abnormal response, new
preparations are tested with larvae hatched from eggs
held under refrigeration for 175-185 days.

NJSS larvae are also being used to study how secondary
metabolites affect insect growth and survival and to
screen insecticides. In each case, comparisons are being
made with larvae reared from field-collected eggs (M. E.
Montgomery and W. McLane, personal communications).
NJSS adults are being used by researchers at Michigan
State University, East Lansing, to characterize the
locomotor activity and pheromone response of males in
various field situations. NJSS adults are also being used
by APHIS GMMDL scientists to evaluate pheromone-
trap designs. These scientists (Michigan State and
APHIS GMMDL) use the adult quality-control evalua¬
tions developed for NJSS sterile-male release.

Challenges

There have been many advances in production, use, and
quality control of the NJSS since its establishment in the
laboratory in 1967. But, in making these advances, we
have learned how much more we need to know to improve
the scientific production and use of the gypsy moth for
biological control and research. The following is a list of
the many challenges we still have facing us at the
APHIS GMMDL.

1. Determine and maintain optimal storage conditions
and shelf life for perishable dietary ingredients, par¬

ticularly wheat germ, casein, vitamins, and gelling
agents.

2. Standardize through automation the processing and
dispensing of diet (mainly to eliminate day-to-day
variation).

3. Automate implantation of eggs and larvae on diet,
and automate harvesting of pupae.

4. Simplify monitoring of developmental and reproduc¬
tive performance.

5. Determine how many insects can be successfully
reared per unit of space if the rearing system is fur¬
ther refined.

6. Determine what environmental conditions will foster
the best rate and synchrony of development (par¬
ticularly for egg hatch, pupation, and adult eclosion).

7. Develop a method for rearing mostly males for
sterile-male production and mostly females for virus
production. (The female produces more virus because
it is larger than the male.)

8. Develop and explore the use of nondiapause or short-
diapause genotypes to augment or replace the ex¬
isting diapause strain (to reduce or eliminate the
requirement for chilling and to shorten generation
time).

9. Develop bioassays, and establish standards for
evaluating insects for consistency of performance in
tests for standard toxicity and activity.

10. Develop systematic methods for investigating devia¬
tions in production standards and deterioration in
product quality.

11. Develop a communication system for informing users
of production and product change, and encourage the
establishment of internal standards for periodic
evaluation of the product received.

12. Develop an antiserum for desensitizing personnel
acutely allergic to gypsy moth setae and scales.

13. Investigate relative chilling effect of air movement
on eggs being held at various temperatures (0°-7° C)
and at 70%-100% relative humidity.

Acknowledgments

We thank Pedro Barbosa, Joseph Elkinton, Robert
Fusco, Franklin Lewis, and David Lance for their review
of this paper. We are also grateful to the staff of the
APHIS GMMDL for their timely suggestions, adherence
to quality-control procedures, and good humor, all of
which have contributed significantly to the advancement
of NJSS rearing technology.

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Research Toward Integrated Pest Manage¬
ment, pp. 669-679. U.S. Dep. Agric. Tech.
Bull. 1584.

Mayr, E.

1942. Systematics and the origin of species. 334 pp.
Columbia University Press, New York.

Medford Mercury Press.

1906. Illustrations of the ravages of the gypsy moth
and brown tail moth. Medford Mercury Press,
Medford, Mass.

ODell, T. M., and Godwin, P. A.

1979. Laboratory techniques for rearing Blepharipa
pratensis, a tachinid parasite of gypsy moth.
Ann. Entomol. Soc. Am. 72: 632-633.

ODell, T. M., and Rollinson, W. D.

1966. A technique for rearing the gypsy moth, Por-
thetria dispar (L.), on an artificial diet. J.

Econ. Entomol. 59: 741-742.

Paschke, J. D.; Lowe, R. E.; and Giese, R. L.

1968. Bioassay of the nucleopolyhedrosis and

granulosis viruses of Trichoplusia ni. J. In-
vertebr. Pathol. 10: 327-334.

Podgwaite, J. D., and Bruen, R. B.

1978. Procedures for the microbiological examina¬
tion of production batch preparations of the
nuclear polyhedrosis virus ( Baculovirus ) of
gypsy moth, Lymantria dispar L. U.S. For.
Serv. Gen. Tech. Rep. NE-38, 8 pp.

Raulston, J. R.; Graham, H. M.; Lingren, P. D.; and
Snow, J. W.

1976. Mating interaction of native and laboratory-
reared tobacco budworms released in cotton.
Environ. Entomol. 5: 195-198.

Richerson, J. V.

1976. Relative attractiveness of irradiated

laboratory-reared female gypsy moths and
non-irradiated laboratory-reared and feral
females. J. Econ. Entomol. 69: 621-622.

Richerson, J. V., and Cameron, E. A.

1974. Differences in pheromone release and sexual
behavior between lab-reared and wild gypsy
moth adults. Environ. Entomol. 3: 475-481.

Ridet, J. M.

1972. Etude des conditions optimales d’elevage et

d 'alimentation de Lymantria dispar L. Aim.
Soc. Entomol. Fr. 8: 653-668. [English sum¬
mary.]

Rollinson, W. D.

1964. An artificial diet for the gypsy moth. U.S.

For. Serv. Northeast. For. Exp. Stn. Tech.
Prog. Rep. Nov. 1963-Apr. 1964., pp. 7-8.
Secrest, J. P., and Collier, C. W.

1967. Mass rearing summary report (Jan.

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Animal and Plant Health Inspection Service,
Otis Methods Development Center, Bldg.

1398, Otis Air National Guard Base, Mass.
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Shapiro, V. A.

1956. The influence of the nutritional regimen of the
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[In Russian.]

Shapiro, M., and Bell, R. A.

1977. Gypsy moth mass rearing: evaluation of
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and/or pupae (Oct. 1976-Mar. 1977), pp.

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Otis Methods Development Center, Bldg.

1398, Otis Air National Guard Base, Mass.
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Tardif, J. G. R.

1975. Techniques for large-scale production of gyp¬
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U.S. Environmental Protection Agency.

1975. Guidance for safety testing of baculoviruses.
In M. Summers, R. Engler, L. Falcon, and P.
Vail (eds.), Baculoviruses for Insect Pest Con¬
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American Society of Microbiology,
Washington, D.C.

Vasiljevic, L. J., and Injac, M.

1971. Artificial diet for the gypsy moth (Lymantria
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Waldvogel, M. G.

1980. Evaluation of lab-reared irradiated, lab-reared
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22: 262-266.

166

Rearing the Tobacco Budworm,
Heliothis virescens, and the Corn
Earworm, Heliothis zea

By J. R. Raulston1 and E. G. King2

Introduction

Scientists doing entomological research with Heliothis
spp. have long recognized that laboratory colonies of
these insects are an important research instrument.
Ellisor (1935), having found himself without enough field-
reared com earworms, Heliothis zea (Boddie), to complete
certain laboratory studies, devised a way to rear them in
the greenhouse. His method required caging adult moths
on young corn plants to lay eggs and then transferring
third- and fourth-stage larvae to soaked sweet com
kernels for completion of larval development. Barber
(1936) described a culturing method that required placing
first-stage corn earworm larvae on the silk of corn ears
that had been silking for less than 10 days. Barber found
that larval development was completed in 2 to 3 weeks,
development periods that corresponded with those ob¬
served in the field. These early attempts to culture corn
earworm in the laboratory had drawbacks, and Stahler
(1939) described a disease that decimated populations be¬
ing reared on lettuce and alfalfa in the insectary. The
disease symptoms described indicate that the larvae were
dying from a nuclear polyhedrosis vims.

Methods similar to these were used for many years in
propagating corn earworm in the laboratory. As recently
as the early 1960’s, Callahan (1962) described techniques
and apparatus for rearing corn earworm larvae on corn
and for enhancing mating and oviposition by adults. But
he was unable to maintain the laboratory culture for more
than five generations and concluded that the diet lacked
certain ingredients needed for continuous culturing.

It became possible to rear Heliothis spp. under con¬
tinuous laboratory culture divorced from its natural hosts
after the nutritional requirements for various insect
species were elucidated and artificial diets were developed
(see reviews by House 1965 and Dadd 1970). Here, we

‘Research entomologist, Cotton Insects Research Unit, Agricul¬
tural Research Service, U.S. Department of Agriculture,
Brownsville, Tex. 78520, in cooperation with the Texas
Agricultural Experiment Station, Texas A&M University.

2Research entomologist, Southern Field Crop Insect Manage¬
ment Laboratory, Agricultural Research Service, U.S. Depart¬
ment of Agriculture, P.O. Box 225, Stoneville, Miss. 38776.

will review several subjects important to the laboratory
propagation of Heliothis spp.— diet development and
evaluation; diet preparation and handling; rearing equip¬
ment and techniques; facility design and flow patterns;
and how pathology, physiology, and behavior affect quali¬
ty control.

Diets

The first published attempt at rearing the com earworm
on an artificial diet was that of George et al. (1960) using
a diet similar to one developed by Bottger (1942) and
Beck et al. (1949) for the European corn borer, Ostrinia
nubilalis (Hiibner). George et al. (1960) were unable to
rear the corn earworm through its complete larval cycle
on their diet and had to feed the first-stage larvae on
corn silk. Vanderzant et al. (1962a) were able to rear the
com earworm from egg to adult on a purified diet, and
they determined a dietary requirement for ascorbic acid.
Because this diet had so many ingredients, Vanderzant et
al. (1962b) went on to rear the corn earworm on a wheat
germ diet modified from one developed by Adkisson et al.
(1960) for the pink bollworm. The wheat germ diet, which
contained fewer ingredients than the previous diet, was
adequate for normal larval development. Using as a base
diet the one she and others reported earlier (1962a),
Vanderzant (1968) later determined the lipid and sterol re¬
quirements of the com earworm. She also determined
that sterol, linoleic acid, linolenic acid, choline, and in¬
ositol were needed in the diet.

Based on these initial investigations and on the nutri¬
tional requirements established by Vanderzant and
others, several diets, many only slightly modified from
that reported by Vanderzant et al. (1962b), have been de¬
veloped for rearing Heliothis spp. But Shorey (1963)
developed a radically different diet based on lima beans
and brewer’s yeast for the cabbage looper, Trichoplusia ni
(Hiibner), and Shorey and Hale (1965) modified this diet
slightly for rearing nine noctuid species including the
com earworm. Burton (1969) further modified the bean
diet for rearing the corn earworm and tobacco budworm,
H. virescens (Fabricius), by adding wheat germ and
replacing brewer’s yeast with the less expensive torula
yeast. Patana (19g9) used baby lima beans in the bean
diet for rearing several lepidoptera, including the corn
earworm and tobacco budworm and reported larvae fed
more readily on this diet. Patana and McAda (1973) also

167

made dry flakes from the bean diet for use as a physical
barrier to separate feeding tobacco budworm larvae,
which are cannibalistic in the later instars.

Most of the recent diet modifications for rearing Helio¬
this spp. have emphasized reducing cost. Burton (1970)
reported the development of a diet for the com earworm
based on a child-food supplement, CSM (corn-soya-milk),
and Burton and Perkins (1972) developed a diet for rear¬
ing com earworm and tobacco budworm that is based on
a blended human-food product, WSB (wheat-soya blend).
Insect growth and development on these diets are com¬
parable to those on the bean diet but cost about half as
much. Perkins et al. (1973) made further comparisons of
the bean, CSM, and WSB diets with another diet contain¬
ing opaque-2 com and concluded that the WSB is nutri¬
tionally and economically best for rearing Heliothis spp.
Other researchers have modified the casein and wheat
germ diet reported by Vanderzant et al. (1962b) to reduce
its cost. For rearing tobacco budworm, (1971) Shaver and
Raulston replaced casein with a toasted, defatted
soyflour. Prepared with the agar-reduction method
reported by Raulston and Shaver (1970), this diet has
been about 50% cheaper than the casein-wheat germ diet.
For corn earworm, Brewer et al. (1975) concluded that
soyflour is better than cottonseed meal or corn germ flour
as a source of protein. Brewer and Martin (1976) also
reported using various sawdusts or corncob grits and
found them to be an economical means of reducing the
amount of agar used in the com earworm’s diet. Brewer
and King (1979) reported that the corn earworm requires
3.4 g of the soyflour diet containing corncob grits to com¬
plete larval development and that the tobacco budworm
needs 2.7 g. Brewer and Tidwell (1975) and Raulston
(1975a) also evaluated the effect of reducing the
B-vitamin supplement used in their diets and found it can
be reduced by as much as 20% with no harmful effects.
Generally, institutions or laboratories that maintain small
laboratory colonies of Heliothis spp. still rely on the orig¬
inal casein and wheat germ diet reported by Vanderzant
(1962b); however, most researchers doing large-scale rear¬
ing have been forced by high costs to develop new diets
or modify existing ones.

Diet Preparation
and Handling

Maintaining small laboratory colonies of Heliothis spp.
requires little sophistication in diet preparation and dis¬
pensing. In fact, both are often done with methods first
reported by Berger (1963), who prepared his diet in a
small blender and dispensed it into rearing containers
from squeeze bottles. Later, as demand increased for
more production, researchers devised equipment to facil¬
itate the preparation and handling of the diets. Burton et
al. (1966), Patana (1967), and Stadelbacher and Brewer

(1974) reported dispensing diet from containers under
pressure. Diet-preparation capacity was increased by the
use of large blenders— for example, the 38-liter Hobart
VCM 40E used by Burton (1969). And Raulston and Lin-
gren (1972) reported use of a stainless steel tank capable
of preparing 177 liters of diet. This system used an over¬
head Lightnen Mixer, Model NM-1 to mix the diet in¬
gredients rapidly. Raulston and Lingren (1972) dispensed
their diet with a peristaltic pump, but later they changed
to a positive-displacement pump like the one described by
Patana (1977). Harrell, Sparks, Hare, and Perkins (1974)
described a mechanized system for mixing large quan¬
tities of dry ingredients (225 kg) and a continuous sub¬
sequent flow of flash-sterilized diet to a form-fill-seal
machine modified for rearing com earworm larvae. This
system maximized production. Brewer (1977) agreed that
flash sterilization of the com earworm diet did not affect
larval growth and development, but found that anti¬
microbials were still necessary in the diet because of
recontamination.

Larval Rearing Methods

At first, larval-rearing techniques were labor-intensive
and evolved from nutritional studies where only a few in¬
sects were handled. Glass vials plugged with cotton were
commonly used by George et al. 1960; Brazzel et al. 1961;
Vanderzant et al. 1962a, 1962b; Berger 1963; Chauthani
and Adkisson 1965; and Vanderzant 1968). Diet was typ¬
ically dispensed into the vial from squeeze bottles, and
neonate larvae were placed on the gelled diet with a fine
brush.

Automated methods for dispensing the diet and handling
the vials were developed in response to increased demand
(Burton et al. 1966). These vials required cleaning and
sterilization for reuse; so disposable plastic cups were
substituted for the vials. Burton and Cox (1966) and Bur¬
ton and Harrell (1966) modified a foodpacking machine to
dispense the cup, dispense diet into the cup, and cap the
cup. Burton (1969) modified this procedure for corn ear-
worm rearing, but the neonate larvae had to be placed in
the cups by hand. Harrell et al. (1969) developed a ma¬
chine that crushed the cups, so pupae could be harvested,
then separated the pupae from the diet and crushed cups.
Harrell et al. (1970) described a device to glue com ear-
worm eggs to the cup lid, thus eliminating the need for
larval implantation. Use of these automated devices
allows processing of about 3,600 cups per hour. Other
nonautomated methods were developed that use plastic
cups for rearing Heliothis spp. Patana (1969) described
some of these procedures, which are often used for main¬
taining small laboratory colonies.

Shorey and Hale (1965) first used compartmentalization
for rearing Heliothis spp. larvae by fitting cardboard

168

dividers into 175-ml cups. Compartmentalization became
an accepted technique for rearing Heliothis spp. larvae
after Ignoffo and Boening (1970) reported rearing larvae

I in compartmented disposable plastic trays covered with
Mylar film. Ignoffo and Boening transferred neonate lar¬
vae into the containers with a brush before sealing the
unit; or they used the larval transfer system described by

IGard (1969) after the unit was sealed. E. A. Harrell and
others (Harrell et al. 1973; Harrell, Sparks, Hare, and
Perkins 1974; Harrell, Sparks, Perkins, and Hare 1974)
automated procedures for using the compartmented trays
for rearing corn earworm larvae by fitting an inline form-
fill-seal machine (that formed the trays) to a device for
dispensing diet and implanting eggs. Equipment was also
devised to collect the pupae from the trays. Sparks and
Harrell (1976) described the complete automated rearing
system and reported that at top speed the system could
process 261 of the 120-cell trays per 8-hour day.

Another system that diverged from the compartmental¬
ized trays was the use of cells that could be placed into
trays. Roberson and Noble (1968) first reported rearing
tobacco budworm larvae in honeycomb-like cells formed
from Mylar. Raulston and Lingren (1969) improved this
technique and later developed a rearing cell from Poly¬
styrene light-diffusion louvers that could be reused in¬
definitely (Raulston and Lingren 1972). These authors
also devised a simple technique for implanting eggs into
the cell unit. Other researchers have further modified the
cell-rearing technique by replacing the top made of
screen-polypropylene cloth and polyurethane foam
described by Raulston and Lingren (1972) with a single
sheet of 3.2-mm-thick polypropylene-Porex filter (Hartley
et al. 1982). This unit is now being used to mass-produce
the sterile backcross (BCnQ X tobacco budworm^) from the
intercross [tobacco budworm Q X H. subflexa (Guenee)cJ].
About 4 million BC pupae have been produced by this tech¬
nique over the last 3 years (1979-81); also in the late 1970’s,
during another pilot sterile-male release project involving
tobacco budworm pupae, a production level of about
60,000 pupae per day was sustained for 4 years at a cost
of less than $5.00 per 1,000 pupae.

Adult Handling

Various container types have been used as oviposition

I cages for Heliothis moths. Two of the most common con¬
tainers have been the 3.8-liter glass jar and the 3.8-liter
cardboard ice-cream carton. About 10 pairs of moths can
be maintained in these ice-cream cartons without affect¬
ing fecundity or fertility, but more (20 pairs per carton)
reduce fertility (Burton 1969). In each oviposition con¬
tainer, the top is replaced with a piece of cloth, and cloth
strips (streamers) are suspended from the container top’s
edge to the bottom. The cloth top and streamers serve as
oviposition substrates. The moths are fed diluted honey

or a sucrose solution. Callahan (1962) used plywood cages
measuring 25 by 25 by 30 cm as moth holding and ovi¬
position cages. The cage front was fitted with a cloth or
paper cover that served as an oviposition site. He con¬
cluded that four to six pairs of corn earworm moths are
best for optimum oviposition in this size of cage. His
studies showed that light, temperature, and humidity are
critical elements in obtaining maximum mating and ovi¬
position. In fact, no mating by corn earworm moths oc¬
curred when the moths were held at temperatures above
29.4° C.

Devices have been developed for continuous removal of
Heliothis spp. moth scales because they are a human
health hazard. Burton (1969) described methods for filter¬
ing scales from the work area while collecting oviposition
cloths. Raulston and Lingren (1972) developed emergence
cages for large-scale production that have an enclosed
scale-filtering system and a device for transferring adults
from emergence cages into the oviposition cages, mini¬
mizing handling.

Recently, Gross et al. (1975) and McWilliams et al. (1981)
have developed techniques for managing the moths in
large quantities with minimum handling. Gross et al.
(1975) described a plywood box partitioned vertically by
five waxed-paper sheets into four oviposition chambers.
Each chamber was fitted with a continuous flow of beer
as a diet source and provided oviposition sites for 100
pairs of adults. The waxed-paper sheets could be pulled
through the bottom of the chamber and the eggs brushed
off with a hog-bristle brush. McWilliams et al. (1981)
reported using oviposition cages that also include devices
for adult emergence and feeding. The cages are housed in
a chamber that confine and collect the moth scales. The
moths lay eggs on a standard aluminum-frame, fiber¬
glass-mesh screen that fits into grooves in the cage
framework and can be easily inserted or removed.
McWilliams has gotten sustained production of about 1.5
million eggs per week by operating 15 oviposition cages
needing about 5,500 pupae per week. The operation re¬
quires about 15 hours of labor per week.

Facilities

for Rearing Heliothis

The advances in dietary and technological developments
facilitated the rearing of Heliothis spp. and allowed their
continuous colonization in the laboratory. But many col¬
onies still fail; Sparks and Harrell (1976) cited both the
lack of facilities to maintain proper environment and the
inability to eliminate massive disease outbreaks as the
main causes of failing to increase production. So, however
sophisticated rearing technology has become, large-scale,
or even small-scale, production of Heliothis spp. will not
be uniform unless facilities are properly designed for such

169

19

A

I Air locks

Steam autoclave
Ethylene- oxide
sterilizer

BROOD COLONY:

1 Equipment room
2 Shower
3 Hallway

4 Media preparation
5 Larval holding
6 Pupal harvesting
7 Emergence

8 Oviposition

9 Egg sterilization
& infesting

I 0 Storage room
PRODUCTION AREA

II Entrance foyer

12 Media preparation

13 Storage

14 Sterile tray storage

15 Media curing

16 Pupal har vesting

17 Infesting

1 8 Rack storage

1 9 Rack wash

20 Hallway

21 Larval holding

2 2 Shower

Figure 1.— Floor plan and flow pattern of the U.S. Agricultural Research Service’s tobacco budworm
rearing facility, Brownsville, Tex.

rearing. (See Chambers 1978, specifically, and Leppla and
Ashley 1978, generally, for some of the requirements for
such rearing facilities.) Here, we will concentrate on the
design and operation of two facilities that have been suc¬
cessfully used for rearing tobacco budworm on a relative¬
ly large scale.

In the U.S. Agricultural Research Service’s rearing facili¬
ty at Brownsville, Tex., Raulston and Lingren (1972) used
a chilled-water environment control and relatively un¬
sophisticated conventional cleanroom technology as
described by Austin and Timmerman (1965). Use of
chilled-water environment control eliminated the transfer
of air between rooms and allowed the environment in
each room to be separately controlled. The system was
designed so that all rooms, except the pupal-harvest
room, were under positive pressure. All the recycling air
and the fresh air being introduced into each room passed

over the conditioning coils and through a HE PA (high-
efficiency particulate air) filter with a pore size of 0.3 gm
before entering the room. One-hour exposures of nutrient
agar plates in each of seven rooms in the stock-culture
area resulted in an average mold-colony growth of 0.16
per hour and an average bacterial-colony growth of 4.2
per hour. These data demonstrate how well the system
reduced airborne contaminants.

The system (fig. 1) was designed as two separate and
autonomous facilities comprising areas for stock culture
and large-scale production. The only junction between the
two areas was a passthrough box that allowed eggs to
pass from the stock culture into the mass-production
area. The one-way flow pattern was designed to move
both personnel and materials through the system from
clean to less-clean areas. The stock-culture area was main¬
tained under strict quarantine, and personnel were re-

170

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Equipment room

8

Diet sterilization

9

Entrance

10

Dressing room

1 1

Shower

12

Corridor

Pupae sexing

13

Harvest
Larval holding
Supply
Foyer

Diet dispensing
<£ egg infesting
Emergence & oviposition

Air locks

Figure 2.— Floor plan and flow pattern of brood-colony units for producing tobacco budworms and
sterile backcrosses at the U.S. Agricultural Research Service’s Delta States Research Center at
Stoneville, Miss.

quired to shower and change into clean garments before
entering the work area. The quarantine was placed on the
stock-culture area because it was involved in culture
propagation, and it was believed that only disease-free
cultures would result in uniformly high quality. Since the
production area produced only pupae for use outside the
facility, no adult-handling facilities were incorporated into
its design; however, rigid restrictions were still main¬
tained on movement of personnel into the facility and
through the work areas. The tobacco budworm culture in¬
troduced into this facility was thoroughly examined for
nuclear polyhedrosis virus, cytoplasmic virus, and
microsporidia before its introduction. The culture was
monitored weekly, and the colony was kept free of

disease from the time of its introduction in 1969 until the
facility was closed in 1977.

The second facility (fig. 2) was constructed for brood col¬
onies to produce sterile Heliothis hybrids at the U.S.
Agricultural Research Service’s Delta States Research
Center, Stoneville, Miss. (Brewer et al. 1978). This facility
was also designed to use a chilled-water environment con¬
trol and absolute air filtration for reducing microbial con¬
taminants. It is maintained under positive pressure
except for the harvest area, which is maintained under
negative pressure. The two sides of the facility are mirror
images. On one side, tobacco budworm has been produced
for culture maintenance; that side has also been used to

171

produce sterile males to be used in the other half of the
facility for backcrossing to Fn -progeny females for egg
production. Mass-production of sterile hybrids has then
been done in another facility. Located between the two
halves of the facility is the diet-preparation area, which is
not maintained as a cleanroom. After preparation, the
diet is flash-sterilized and pumped through a closed
system to the desired area. So a sterile diet kitchen is not
needed. Colonies being reared are maintained under
quarantine to prevent introduction of disease organisms.

Quality Control

The application of cleanroom techniques and the best
available environment-control systems has greatly fa¬
cilitated the production of Heliothis spp. in continuous
closed cultures. These advances and the automated and
highly efficient rearing procedures provide the technology
for mass producing Heliothis spp. as needed. Such
production efforts require that rigid standards be placed
on the product (the insect) and that a monitoring system
be established to insure that these standards be met. Sev¬
eral aspects of quality must be considered in the develop¬
ment of monitoring systems (see, for example Mackauer
1972, 1976; Huettel 1976; and Chambers 1977). Huettel
(1976) lists the major components associated with insect
propagation in the laboratory, their measurable traits,
and possible methods for monitoring such traits.

Both com earworm and tobacco budworm are susceptible
to several diseases that can rapidly become epidemic in
the confines of the laboratory. So one of the first needs of
quality control is monitoring for disease organisms. Three
of the major disease organisms associated with labora¬
tory propagation of Heliothis spp. are nuclear polyhedro-
sis vims, cytoplasmic polyhedrosis vims, and the proto¬
zoan Nosema heliothidis (Lutz and Splendor). Outbreaks
of nuclear polyhedrosis are normally self-limiting since
those insects that contract the disease die. Both the
cytoplasmic vims and the protozoan, however, are easily
transmitted across generations— cytoplasmic virus on the
egg’s surface (Mery and Dulmage 1975) and Nosema in
the egg (Brooks 1968). So these two diseases may cause a
general decline of the laboratory culture with prolonged
development, increased mortality, and decreased fecundi¬
ty and fertility. A disease-monitoring system should in¬
clude direct measurements on these physiological traits
and autopsies of suspect individuals by a competent
pathologist. The best method for handling disease in
laboratory colonies is prevention. In fact, disease
organisms in a colony are intolerable since the quality of
the product can no longer be guaranteed.

Quality insect rearing is dictated by intended use of the
product. So traits desirable for specific culture uses, as in
a laboratory bioassay or in a hybrid or sterile-insect

release, may not be consistent and may be mutually ex¬
clusive. H. T. Dulmage (personal communication) lists
desirable traits in a tobacco budworm culture used to
bioassay Bacillus thuringiensis Berliner isolates; these
are uniformity in larval growth, reproducibility of results
over time, and the culture’s capacity to reproduce enough
insects to meet bioassay demands uniformly and predict¬
ably. These traits would describe a culture that had
already undergone the dynamic period of laboratory adap¬
tation, was genetically stable and disease free and was
being reared on a uniform artificial diet under a uniform
set of laboratory conditions. But traits that would be con¬
sidered desirable in a culture of Heliothis spp. destined
for use in a sterile-male program would be much different.
Here, desirable traits would be ability to disperse within
the ecosystem, ability to locate and use host plants, and
ability to locate and compete for native virgin female
mates. Maintenance of these characteristics would require
developing a rearing regimen that would insure a wide
genetic base and minimize genetic changes caused by
laboratory adaptation.

Young et al. (1976) developed a crossing scheme to
minimize inbreeding in a culture of com earworm and
concluded that the method improved colony performance
in mating, fecundity, and fertility. No behavioral traits
were measured in this investigation. Raulston (1975b)
reported laboratory adaptation of a wild strain of tobacco
budworm and found that selection pressure changes the
mating and oviposition patterns beginning in the sixth
and seventh generations. Crosses between the laboratory
and wild strains showed that these changes are genetical¬
ly controlled; however, a genetic analysis was not made.
Raulston et al. (1976) and Lingren et al. (1979) further
observed a mating asynchrony of about 2 hours be¬
tween a long-established laboratory colony and wild
tobacco budworms in field conditions. A similar asyn¬
chrony with a lack of mating interaction was observed
when this same laboratory strain of tobacco budworm
was used in producing backcross progeny from hybrids
resulting from a cross between the tobacco budworm and
H. subflexa (Raulston et al. 1979). But, when J. R. Raul¬
ston and A. N. Sparks (unpublished data) used trapped
wild males in producing the backcross progeny (having
the laboratory-colony background), this asynchrony and
lack of mating interaction were no longer observed.

It is the responsibility of personnel involved in the
development and production of Heliothis spp. to deter¬
mine traits that must be retained or incorporated into the
colony. Without this input, a scientifically acceptable
product cannot be developed, regardless of our physical
sophistication and production capability.

172

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175

Mass Rearing the Pink Bollworm,
Pectinophora gossypiella

By Fred D. Stewart1

Introduction

The main reason for mass rearing the pink bollworm, Pec¬
tinophora gossypiella (Saunders), at the U.S. Animal and
Plant Health Inspection Service’s (APHIS) Pink
Bollworm Rearing Facility in Phoenix, Ariz., is to supply
enough competitive, sterile moths to prevent establish¬
ment of the pink bollworm in the San Joaquin Valley of
California (the only major Southwest cotton-growing area
not generally infested). The pink bollworm became a
serious cotton pest in California’s Imperial Valley in
1966. The first moth detected in the San Joaquin Valley
was trapped in 1967. At this time, much of the ground¬
work in techniques for rearing pink bollworms had been
laid for starting up a sterile-insect-release program. This
program began on a large scale in 1970.

In the late 1950’s and early 1960’s, Knipling’s (1955)
discussion of the feasibility of using the sterile-insect
technique and the new emphasis on insect pheromones
led to research on pink bollworm nutrition and practical
rearing diets (see Vanderzant and Reiser 1956a, 1956b;
Vanderzant 1957; Vanderzant et al. 1957; Adkisson et al.
1960a, 1960b; Clark et al. 1961; Ouye 1962; and Ouye and
Vanderzant 1964), irradiation (see Bartlett 1978), and
mass rearing. Vanderzant et al. (1956), the first study to
report on mass rearing the pink bollworm, used diets of
immature peas and sprouted beans and of cottonseed and
peas. The diet of cottonseed and peas gave better results
when insect size was used as an indicator of quality. A
food preservative, methylparaben (methyl p-hydroxy-
benzoate) with methyl cellulose, was used to coat the
seed and inhibit mold growth for 14 days. These research¬
ers were the first to use antimicrobial substances in the
pink bollworm 's diet.

Richmond and Ignoffo (1964) were the first to report on
attempts to mass rear the pink bollworm on an artificial
diet developed by Ouye (1962). Their goal was to produce
moths easily and inexpensively for attractant and irradia¬
tion tests. They increased the yield per container by us¬
ing physical barriers between layers of diets. The barriers
presumably prevented not only the attraction of neonatal

’Entomologist, Pink Bollworm and Range Pests Station, Animal
and Plant Health Inspection Service, U.S. Department of
Agriculture, 4125 East Broadway, Phoenix, Ariz. 85040.

larvae to light but also movement and competitive in¬
teractions in the containers. Richmond and Ignoffo also
increased the surface areas by cubing the diet, reducing
interactions among neonatal larvae. And they presented
data suggesting that the moisture content of diet after
drying at 50° C for various periods was responsible for
variation in yields. Their data generally showed the
highest yield (41%) of mature larvae occurring at 75.7%
moisture content and yields decreasing as the water con¬
tent increased or decreased. An adult-collection system
(based on light-trap information of Richmond and
Husman 1957) was used to capture 98% of the emerged
moths. In a pilot program, 75,225 moths were produced
over 45 days when physical barriers of cotton or chipped
plastic foam were used between layers of diet in 8-oz
glass jars and plastic-coated paper cups and 6- and 9-oz
paper cups.

Many of the rearing techniques developed in this early
period (see Martin 1966 for a comprehensive review) have
been modified or expanded and are now used at the Pink
Bollworm Facility. Among these are the use of dark
rooms during the first few days of larval incubation, a
double thickness of screening to stimulate oviposition,
and, most important, adequate sanitation. Martin (1966)
foresaw that nearly all the hand procedures for rearing
could be completely automated, except the egg-laying
culture, and Mangum et al. (1969) reported techniques
able to produce 1 million sterile moths per week.

Since the 1968 commitment to a sterile-insect program by
the U.S. Department of Agriculture, the California
Department of Food and Agriculture, and California cot¬
ton growers, techniques and equipment have been con¬
tinuously changed and refined by the APHIS Methods
Development group at Phoenix. For example, use of
Honeycomb (Hexcel Corp., Long Beach, Calif.) as a pupa¬
tion substrate has allowed rapid and safe mass collection
of pupae; ceasing to layer with barriers in the diet has
saved an immense amount of labor without loss of rear¬
ing efficiency; use of caged pupae rather than adults has
reduced physical handling and injury to moths and in¬
creased oviposition yields; use of larval barriers that
restrict larvae to pupation sites has increased the collect¬
able yield; use of a new moth-collection system has in¬
creased moth yields and quality; and elimination of
pathogens from the rearing facility and the control of
fungi and bacteria that multiply in the diet have also in¬
creased yields and quality. This paper gives details of
these techniques, how we use them at the Pink Bollworm

176

Rearing Facility, and what procedures we use to assess
and insure quality.

Colony Maintenance
and Production

Oviposition and egg incubation

The egg-production units at the Pink Bollworm Rearing
Facility are housed in four 3.7-m by 13.8-m housetrailers
modified for this purpose. Two units are used for
maintenance of three different stock colonies, and two
units are used for producing eggs during the production
season (mid- April to November 1). The four units hold a
total of 2,560 oviposition cages. The genetic integrity of
each of the three colonies is maintained as much as possi¬
ble by strict separation of pupae when the oviposition
cages are set up and afterwards by separation in the
units. Genetic contamination from another strain (a
remote possibility) may result from moths escaping then-
cages. To prevent this, escaped moths are removed from
the units by black-light traps in the trailers and also by
daily vacuuming of walls, ceiling, and other surfaces.
Moths housed in the production units are a mixture from
the three colonies.

Each oviposition cage (fig. 1) is assembled from three ma¬
jor components: the plastic bottom, the fiberboard
sidewall, and the steel-mesh top. The white plastic bot¬
tom of the cage is a vacuum-formed dish 30.5 cm in
diameter with a 5-cm-high sidewall: a central cylinder 6.4
cm in diameter by 3.8 cm long projects from the bottom
and attaches to the scale-collection line. A 20-mesh
fiberglass screen is cemented to the inside of the cage to
prevent moths escaping through the bottom; beneath the
screen is a retainer ring that protects the screen when the
cage is placed on the scale-collection line. The 11.4-cm-tall
fiberboard sidewall is made from the precision cutting of
a 3-gal (11.4 liter) size fiberboard sidewall (Sealrite Co.,
Kansas City, Kan.). The fiberboard fits in a molded
groove inside the plastic sidewall. The 20-mesh stainless
steel top fits over the sidewall and is sealed with wide
tape (6.5 cm) to prevent moth escapes.

Twenty-six cages are set up each day for colony
maintenance in each of the three stock colonies, and 100
cages are set up for production. The cages are set over a
90° sanitary tee on a white polyvinyl-chloride pipe (6.4 cm
outside diameter, 3.8 cm inside diameter). The pipe not
only supports each cage in an upright position but also
forms the basis of the sealed scale-collection system. A
2.5-cm2 tubular steel rack fastened to the floor supports
80 cages on 10 pipelines. Each of these lines (two each in
five tiers) connects to a 5-cm pipeline that, in turn, con¬
nects to 15.2-cm pipeline. The largest line connects to
three cyclone dust collectors (Ridgeway and Billingsley

Figure 1.— Pink bollworm oviposition cage.

1973) in series on the outside wall of the trailer. Air is
drawn through the pipe-and-cyclone system with a
1 -horsepower Dayton (AC 108), high-pressure, direct-drive
blower (Granger, Inc., Skokie, Ill.). The cyclone system
removes about 95% of the moth scales and recycles the
clean air to the units. Efficiency requires that the
cyclones be emptied each day.

In actual use, 49 g of 6- to 7-day-old pupae (about 3,000)
are spread evenly over the bottom of the cage. On emer¬
gence, usually the 2d day after setup, the moths are sup¬
plied a 7.5% sucrose solution made with autoclaved water
containing 0.125% methylparaben. Methylparaben is in¬
cluded in the solution to prevent the growth of bacteria,
particularly Pseudomonas aeruginosa (Schroeter), which
is a common contaminant of municipal water supplies
and also a pathogen of the adults of several lepidopteran
species (F. D. Stewart, unpublished data). One-third of a
New Freedom minipad, a feminine sanitary napkin, is
soaked in the solution and placed in the hole at the center
of the pad, screen, and weight assembly. The minipads
are replaced daily with new pads and fresh solutions

177

Figure 2.— Spindles used for semiautomated
surface disinfection of pink bollworm eggs.
Spindle on right holds egg pads and screens.

because they quickly dry out under oviposition conditions
of 26.6° C and a minimum of 60% relative humidity.

After the cage has been on the support rack for 4 days, a
ring-shaped piece of 18-mesh fiberglass screen is centered
on the screen top of the cage. A similarly ring-shaped,
flexible weighted pad sheathed with vinyl is put on top of
the 18-mesh screen. The weighted pad smooths and
brings the paper and screen layers close together. The
egg pads with the screen rings are collected on a spindle
(fig. 2) each day for 8 days and incubated at 25.5° C for 3
days before disinfection and use. Collection of the screen
with the pad prevents damage to the eggs during collec¬
tion and disinfection.

Surface disinfection of eggs

The egg-disinfection room is a single room maintained
under constant temperature; air is filtered by a HE PA
(high-efficiency particulate air) filter with positive
pressure. The room is equipped with a water-mixing valve
to provide water at 28.9° C for solutions and rinse water.
A time stamp is used to record the starting and finishing
times for each egg-disinfection treatment. On the time¬
card are recorded solution temperatures and total volume
of eggs. Egg treatments are timed so that, preferably, no
more than one-half hour elapses from the completion of
disinfection to egg implantation in rearing containers.
Prolonged submersion in water decreases the hatch rate
of eggs, survival of larvae, and yield.

Figure 3.— Semiautomated egg-disinfection
system. Long tanks are used for egg disinfec¬
tion; shorter tanks are used to rinse scales
away from the egg pads.

Before the egg spindles are passed into the egg-treatment
area, the pads are counted because not more than 140
pads should be on 1 stainless steel spindle. To wet the
scales and prevent them from becoming airborne, the
pads and spindle are submerged and lightly agitated by
hand in 0.26% Tween 80 (polysorbate 80) solution in
water at 28.8° C. The eggs are removed from the egg
rings and surface-disinfected by a modification of Rich¬
mond and Martin’s (1966) technique (a 1% bleach solution
is used, and the sodium thiosulfate is omitted). In our
operation, egg removal and disinfection are facilitated by
the use of a motorized semiautomated egg-removal appa¬
ratus consisting of three transparent polyvinyl-chloride
treatment tanks (38-liter capacity) on a support rack (fig.
3). Agitation is provided to a cradle that supports the
submerged egg pads on a spindle connected to a teo-
horsepower, 30-revolutions-per-minute, eccentric-drive-
gear motor (C. H. Billingsley and J. L. Roberson, unpub¬
lished data).

In practice, the end tanks are used for the disinfection of
eggs and the middle tank for rinses. Normally, a wet egg
spindle is put into the middle tank, and a stream of
aerated water is directed near the submerged egg rings.
The scales stick to the bubbles and are carried to the sur¬
face and disposed of via an overflow drain. The egg
spindles are then immersed in one of the two outer bleach
tanks. Each tank contains 1% bleach and 0.42% Tween
80. One tank of bleach is normally enough for the treat¬
ment of three spindles. Eggs begin to loosen and settle to

178

Table 1. — Ingredients and quantities of 1 liter of artificial pink bollworm diet

Ingredient Quantity

Agar . g . . .24.2

Toasted soy flour . g . .80.0

Sugar . g 17.6

Wheat germ . g .34.8

Alphacel . g 3.0

Wesson Salts mixture . g 11.6

Methylparaben . g ... 1-9

Choline chloride (10% solution in water) . ml ... 11.6

Formaldehyde (10% solution in water) . ml , . .4.8

Potassium hydroxide (22% solution in water) . ml ... 5.8

Acetic acid (25% solution in water) . ml . . 16.4

Calco Oil Red dye . mg 150.0

Corn oil . ml 0.42

Aureomycin (chlortetracycline; 5.5% veterinary grade) . mg .53.7

FumidilB (fumagillin) . g .0.2

Locust bean gum (optional) . g 1.22-2.44

Potassium pentothenate . mg 90.97

Nicotine acid amide . mg . .46.45

Riboflavin . mg .23.23

Folic acid . mg 23.23

Thiamine hydrochloride . mg . .11.61

Pyridoxine hydrochloride . mg . .11.61

Biotin . mg . 0.929

Vitamin B12 . mg . 0.0464

Water . q.s. to 1.0 liter

the bottom after 2-3 minutes. After 5 minutes, the eggs
are siphoned off into a plastic bucket before being trans¬
ferred to a 2-liter beaker. Normally, the eggs are exposed
to the bleach solution for about 8 minutes. The viability
of the eggs begins to decrease drastically after 13 min¬
utes exposure to the bleach solution (J. L. Roberson, un¬
published data). Once all the eggs have been collected,
they are gently rinsed three times in water (28.8° C) and
then disinfected again for 10 minutes in a 9% formal¬
dehyde solution. The final step is to rinse the disinfected
eggs four times with autoclaved water. The second
disinfection is optional but is used as a prophylactic
against any cytoplasmic polyhedrosis virus and Bacillus
thuringensis (Berliner) surviving the first disinfection
treatment (Stewart et al. 1976).

A volumetric estimate of the number of eggs collected is
made for each unit. The eggs are poured into a 100-ml
graduated cylinder; the volume is noted after they settle
for 5 minutes. With 18,000 eggs/ml as a constant, the
volume of eggs is converted to the number of eggs.

Diet preparation,

storage, and shredding

From the start of the sterile-release program until 1976,
the diet used was essentially that of Ouye (1962) except

for the additions of acetic acid, Calco Oil Red N-1700 dye
(American Cyanamid Corp., Princeton, N.J.) (Graham and
Mangum 1971) dissolved in corn oil, Fumidil B (bycy-
clohexylammonium fumagillin, Abbott Laboratories, W.
Chicago, Ill.), locust bean gum, and potassium sorbate. In
1977, I modified the diet (table 1) by replacing casein as a
protein source with toasted Nutrisoy flour (soy flour, Ar¬
cher Daniels, Midland Co., Decatur, Ill.), and reducing the
amount of Alphacel (cellulose powder) and sugar (F. D.
Stewart, unpublished data). This diet is similar to a diet
developed for Heliothis virescens (Fabricius) by Shaver
and Raulston (1971). During the production season, three
or four batches (558 liters each) of diet are made each day
in a 775-liter Groen (Elk Grove Village, Ill.) steam kettle.
This amount of diet is used to fill 5,280 rearing
containers.

Cooking.— The first step in preparing the diet is to bring
the agar to a rolling boil for 5 minutes in about 225 liters
of water. Methylparaben, potassium sorbate, and Calco
red dye, partly dissolved in com oil, are added to the
steam kettle before boiling to insure the maximum dis¬
solution and dispersion in the final diet. To minimize the
preparation time, other diet ingredients are made at the
same time into a slurry in a 232-liter kettle with a
0.5-horsepower Lightnen blender, (Mixing Equipment Co.,
Rochester, N.Y.) and a 0.5-horsepower Tri-Clover blender

179

Figure 4.— 1-quart Convocans used for mass¬
rearing the pink boUworm.

(Ladish Co., Kenosha, Wis.). The Tri-Clover blender great¬
ly reduces the size of the wheat germ particles. Wesson
salt mixture (a nutritional salt mix, ICN Pharmaceuticals,
Cleveland, Ohio), sugar, Alphacel, and locust bean gum
are thoroughly mixed before the addition of soy flour and
wheat germ. All ingredients are mixed for 5 minutes.

The slurry is then pumped into the boiling agar.
Potassium hydroxide, formaldehyde, and choline chloride
solutions are added separately, in that order, and mixed
in. Once the temperature is below 65° C, vitamins,

Fumidil B, and Aureomycin (chlortetracy cline) are added.
Acetic acid is added 5 minutes after the potassium
hydroxide. The preparation of the diet is completed after
an additional 10 minutes of mixing. The diet is then
pumped into 76- by^6-cm stacking fiberglass trays at the
rate of 10-11 kg per tray.

Storage and drying.— The diet-filled trays are stored for 3
days. Storage for the first 24 hours is in a large fume
hood to facilitate the removal of acetic acid and form¬
aldehyde fumes from the facility (both are toxic to
employees and pink bollworm larvae). In a similar diet,
formaldehyde completely reacted with the diet ingred¬
ients or evaporated over 24 hours, leaving the diet
nontoxic (David et al. 1972). The diet is then dried an ad¬
ditional 2 days in a 7.8-m-long, 0.86-m-wide, and
1.8-m-high wind tunnel. A nearly laminar flow of HEPA-
filtered, fresh outdoor air passes through the tunnel and
is expelled from the building, drying the diet. During
high humidity, the drying system can also heat the air
before it passes through the tunnel. The diet is dried

because Richmond and Ignoffo (1964) reported that doing
so from 84% to 75% moisture produced the highest
yields. The APHIS diet contains 79% moisture after it
has been prepared. About l%-2% of the total weight in
moisture is lost by the time the diet solidifies; an addi¬
tional 7%-15% of the total weight is lost during the
3-day holding period. Both poor binding characteristics of
the agar and insufficient drying can lower the quality of
the diet after it is shredded. Either will produce a wet
diet that invariably gives lower yields. I think these
lower yields are related more to the amount of free sur¬
face moisture released by shredding than to the water
content of the diet. Free water can trap or drown the
neonatal larvae, or disrupt their normal behavior by caus¬
ing them to leave the diet and wander about the walls of
the rearing container instead of mining and feeding.

Shredding.— The diet is shredded with an Artofex
TR-201 (Excelsior Industrial Corp., Fairview, N.J.) com¬
mercial stainless steel food shredder with a 5-mm-mesh-
size basket. The diet is shredded into fiberglass contain¬
ers 1.4 m by 0.9 m by 0.3 m. These fiberglass containers
are raised to a workable height for the filling of the rear¬
ing containers with a mobile, custom-built, motor-driven
cable hoist.

Containerization,

development, and harvesting

Filling of rearing containers and implantation of eggs. —

The container currently used for rearing the pink
bollworm is a Sealrite (Long Beach, Calif.) 1-qt Convocan
(fig. 4). The inside of the solid container lid is laminated
with a polypropylene film that prevents excessive evap¬
oration from the cans. Both sides of the sidewall and bot¬
tom of the can are lined with polyethylene film. The cans
are made in the facility by an automated machine that
forms them from stockboard blanks at a maximum of
38/minute. They are carried to the filling-implantation
room via a 23-m-long overhead cable conveyor.

Currently, the can-filling procedure is done by hand. Per¬
sonnel doing the filling and implanting are required to
disinfect exposed skin, wear sterile surgical gowns, foot
and head covers, surgical gloves, and beard covers if ap¬
plicable. About 300 g of shredded diet is handplaced into
each container. The disinfected eggs are suspended in a
0.1% sterile agar solution (26.6°-28.9° C) used to dilute
and maintain distribution of the eggs during implanta¬
tion (Richmond and Martin 1966). The normal implant
rate may vary according to needs, but is usually
2,000-2,600 eggs suspended in a 4 ml of agar per can; the
eggs are dispensed into each container with an F-400X
filler pump (Cozzoli Machinery Co., Plainfield, N.J.) and a
20-cc stainless steel syringe (autoclaved each day). The
eggs are dispensed directly onto the diet while the can is

180

Figure 5.— Honeycomb pupation substrate.

rotated to distribute the eggs in the can. Eggs that are
accidentally splashed onto the container sidewalls usually
do not hatch, because they dry out; but, even when part
of the chorion is removed during disinfection, the egg
hatch remains high because of an advanced stage of em-
bryological development (3 days incubation at 26.6° C) at
the time of implantation into the moist environment of
the shredded diet. Disinfected eggs dry out in 1.5-2
hours when they are exposed to ambient room tempera¬
ture and humidity. Filled and implanted containers are
passed out of the implant area to be stacked in screened-
in roller racks and transported by a refrigerated panel
truck to incubation buildings.

Although nearly all the filling and implanting procedures
had been done by hand, a totally automated system was
completed in 1980. The various components— a can filler,
implant machine, and can capper— are connected with ap¬
propriate conveyor systems.

Incubation of larvae and collection of pupae. — Rearing
containers are incubated at 28.9° C for 16 days. The con¬
tainers are stacked on rearing carts and held in
darkrooms for 8 days; each cart is labeled with its unit
number, egg-collection and implantation dates, and diet
batch. Darkness is required during the early phase of in¬
cubation because neonatal larvae wander, are attracted to
normal indoor lighting (Mangum and Ridgeway 1968),
and could escape. Flashlights or red overhead lights are
used to aid personnel working in the darkrooms and to
reduce the number of larval escapes. On the eighth day of
incubation, the racks are removed from the darkroom and
moved to a pupae collection room (cutout room), where
they are held for 8 more days. Normal overhead lighting
is used when personnel are working in the area, and dim
lighting is used otherwise.

In the cutout room, two large, galvanized steel, larval-
barrier trays with rippled (Pittsburg seam) overlapping
edges (similar to the tray reported by Foster 1978) are
placed beneath each stacked tier of 120 cans. An angled,
peaked galvanized strip connects the two trays and
prevents larvae from falling between them. In each col¬
lection tray are centered a 61- by 41 -cm piece of corru¬
gated cardboard covered with a 58-cm by 38-cm by 6.4-
mm-thick sheet of Honeycomb containing about 20,000
(about 9 cells/cm2) pupation sites 3.2 mm in diameter (fig.
5).

A nondiapausing mature larva behaves the same in a
rearing container as it does in a cotton boll; it cuts an ex¬
it hole in the carpel or sidewall and drops down to
pupate. This behavior by the pink bollworm, unlike that
of many other lepidopterans now being reared, allows
easy collection of pupae and makes mass rearing the pink
bollworm possible. Most larvae enter cells, always pupate
with head end up, and usually spin silk caps over the
cells. In the Pink Bollworm Rearing Facility, the sheets
of Honeycomb with pupae are replaced in the trays on a
schedule or by necessity when they are about 60% filled
with pupae. A variable loss of pupae and larvae caused
by larval competition for pupation sites occurs if the cells
exceed 60% occupation. Once the pupae are collected in
the Honeycombs, they are not to be stacked tightly
together, because the metabolic heat accumulated will
hinder future adult mating and sperm transfer, oviposi-
tion, and egg viability (Fye and Poole 1971, Henneberry
et al. 1977) and cause premature eclosion. Partly filled
Honeycomb is replaced twice during the 8 days; each
sheet, with the cardboard backing firmly attached by
webbing, is labeled by source and stacked horizontally
with a spacer (5- by 10-cm fence wire) between each piece.

Another potentially serious problem can be triggered by
crowded conditions in Honeycomb. Spores of the
bacterial pathogen B. thuringiensis are airborne con¬
taminants present in all areas of larval rearing. At least
five varieties of B. thuringiensis have been identified
from isolates from the Pink Bollworm Rearing Facility
(C. C. Beegle, personal communication). At first, a low
proportion of larvae that have left the containers die from
wounds inflicted by other larvae. These wounded and
dead larvae readily support the saprophytic growth of
these bacteria. If this high larval density is left unattend¬
ed in the Honeycomb and becomes chronic, the concen¬
tration of airborne spores is greatly increased. In time,
nearly every dead larva will contain spores, though the
cause of death is directly related to wounds. These air¬
borne spores can enter the cans through the larval exit
holes on air currents (F. D. Stewart, unpublished data).
Larvae may also wander out of a rearing container,
become wounded or physically contact B. thuringiensis
spores, and introduce them into the same or another con-

181

Figure 6.— Moth-collection system showing
pupal holding boxes connected to moth-
collection lines.

tainer via the exit holes. The problem may be compound¬
ed if wounded larvae die in a container after contacting
spores via ingestion or wounds. Larvae still in the con¬
tainer are then directly exposed to high concentrations of
B. thuringiensis spores and crystals from cadavers, and
these conditions may start epizootics within containers.
Only in these circumstances has B. thuringiensis been
responsible for a decrease in the yield of pink bollworm at
the Pink Bollworm Rearing Facility. Similar problems
may arise when other micro-organisms, small beetles, and
mites invade or are introduced into the cans through the
exit holes.

The pupae used for oviposition are allowed to develop for
7 days (at 28.9° C) and are then collected when the
Honeycomb is gently separated from the cardboard sheet.
Pupae are allowed to fall not more than 15 cm onto a
plastic or cloth surface. Unfreed pupae are dislodged
when the Honeycomb is struck gently against a wire
frame. Pupae used for the production of sterile moths are
incubated for 3-6 days at 28.9° C before they are set up
for eclosion at the moth-collection facility.

Moth eclosion and collection. — Tests have shown that the
eclosion rate of pink bollworm moths normally exceeds
95%. Our moth-collection system uses a drawn air cur¬
rent to transport moths to collection cyclones housed in
walk-in refrigerators (4.4°-5.6°) and averages a collection

rate of 85% (range=80%-95%). A box is used to store
the pupae during eclosion. An ultraviolet light attracts
the moths from the box directly to the collection air-
stream. The boxes are large, galvanized 20-gage sheet-
metal boxes that taper to a 7.5-cm diameter pipe ending
(fig. 6). The pipe end of the box is gasketed and fits into
a 7.6-cm right 90° polyvinyl chloride tee. The large end of
the box has runners to hold six 44- by 44- by 2.5-cm
aluminum trays that allow maximum conduction of heat
away from the pupae. A door, gasketed around the edge
with neoprene (1.25-cm-wide), fits over the large end of
the box to seal it from air and light leaks. A 2.5-cm
tubular frame supports the box and permits its pipe end
to be easily slid into the 90° tee of the air line. Directly
opposite this connection in the pipe wall is a 2.5-cm Plex¬
iglas lens, and immediately behind this is a ultraviolet
lamp that radiates through a 2-mm-wide slit directly into
the box. The internal flanges of the polyvinyl-chloride
pipe have been beveled and polished to prevent injury to
the moths. Two lines, each holding eight boxes, are used
for the production from one egg-implant day. The daily
setup capacity is 3.7 million pupae at the rate of 550 g of
pupae per tray.

An air current is drawn through the screened end of the
collection pipe with a 0.5-horsepower blower motor.
Airspeed (11.9 km/h) is adjusted with a sliding sleeve
valve. An airspeed that is too slow will allow the moths
to attach to the sidewalls and lenses, obscuring the
ultraviolet light and eventually filling the collection tube
while further reducing airspeed. Too much airspeed will
cause excessive scale removal, injury, or death. In either
case, the moth quality and yield are greatly diminished.
Prevention of such losses is assured by monitoring the
line airspeeds twice daily with a Taylor anemometer
(Sybron Corp., Arden, N.C.).

A maximum of 550 g of pupae is evenly spread in each
tray and placed in the boxes. The boxes are attached to
the collection pipes, all empty tees are plugged, and the
pupae are incubated for 7 days at 27.7° C. The boxes
must be kept below 32.3°. C, a temperature that Hen-
neberry et al. (1977) reported to affect mating in pink
bollworm moths. Moths attracted to the ultraviolet light
are carried by the air current into a collection cyclone in
the coldroom. The top funnel is thickly insulated to
reduce condensation, which drowns moths. The bottom
funnel is uninsulated to facilitate rapid conduction of
heat to inactivated moths which are collected from the
cyclone one to four times daily, depending on moth densi¬
ty. A high density of moths in the collectors permits an
accumulation of metabolic heat, precludes inactivation,
and permits scale loss and moth deaths. Yields are deter¬
mined gravimetrically for each collector. Emergency
boxes are removed after 7 days, and the uncollected in¬
sects are killed by freezing.

182

Quality Control

As in most production endeavors, commercial or
biological, product quality at the Pink Bollworm Rearing
Facility depends directly or indirectly on quality of the
materials and conditions used to produce that product.
Diet quality is especially important because it greatly
influences insect quality. Quality of environmental
conditions is also carefully controlled as is quality of
nondietary supplies used.

Diet quality

To successfully mass-rear pink bollworms, all diet ingre¬
dients must be of a consistantly high quality. To get the
best quality ingredients for the least money, practical,
pharmaceutical-grade materials are used; and, if the in¬
gredient has a high use rate, a supply contract is
negotiated that legally binds the supplier to meet product
specifications. When feasible, each lot or batch is sampled
at the facility, and the samples are assayed by quality-
control personnel or by commercial laboratories before be¬
ing accepted and used. When an ingredient does not meet
product specifications, it is rejected. Rejection of an in¬
gredient has occurred, for example, when vitamins were
less than 95% potent, when bleach was less than 5.25%
sodium hypochlorite, and when Wesson Salts did not ap¬
proximate specified composition.

The quality of wheat germ is a special problem because it
is relatively unrefined. It is the most variable ingredient
in the pink bollworm diet, and annual consumption ex¬
ceeds 12.7 metric tons. Variation in protein and oil con¬
tent is caused by weather conditions and soil type where
the wheat was grown and by the milling process. Wheat
starch and bran in the wheat germ proportionately
decrease the concentration of usable nutrients. Instability
in the wheat germ caused by prolonged storage produces
even more variability in its quality.

Agar is another highly variable ingredient used in
relatively large quantity (about 10 metric tons annually).
Through experience, we have developed specific re¬
quirements for properties of agar for which we conduct
our own quality tests before awarding a contract. We re¬
quire agar of high gel strength, low viscosity at 65° C,
and a high water-holding capacity after shredding. Tests
routinely conducted include gel strength, viscosity, gela¬
tion temperature, and water retention of prepared gels.
We test for moisture content of the dry material, particle
size, and other standards found in the United States
Pharmacopeia. We also make a visual examination for
foreign matter. When agar shipments arrive, gel strength
is measured for each barrel (45.4 kg); those that do not
conform to specified gel strength are rejected.

We cannot afford to buy the diet already mixed by a
commercial source because diet quality is so important.
Although doing so would seem to be more convenient and
economical, we would lose control over a crucial step in
diet preparation. If diet problems did occur, we would
have much more difficulty tracing the cause if the diet
had been bought already mixed and weighed.

Sanitation and traffic flow

Experience has proven that the pink bollworm cannot be
successfully mass-reared without control of the insect
pathogens and of micro-organisms that make the diet un¬
palatable. A cytoplasmic polyhedrosis virus was intro¬
duced into the facility via the original laboratory culture
brought from Brownsville, Tex., in 1969 and continued to
hamper insect production until early 1976 when it was
finally eradicated. In 1972, production was hampered by
severe mold problems, particularly a strain of Aspergillus
niger (Tieg.) that may have been resistant to methyl-
paraben (a similar resistance problem was detected in
1978). In 1973, the mold problem did not recur, but pro¬
duction was limited by two pathogens, B. thuringiensis
and a transovarially transmitted Nosema sp. Midway
through the production season, the Nosema sp. was
eradicated when Fumidil B was added to the diet. Pro¬
duction reached its low point in 1974 from a widespread
epizootic of cytoplasmic polyhedrosis virus.

A milestone was reached during 1974-75 when Stewart
et al. (1976) revised techniques for egg-surface disinfec¬
tion by using 1% sodium hypochlorite solution followed
with a second disinfection in 9% formaldehyde solution.
These simple changes helped break the infection cycle; so
production increased 300% in 1975. During midseason
1975, however, a chronic cytoplasmic virus infection was
detected; investigation (F. D. Stewart, unpublished data)
revealed that moth scales from oviposition areas were
acting as vehicles for polyhedra picked up from adult
fecal matter (2% of the scales examined had at least one
polyhedral body that could be seen through a microscope).
Further observation showed that moth scales (easily dis¬
persed by air currents) were being deposited on rearing
containers in the larval incubation rooms and that
wandering neonatal larvae (capable of crawling between
the container and lid) could pick up contaminated scales
and their integument and carry them back into rearing
containers. Expansion of these observations showed
that mature larvae that commonly exit and reenter
rearing containers have a far greater vector potential
than do neonatal larvae, because the mature larvae
have greater surface area and mobility.

Before 1975, the three parent cultures of pink bollworm
were largely self-sufficient because each had its own areas

183

Egg

Treatment

CPV and Bt

on Oviposition Cages (10)

CPV and Bt on

Scales, Feces, and in Air (9)

Bt and CPV on Frames (8)

Bt and CPV on Hexcel (7)

Entry of Carrier and Infected Larvae,
Bt and CPV (6)

Inefficient Egg Treatment (I)

. Post-treatment Contamination (2)

Loose and Damaged

Rearing Lids (3)

CPV and Bt via
Moth Scales and Air (4)

CUTOUT (begins at 10 days)

Bt and CPV, Airborne and Scaleborne (5)

Figure 7.— Sources of contamination at Pink Bollworm Rearing Facility in 1974 and 1975. The in¬
crease in width of the circular arrow represents the increase in the contamination load in the facility
from different identified sources. (Note: Hexcel is a synonym for Honeycomb.)

of egg-disinfection, container inoculation, larval-
incubation, pupation, and oviposition. Sanitation
measures were limited to disinfecting work surfaces,
floors, walls, and ceilings and to some use of protective
garments. The flow of people and supplies from one area
to another was given little consideration. Given the key
to breaking the cytoplasmic virus cycle late in 1974, the
Pink Bollworm Rearing Facility underwent major modifi¬
cation early in 1975. This modification included centrali¬
zation of all egg disinfection and container inoculation,
positive pressurization of diet preparation and egg disin¬
fection, installation of HEPA-filtered air in container-
inoculation areas, elimination of people movement from
larval-development and oviposition areas to diet-
preparation and container-inoculation areas, and exten¬
sive use of scrubbing and sterile clothing.

After the discovery that moth scales could carry patho¬
gens, further modification of the facility became im¬
perative. The changes made before the 1976 season were:
restrictive routes for people and materials were
developed; to eliminate the possibility of contaminated
scales settling on disinfected eggs, egg pads were wetted
before entering the egg-disinfection room; oviposition
areas were semi-isolated; and larval incubation and pupa¬
tion areas were centralized to take maximum advantage
of prevailing winds and to allow logical and effective traf¬
fic patterns in the facility. Cleaning of equipment also

became more stringent, especially on those objects likely
to be contaminated with scales.

Once the problems were understood and remedial pro¬
cedures begun, the virus was eradicated within 2 months
and has not been detected since February 1976. Because
airborne moth scales are much easier to detect than air¬
borne polyhedra, a simple air-sampling technique (24-hour
exposure of a gridded filter paper in a 50- by 9-mm petri
dish) was used to monitor rates of scale fallout in key
areas. Counts of scales on exposed plates were used to
track progress and effectiveness of actions taken to
eradicate cytoplasmic polyhedrosis virus from the facili¬
ty. The measures taken to eliminate the cytoplasmic
virus have also proven effective against other microbial
contaminants. Figure 7 summarizes and illustrates the
magnitude of problems associated with cytoplasmic
polyhedrosis virus and B. thuringiensis at this facility.

Controlling microbial contaminants is not the only key to
successful mass rearing of the pink bollworm; levels of
airborne contaminants must be continually monitored in
all sectors to detect new organisms or increases in quanti¬
ty. Early detection is the first step in preventing
catastrophes. For example, pigmented and unpigmented
colonies of Serratia marcescens (Bizio), a known pathogen
of the pink bollworm (W. L. Belser, personal communica¬
tion), were detected on air-exposure agar plates.

184

B-po

F-2

V-a

N-a

2.0-1

CO

c

o

r 1.5

o

Q

v_

a>

a

CO

1.0-

o

v_

a>

-O

io.5

c

o

a>

0 J

B Mean Daily Sterile Moth Production
I | Mean Production Goal

B-p

B-l

B-l

B-po

F-3

F-po

F-a

F-a

V-l

V-l

V-3

V-a

N-pp

N-3

N-a

B-po N-a

B-pp

F-P

V-l

N-a

n

B-po

F-a

V-o

N-a

1970 1971 1972 1973 1974 1975 1976 1977 1978 1979

Figure 8.— Summary of the daily production goals for each year from 1970 through 1979 and the in¬
fluence of associated micro-organisms on the mean daily sterile moth production. Micro-organisms:
F=Aspergillus niger; B=Bacillus thuringiensis; V =cytoplasmic polyhedrosis virus; N=Nosema sp.
Letters followed by characters indicate presence and effect on production; a=absent; p=present, ef¬
fect unknown; pp=possibly present, no examination made; po=present, no effect; l=slight effect;
2=moderate effect; 3=severe effect.

Bacteriological smears were taken from different sites in
the contaminated pupation area, the sources were iden¬
tified, and intensive sanitation measures eradicated the
bacterium. Increases in plate counts may be by chance,
but they may also indicate serious chronic problems or il¬
licit changes in sanitation procedures or traffic flow.

In general, the measures now used to sanitize the facility
include mopping floors and washing walls and ceilings
with bleach, quaternary ammonium compounds, phenolic
compounds, and stabilized chlorine dioxide solutions.

Chlorine dioxide is advantageous because it is relatively
stable and noncorrosive, and it can be rapidly and effec¬
tively applied with airless spray guns to almost any sur¬
face— particularly walls, ceilings, and supplies entering
the facility. All glassware, rinse water, and clothing used
in egg disinfection are autoclaved daily. Every failure to
meet production quotas at the Pink Bollworm Rearing
Facility has been directly or indirectly related to the
dominating, harmful effects of micro-organisms (fig. 8), so
sanitation is extremely important in insuring that pro¬
duction goals are met.

185

Insect quality

Thus far, standards used to define overall quality of the
insect have been limited by a lack of measurable traits
showing how sterile moths will perform in the field.
Regardless of what those traits might be, the measure¬
ment system must lend itself to routine daily use with
several individuals or samples. For example, measure¬
ment of flight capability with flight mills (Flint et al.

1975) is not a practical system because of size and fragili¬
ty of the moths and the nonreproducibility of results (R.
T. Staten, unpublished data). Quality traits of sterile pink
bollworm moths that have been studied in the field in¬
clude competitiveness (Van Steenwyck et al. 1979), at¬
tractiveness of irradiated females (Flint et al. 1973), and
time and duration of pheromone release by irradiated
females in the field (P. Lingren, unpublished data).
Routine monitoring of these traits would greatly en¬
hance assessment of overall moth quality, but field
measurements are expensive, time consuming, and not
readily adaptable to routine quality testing. Laboratory
tests have only been used to monitor mating and longevi¬
ty characteristics.

At the APHIS field laboratory in Bakersfield, Calif., ir¬
radiated moths are sampled daily just before release. The
samples are subjected to various measurements including
mating, mating potential, and longevity. Data on mating
and mating potential— tests determined by spermato-
phore counts (Ouye et al. 1965)— are used as tests of
handling and shipping procedures and of potential
usefulness of moths in the field. Measurement of longevi¬
ty over 14 days is used to estimate residual or working
populations of moths in the field. More importantly,
these data are used as signals of production problems
(high rates of mortality early in the 14 days have always
been disease related) that require investigation. In all
cases, interpretation is largely a matter of comparing
present performance with past performance.

Other things we might consider as part of the quality-
control system include daily monitoring of comprehensive
production data (eggs, pupae, and moths produced) and
daily measurement of moth size and pupal size. Such
data are also used as indicators of potential problems and
stimulate thorough investigation.

References

Adkisson, P. L.; Bull, D. L.; and Allison, W. E.

1960a. A comparison of certain artificial diets for lab¬
oratory culture of the pink bollworm. J. Econ.
Entomol. 53: 791-793.

Adkisson, P. L.; Vanderzant, E. S.; Bull, D. L.; and
Allison, W. E.

1960b. A wheat germ medium for rearing the pink

bollworm. J. Econ. Entomol. 53: 759-761.

Bartlett, A. C.

1978. Radiation-induced sterility in the pink boll¬
worm. U.S. Sci. Educ. Adm. Agric. Rev. Man.
West. Ser. 1, 25 pp.

Clark, E. W.; Richmond, C. A.; and McGough, J. M.

1961. Artificial media and rearing techniques of the
pink bollworm. J. Econ. Entomol. 54: 4-9.

David, W. L.; Ellaby, S.; and Taylor, G.

1972. The fumigant action of formaldehyde incor¬
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granulosis virus of Pieris brassicae and its
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Flint, H. M.; Staten, R. T.; Bariola, L. A.; and Palmer,

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1973. Gamma-irradiated pink bollworms. Attrac¬
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Flint, H. M.; Wright, B. S.; Sallam, H. A.; and Horn, B.

1975. Dispersal and mating in the field by pink
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Foster, R. N.; Staten, R. T.; and Moss, L. K.

1978. A procedural manual for rearing the parasites
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Fye, R. E„ and Poole, H. K.

1971. Effect of high temperatures on fecundity and
fertility of six lepidopterous pests of cotton in
Arizona. U.S. Dep. Agric. Prod. Res. Rep.

131, 8 pp.

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1971. Larval diets containing dyes for tagging pink
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64: 376-379.

Henneberry, T. J.; Flint, H. M.; and Bariola, L. A.

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1955. Possibilities of insect control for eradication
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Mangum, C. L., and Ridgeway, W. O.

1968. Phototactic response of first instar larvae of
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1966. Pink bollworm. In Smith, C. N. (ed.), Insect
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1965. Mating studies on the pink bollworm, Pec-
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1964. Mass rearing pink bollworms. J. Econ. En¬
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1966. Technique for mass rearing of the pink
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1957. Growth and reproduction of the pink

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1957. The role of dietary fatty acids in the develop¬
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Vanderzant, E. S., and Reiser, R.

1956a. Aseptic rearing of the pink bollworm on syn¬
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1956b. Studies of the nutrition of the pink bollworm
using purified casein media. J. Econ. En¬
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1956. Methods for mass rearing of the pink

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187

Production of Boll Weevils,
Anthonomus grandis grandis

By J. L. Roberson1 and J. E. Wright2

Introduction

History and distribution

of the boll weevil

The boll weevil, Anthonomus grandis grandis Boheman,
is a New World pest suspected to have originated in
southern Mexico or northern Central America. The first
specimen was collected near Veracruz, Mexico, in the
1830’s. Its taxonomy was described in 1843 by Boheman
from specimens collected in Veracruz. The first report of
the boll weevil in the United States was during the fall of
1894 near Brownsville, Tex. By 1898, it had spread
across Texas; it was in Louisiana by 1903 and in Okla¬
homa, Arkansas, and Mississippi by 1907. It completed
its distribution across the southeastern Cotton Belt of
the United States by 1922. (For a detailed history and
taxonomy of the boll weevil, see Burke 1968.) The boll
weevil develops on cultivated and wild cotton of the
genus Gossypium and other closely related genera. (For
more information on host plants of the boll weevil, see
Cross et al. 1975.)

Life cycle of the boll weevil

in the field

The boll weevil is a holometabolous insect with an egg,
larval, pupal, and adult stage. The life cycle may be com¬
pleted in 18-21 days at temperatures of about 30° C and
60% relative humidity, so five or more generations may
occur each year. The adult is 6-7 mm long, weighs about
15 mg, and ranges in color from tan to dark gray or
brown. The proboscis is about half the length of the
body, and the adult has a spur on the inner surface of
each front femur.

With its proboscis, the adult punctures squares (flower-
buds) or bolls for feeding and oviposition. The females lay
eggs in feeding punctures and then seal the holes with a
gluey substance. These eggs hatch in 3-5 days; hatching
is followed by a 7-12 day feeding period that causes the
squares or bolls to abscise. Larval feeding is completed in
the fruit on the ground. The pupal stage lasts 3-5 days

'Insect product manager, Boll Weevil Research Laboratory,
Agricultural Research Service, U.S. Department of Agriculture,
Mississippi State, Miss. 39762.

'Research leader, Boll Weevil Research Laboratory.

and is followed by adult emergence. Female weevils begin
to lay eggs 3-7 days after feeding and mating. This life
cycle occurs repeatedly until cotton is no longer available.
Then, adult weevils overwinter in diapause in surface
woods trash, along ditch banks, in cottonfields, and in
trash and litter around gins and farm buildings. Diapause
generally ends in the spring about the time squares are
being formed on the cotton plant.

Laboratory Rearing
of the Boll Weevil

Rearing procedures for the boll weevil at the U.S. Animal
and Plant Health Inspection Service’s Robert T. Gast
Boll Weevil Rearing Facility are a compromise between
the insect’s basic necessities and the artificial conditions
required to produce the numbers and quality of insects
needed for research and proposed field testing of sterilized
weevils. Insect-control plans using sterile-insect techniques
demand massive numbers of high-quality insects to be pro¬
duced and delivered on specific use dates. Since mass¬
rearing procedures were first reported (Gast and Davich
1966), research has been directed to development and in¬
corporation of mechanized operations where possible. The
aim is to reduce risks of microbial contaminants in sen¬
sitive work areas and increase production capacities. Most
recent advancements include incorporation of quality-
control techniques to identify problems and evaluate pro¬
duction standards.

Colony selection

The colony presently being mass-reared at the R. T. Gast
Rearing Facility originated from an ebony strain selected
by Bartlett (1967); this character is now used as a marker
to distinguish the laboratory weevils from native weevils.
The ebony characteristic is semidominant and requires
selection and backcrossing from the colony for its
maintenance. In 1975, native specimens were collected in
North Carolina and crossed with laboratory ebony; the pro¬
geny were selected for ebony color. In October 1978, ebony
adults were selected from pupal cells; these adults were
colonized, and the following generation was inspected and
nonebony weevils discarded. The selected colony was
isolated and increased to replace the existing colony.

Adults reared for release in the 1979 North Carolina boll
weevil eradication trial were inspected daily; they exhibited
98% ebony characteristic. Calco red dye was added to the
larval diet (Lindig et al. 1980) to provide an additional
marker.

188

Table 1.— Contents of a prennix batch of dry ingredients for adult
and larval diets of the boll weevil

Ingredient

Amount (g)

Adult diet Larval diet

Cottonseed meal, sifted .

.... 28,500

31,800

Agar

8,400

8,280

Sugar

6,850

7,800

Promine D .

10,800

12,280

Corncob grits, 60 mesh

. 6,000

Cholesterol .

300

360

Wesson salt mix .

1,125

1,260

Vitamin mix1 .

1,170

1,260

Potassium sorbate .

585

400

Methylparaben (methyl p-

375

400

hydroxybenzoate) .

Ascorbic acid

. . . . 270

290

Methenamine

60

Cottonseed meats .

. . . 10,800

10,724

‘See table 2.

Laboratory rearing procedures

Preparation of diet premix. — Diet ingredients are received
in bulk. As each shipment of ingredients is received, it is
identified either by the date received or by the industrial-
batch-lot number. As each premix batch of dry ingredients
(tables 1 and 2) is prepared, it is identified numerically;
the industrial- or storage-lot number of each ingredient
used is recorded for reference. This record makes it possi¬
ble to find out when new ingredients were introduced into
the production line and provides valuable information if
diet quality is questioned. Two records are made for each
premix batch; one remains on file in the weighing area,
and the other is attached to the diet container. The work
operations for diet preparation are: All ingredients are
weighed with top loading balances. Cottonseed meal is
sifted through a 32-mesh sieve screen with a
2.5-horsepower automatic feeder/sifter/shaker (Griffin and
Lindig 1974b). All ingredients except agar, Promine D
(soy protein), and corncob grits are weighed and loaded
into a heavy-duty mixer. These ingredients are mixed
about 20 minutes to obtain a uniform distribution of cot¬
tonseed meats. The bulk mix is transferred by conveyor
to a hammer mill (Griffin 1979) where it is milled and
returned to the blender. The remaining diet ingredients
are weighed and added to the blender for a 30-minute
final mixing. When the blending operation is completed,
the mixture is resifted, packaged, labeled numerically,
and transferred to the diet-preparation area.

Operation of the equipment to prepare the blended diet
mixture causes dust problems in the rearing area. The

Table 2. — Contents of vitamin mix in a premix batch of dry ingre¬
dients for boll weevil diet1

Ingredient Amount (g)

Niacinamide . 20

Calcium pantothenate . 20

Riboflavin . 10

Thiamine hydrochloride . 5

Pyridoxine hydrochloride . 5

Folic acid . 5

Inositol . 40

Sugar . 6.160

‘Ingredients weighed on balances and then ball-milled for 1 hour
to give a homogenous blend.

dust contains diet ingredients that harbor bacterial and
fungal growth. So the work area is well-ventilated to min¬
imize dust inhalation and development of potentially ex¬
plosive conditions. Also, the work area is maintained in a
separate building to minimize threat of microbial contam¬
ination to the rearing colony.

Diet preparation. — Both adult and larval diet formula¬
tions are mixed with water and flash-sterilized in one of
the three No Bac Unitherm IV (Cherry-Burrell Corp.,
Cedar Rapids, Iowa) processing systems (Griffin and Lin¬
dig 1974b). The flash-sterilizing units are interchangeable
and serve to back each other up in case of mechanical
malfunction. Diet is prepared at the rate of 151 ±19 liters
per hour. It is sterile as it enters the stainless steel trans¬
fer lines (0.7 i.d. by 1.0 cm o.d.) going to the desired area.
Three forms of diet are prepared in the room: pellets for
adult oviposition, slabs containing diflubenzuron3
(Wright, McCoy, et al. 1980) for feed-out of weevils to be
irradiated, and larval diet for rearing trays. Before larval
diet preparation, the tubing used to transfer sterile diet is
flushed free of a disinfectant solution (1.0% Mikro-Quat,4 *
Economics Laboratory, Atlanta, Ga.) and treated with
Oxine6 (Bio-Cide Chemical Co., Norman, Okla.) at 500 p/m
(parts per million). The Oxine is introduced and main¬
tained in the lines for 15-20 minutes. After use, the equip¬
ment and transfer tubing are treated with a strong, oxidizing
soap to rid the system of adhering organic matter, then
neutralized with an acid detergent. After standard clean¬
up procedures, a germicidal agent (1.0% Micro-Quat) is
pumped into the lines and held until the next use.

W-[[(4-Chlorophenyl)amino]carbonyl]-2,6-difluorobenz amide.

4Active ingredients: Alkyl (50% C14, 40% CI2, 10% C,6) dimethyl

benzyl ammonium chlorides, 9%; trisodium

‘Active ingredient: chlorine dioxide, 2%.

189

Pellet preparation. — A batch of diet is mixed, sterilized,
and pumped to the pellet-forming unit. The pellet-forming
unit is constructed with 12 stainless steel water-jacket
lines. The diet is pumped into each of the lines at
12-second intervals. Cold water flows continually around
each line, causing the diet to gel; introduction of liquid
diet into the line with gelled diet forces the gelled diet
out, in rod form, and into a rotating wire wheel that cuts
the diet into pellets (0.8 cm in diameter by 1±0.5 cm
long). The pellets fall into a vat containing a mixture of
40% beeswax and 60% paraffin maintained at 66° C. The
pellets submerge and are completely coated with the wax
mixture. As they settle in the wax mixture, the pellets
are collected on an inclined endless wire belt that
transfers them from the vat to a catch container. They
are then poured from the catch container into pans (24.1
cm wide by 13.8 cm deep by 40.9 cm long) for holding
and storage in a cool room; or they are transferred to the
oviposition room (Griffin and Lindig 1974a).

Slab preparation. — The adult-diet formulation containing
100 p/m diflubenzuron (25% wettable powder) is processed
by the flash-sterilizing system. Processed diet is cooled to
about 71.1° C and transferred through stainless steel tub¬
ing to a Mateer-Burt diet-filling unit (Mateer-Burt Co.,
Wayne, Pa.). The unit is calibrated to dispense 285 ml of
liquid diet into serving trays (34.3 cm wide by 1.9 cm
deep by 45.7 cm long). The trays are positioned in a chain
drive that moves them through a chill tunnel (53.3 cm
wide by 172.7 cm long) to gel the exposed diet surface.
The machine fills and delivers six gelled trays per minute
for positioning in mobile storage racks (94-tray capacity).
The racks with trays are transferred to the adult feed-out
area for use. The trays are washed and autoclaved before
reuse.

Preparation of larval trays. — Sanitation is vital to suc¬
cessful preparation of the larval trays. So it is done in an
isolated room with maximum security for sanitation; the
room is restricted to assigned personnel who must wear
clean cover clothing when entering and working in it. All
air-conditioning outlets in the room have absolute HE PA
(high-efficiency particulate air) filters and operate 24
hours a day. Microbial levels are routinely monitored.
About 15-20 exposure plates prepared with trypticase
soy agar (TSA) growth medium are put in specified places
in the room for 20 minutes and exposed to atmospheric
microbial settlement while trays are being prepared.

Touch plates prepared with the TSA growth medium are
used to press against surface areas of materials and
equipment to obtain samples showing levels of microbial
contamination. Samples of tray components such as diet,
eggs, and sand-corncob mixture are taken with each new
batch lot that is introduced into the production line
(Sikorowski 1975).

Trays for larval development are made from a stock roll
of polystyrene that is 15 cm wide by 18 mil thick. The
polystyrene sheeting is heated to make it pliable. A
vacuum is introduced, and the polystyrene is pulled
within a mold that forms cavities in the sheeting. The
sheeting with cavities serves as a larval rearing tray. It is
secured and moved by clamps on the Anderson Form Fill-
Seal Machine (Model 655-B, Anderson Bros. Manufac¬
turing Co., Rockford, Ill.) in a synchronized fashion that
mechanically completes all tray-preparation tasks. (See
Harrell et al. 1977 for the original concept and design of
tray-forming equipment in boll weevil rearing. Accessory
equipment to dispense and chill the diet and dispense the
eggs and sand-corncob mixture was improved by Griffin
et al. 1979.)

The larval diet formulation differs from the adult for¬
mulation in ratio of ingredients; however, processing pro¬
cedures with the flash sterilizer are the same. The sterile
diet is delivered to a dispenser that is synchronized with
the Anderson Form-Fill-Seal machine; and, as the
sheeting passes below the dispenser, each cavity is filled
with 185 ml of liquid diet adjusted to 40°±10° C. This
temperature permits the diet to flow evenly in the cavity
but gel quickly with a minimum of cooling. The cavities
filled with diet enter a 1.5-m-long cooling tunnel main¬
tained at 1° C, a procedure that gels the diet surface. The
clamp track moves the cavities from the tunnel and posi¬
tions them beneath a pump and sprayer that deliver 4 ml
of eggs (2,100 eggs) in a furcellaran solution (0.5%) to the
diet surface. The furcellaran solution is used to suspend
the eggs, thus enabling uniform distribution with spray¬
ing. The clamp track advances the cavities with diet and
eggs below a hopper that dispenses a sterile sand-corncob
(70 : 30) mixture containing antibiotics (Sikorowski et al.
1980) over the diet surface to absorb moisture and force
hatching larvae to feed on the diet. The clamp track then
moves the cavities into position so that a Tyvek (Stand¬
ard Packaging Corp., Atlanta, Ga.) cover can be heat-
sealed over the surface. Tyvek is a porous polyethylene
material that allows air and moisture exchange in the
cavities. The clamp track moves the sealed cavities to a
shearing station that cuts the sheeting into trays, each
with two cavities. The trays are agitated by hand with a
circular motion to spread the sand-corncob mixture over
the diet surface then manually positioned on metal tracks
in rackveyors (Griffin 1979) for transfer to the larval
holding area. The rackveyor is designed for maximum
space efficiency, mobility, and tray suspension for good
air movement around trays.

A record sheet to aid in monitoring of production proces¬
ses is attached to the rackveyor and remains as a reference
for quality control. Data corded on the sheet iden¬
tify the trays when different lot numbers of tray compo¬
nents or abnormal procedures are introduced or occur in

190

the production line. Data are normally recorded on rearing
trays by tagging them when changes occur in the diet-
preparation batch number, the polystyrene roll, the Tyvek
roll, the bottles containing egg suspension, etc. All trays
on the rackveyor are inspected for loose Tyvek covers; if
any are found, they are sealed with tape before transfer of
the rackveyor into the larval holding room.

Larval development. — The larval-development room is
maintained at about 31° C, 55%±5% relative humidity,
and 24 hours of light. Egg hatch occurs within 3 days
after eggs are put in trays; larvae feed vigorously. Pre¬
pupae forms appear in isolated cells by the 9th day, and
adults can be observed in the trays by the 13th day. The
larval holding room is kept clean with an absolute air¬
filtering system, and all personnel are required to wear
clean clothing when in the area. Temperature and humidity
are monitored to insure proper drying of the diet for nor¬
mal insect development. If abnormal conditions arise and
adults emerge earlier than usual, the rackveyors are
removed from the area to prevent contamination in the
cleanroom. On the 13th day after eggs are put on the tray,
the rackveyors are transferred to the adult emergence area.

Adult emergence and collection.— The rackveyor contain¬
ing trays of emerging adults is stationed in the workroom.
A small hole is melted in the exposed end of each tray
with a heating element, and the rackveyor is then transfer¬
red to a darkened emergence chamber (2.4 by 2.0 by 2.2
m). Twenty-eight 3.78-liter plastic jars are positioned over
openings (5 cm in diameter) in the outside of the chamber
wall. Light entering the chamber through the jar portals
attracts emerging adults, which become trapped in the
jars. Adults are harvested from the collection jars daily,
more often if necessary to prevent damage to collected
adults. Adults emerging on days 1 and 2 are transferred to
laying cages and used as maintenance stock. Adults
emerging on days 3, 4, and 5 are bagged in feeding
packets and fed for 5 days before irradiation. The
rackveyor with spent trays is transferred to a heating
chamber and treated for 24 hours at 52° C to kill weevils
remaining in the trays. The trays are bulk-packaged in
two-ply paper waste-disposal bags, sewn closed, and
discarded.

Adult feed-out for colony maintenance.— The adult weevils
for egg production are maintained for 14 days at 28° C,
50% ±5% relative humidity, and a photoperiod with 20
hours of darkness, 4 hours of light. These adults are held
in stainless steel cages (45.7 by 96.5 by 5 cm), described by
Griffin et al. (1979). These cages are constructed with
removable tops and 12- by 12-mesh stainless steel cloth
bottoms. About 9,000 weevils are placed in each cage
(estimated by weight). The cage contains two baskets (41.2
by 41.2 by 1.2 cm) to hold diet pellets that serve both as
food and as oviposition sites. After the first 2 days, the

eggs are mechanically extracted from the pellets (Griffin
and Lindig 1977), cleaned, and surface-sterilized
(Sikorowski 1977). They are then mixed with a sterile
furcellaran solution (0.5%) for suspending and dispensing
in rearing trays.

Adult feed-out for irradiation. — Adult weevils collected for
irradiation are placed in 20.5- by 40.6-cm (mosquito net¬
ting) bags (6,000/bag; O. T. Malone, unpublished data) and
maintained for 5 days on diet containing antibiotics and
100 p/m diflubenzuron (Wright, Roberson, and Dawson
1980). They are transferred to fresh diet daily. En¬
vironmental conditions of the feeding rooms are main¬
tained at 28.3°±1° C, 50%±5% relative humidity, and 20
hours of darkness. On the morning of the sixth day, the
bags are transferred to a walk-in refrigerator, and the
weevils are chilled until immobilized (30 minutes at
2.7°±1° C); the dead or very small adults are separated by
processing in a seed separator. About 130,000 adults are
placed in a Plexiglas canister, (28.2 cm in diameter by 10.2
cm deep; J. G. Griffin, unpublished data) and transferred
to the irradiation area for processing. The weevils in
canisters are irradiated with 10 krads from a cesium
source and packaged in cool boxes for shipment to release
sites.

Maintaining production yields

To insure that the required quantity of boll weevils is
produced, production records are kept in each rearing sec¬
tion. These provide a quick review of each production
area so that potential rearing problems can be identified.
Effective control of production is achieved by recording
and monitoring production procedures as follows:

1. When diet ingredients are weighed, the industrial batch
number is recorded and maintained with each diet
batch.

2. Ingredient and operational variables are recorded dur¬
ing diet preparation.

3. Records of tray preparation identify trays that may
be affected by changes or adjustments that have
occurred.

4. The eggs-per-female production rate is recorded daily.

5. Adult emergence for each egg-implantation date is
monitored and is compared with estimates of adult
production based on numbers of eggs implanted, hatch
rates, and larvae per tray.

6. Egg hatch for each implantation date is monitored as
an indicator of viability.

7. Microbial contamination in cleanrooms is continually
monitored to maintain sanitation standards.

The collection and use of these data provide information
essential for management to meet production goals.

191

Quality Control

Quality-control data are needed to certify that all rearing
and sterilization processes are functioning properly so
that the insects produced meet established standards.
Data obtained from sampling boll weevils irradiated for
release are as follows:

1. Bacterial gut-load level (Sikorowski 1977). Twenty
adult boll weevil specimens are collected from each
egg-implantation date. Weevils are analyzed and
classified into groups I-IV according to their bacterial
gut-load level.

2. Sperm transfer. Twenty-five paired matings of ir¬
radiated and nonirradiated adults are made from each
day’s production.

3. Pheromone production. One hundred males are an¬
alyzed for pheromone production (McKibben et al.
1976).

4. Egg viability of irradiated adults. A gross sample,
about 9,000 irradiated mixed-sex adults, are collected
from irradiation canisters daily and maintained in lay¬
ing cages for 5 days. Eggs are collected daily and in¬
cubated to determine viability.

References
Bartlett, A. C.

1967. Genetic marker in boll weevil. J. Hered.

58: 159-163.

Burke, H. R.

1968. Geographic variation and taxonomy of An-
thonomus grandis Boheman. 152 pp. Texas
Agricultural Experiment Station, College Sta¬
tion, Tex.

Cross, W. H.; Lukefahr, M. J.; Fryxell, P. A.; and Burke,
H. R.

1975. Host plants of the boll weevil. Environ. En-
tomol. 4: 19-25.

Gast, R. T., and Davich, T. B.

1966. Boll weevils. In C. N. Smith (ed.), Insect Col¬
onization and Mass Production, pp. 405-418.
Academic Press, New York.

Griffin, J. G.

1979. “Rackveyor” for use in mass rearing of boll
weevils. U.S. Sci. Educ. Adm. Adv. Agric.
Technol. South. Ser. 4, 3 pp.

Griffin, J. G., and Lindig, O. H.

1974a. Mechanized production of boll weevil diet
pellets. Trans. ASAE 17: 15-19.

1974b. Flash sterilizers: sterilizing artificial diets for
insects. J. Econ. Entomol. 67: 689.

1977. System for mechanically harvesting boll

weevil eggs on artificial diets. Trans. ASAE
20: 254-256.

Griffin, J. G.; Lindig, O. H.; Roberson, J.; and Sikorow¬
ski, P. P.

1979. System for mass rearing boll weevils in a
laboratory. Miss. Agric. For. Exp. Stn. Tech.
Bull. 95, 15 pp.

Harrell, E. A.; Perkins, W. D.; Sparks, A. N.; and Moore,
R. F.

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Lindig, O. H.; Wiygul, G.; Wright, J. E.; Roberson, J.;
and Dawson, J. R.

1980. A rapid method for mass rearing boll weevils.
J. Econ. Entomol. 73: 385-387.

McKibben, G. H.; McGovern, W. L.; Cross, W. H.; and
Lindig, O. H.

1976. Search for a super laboratory strain of boll
weevils: a rapid method for pheromone
analysis of frass. Environ. Entomol. 5: 81-82.

Sikorowski, P. P.

1975. Microbial monitoring in the boll weevil rear¬
ing facility. Miss. Agric. For. Exp. Stn. Tech.
Bull. 71, 20 pp.

1977. Method for surface sterilization of boll weevil
eggs, and the determination of microbial con¬
tamination of adults. Southwest. Entomol.

2: 32-36.

Sikorowski, P. P.; Kent, A. D.; Lindig, O. H.; Wiygul,

G.; and Roberson, J.

1980. Laboratory and insectary studies on the use
of antibiotics and antimicrobial agents in
mass rearing of boll weevils. J. Econ. En¬
tomol. 73: 106-109.

Wright, J. E.; McCoy, J. R.; Sikorowski, P. P.; Roberson,
J.; and Dawson, J. R.

1980. Boll weevil: effects of different combinations
of diflubenzuron, antibiotics, fumigation, and
irradiation. Southwest. Entomol. 5: 84-85.

Wright, J. E.; Roberson, J. L.; and Dawson, J. R.

1980. Boll weevil: effects of diflubenzuron on sperm
transfer, mortality and sterility. J. Econ. En¬
tomol. 73: 803-805.

192

Mass Production of Screwworm
Flies, Cochliomyia hominivorax

By Harold E. Brown1

Introduction

The mass production and release of irradiated sterile
screwworm flies, Cochliomyia hominivorax (Coquerel), is
the first and most outstanding program to control an in¬
sect pest by release of the sterilized insects. The develop¬
ment of rearing techniques on artificial media (Melvin
and Bushland 1936, 1940), the initial concept of the
sterile-male technique by Knipling in 1938 (1955, 1959),
and the determination that viable sterile males could be
produced after irradiation of the pupae by X-rays (Bush-
land and Hopkins 1951) and gamma rays (Bushland and
Hopkins 1953) provided the technology that led to the
completion of successful screwworm eradication pro¬
grams. Sterile-male release has eradicated screwworms
from the southeastern (Knipling 1960) and southwestern
(Bushland 1975) United States, Puerto Rico (Williams et
al. 1977), and twice from the island of Curacao, Nether¬
lands Antilles, (Baumhover et al. 1955, Coppedge et al.
1978). Billions of screwworm flies have been artificially
reared, irradiated, and released during these control pro¬
grams.

Presently, the U.S. Department of Agriculture’s South¬
west Screwworm Eradication Program at Mission, Tex.,2
and the Mexican-American Program for the Eradication
of the Screwworm at the rearing plant located in Tuxtla
Gutierrez, Chiapas, Mexico, rear a combined total of
about 500 million screwworm flies each week. This report
summarizes techniques for strain development, selection,
production, and quality control used by these programs
for the production of high-quality screwworm flies.

‘Research chemist. Agricultural Products Quality Research Unit,
Agricultural Research Service, U.S. Department of Agriculture,
P.O. Box 267, Weslaco, Tex. 78596.

2Since I wrote this paper, the program at Mission, Tex., has been
closed down. The eradication program at Mission was under the
auspices of the U.S. Department of Agriculture’s Animal and
Plant Health Inspection Service, which shared a location with a
screwworm research program of the U.S. Department of Agri¬
culture’s Agricultural Research Service.

Colony Selection

Selection of flies used in the colony for mass rearing is
divided into two operations: the development and selec¬
tion of candidate strains and the introduction of the
strain of flies to be used in the rearing colony.

Strain development and selection

Strains of screwworm flies to be used in the mass-rearing
facility are usually generated from egg masses collected
from areas that will be treated with the sterile flies
(Crystal and Ramirez 1975, Crystal and Whitten 1976).
Egg masses are collected from naturally or intentionally
wounded animals and transported to the laboratory for
incubation, larval rearing, and strain expansion. When
egg masses are collected in remote areas, the larvae are
often reared in temporary laboratories and the pupae
transported to the permanent laboratory for further de¬
velopment. More recently, strains have been developed
for release back into the specific geographic locations
they are derived from. It is hoped that matching loca¬
tions in this way will take advantage of selective mating
if, in fact, native females are more likely to mate with
males derived from the same geographic area.

In the southeastern eradication program (Baumhover et
al. 1966), efforts were made to collect egg masses from
scattered areas to get a wide genetic base. These strains
were selected for sexual vigor, longevity, and resistance
to starvation to get aggressive, artificially reared flies.
The early strain (Baumhover et al. 1966), labeled “the
Florida strain,” was successful in eradicating
screwworms from the southeastern part of the United
States. The Florida strain was also used for mass pro¬
duction when the Southwest Screwworm Eradication Pro¬
gram was begun in 1962 since time and resources for
field-testing candidate strains were inadequate. (See
Bushland 1975 for a discussion of strain development and
changes in strains for mass production through 1974.)

How many and what kinds of laboratory and field evalua¬
tions can be used in selecting strains for mass rearing are
limited. But all candidate strains are tested in the labo¬
ratory for larval weights, pupation, emergence, mating,
fertility, oviposition percentages, longevity, and response
to heat and starvation stresses (Crystal and Ramirez
1975, and Crystal and Whitten 1976). Biochemical,
genetic, and physical measurements have also been used

193

sometimes to compare strains. Isoenzyme (Whitten 1980)
and electroretinographic (Goodenough et al. 1977, 1978)
techniques have been used and added to the series of
evaluations to determine whether a strain is suitable for
mass production and release by the eradication program.
Also, field evaluations similar to that reported by Ahrens
et al. (1976) have helped assess candidate strains for
mass production. These evaluations use trap-back studies
of released flies from the various strains and provide in¬
formation about the migration, dispersal, and survival of
the flies.

Colony introduction

At Mission, once a strain has been selected, it is expanded
in the U.S. Agricultural Research Service’s screwworm
research facilities; then pupae are transferred to the
methods development section of the U.S. Animal and
Plant Health Inspection Service’s Mission facility at a
rate of about 4 liters/day for further expansion. Currently,
strain expansion takes place in the methods development
section where enough flies can be reared for production
and for field tests before strain introduction. The expan¬
sion is timed to provide pupae for 21 days, a period that
covers a one-generation cycle of the fly. Production is
reduced during strain changes since, to prevent con¬
tamination, all rearing areas (fly colony, larval starting
rooms, rearing floors, and pupation and pupal holding
rooms) are cleared of all the previous strain before in¬
troduction of the new strain. During these strain
changes, problems often occur in establishing proper
oviposition, egg-incubation, and starting and rearing
schedules. These production bottlenecks must be over¬
come, and the new strain must adapt to the laboratory
environment.

Colony Maintenance
and Oviposition

About 5% of the flies produced are used for the brood
colony and for egg production. The brood colony is
selected from the normal system for pupae handling at
about the midpoint of each work shift. Pupae are col¬
lected just after pupation and are placed in screen-
bottomed trays (46 by 66 by 5 cm), 2.5 liters/tray. Trays
of pupae are held in a constant temperature (24°-25° C)
and relative humidity (50%-60%) in a separate room for 6
days and then placed in the colony cages. These metal
cages (109 cm wide by 152 cm long by 180 cm high) are
stocked with 2 trays of pupae (about 50,000) each.
Emergence occurs in 24-48 hours. Cages are equipped
with five rows of paper curtains hanging from rods run¬
ning lengthwise in the cages. These provide resting areas
for the flies. A bun tray (46 by 66 by 2.5 cm) containing
10.8 kg of adult diet (50% nutria meat and 50% honey
covered with a thin layer of a 50 : 50 mixture of cot¬

tonseed hulls and rice hulls) is provided for each cage.

The Mexican program uses horsemeat, instead of nutria,
in the same proportions. The cages are also stocked with
10 containers of honey (500-ml-jar chick feeders) and 10
containers (1 liter each) of water. The cages are placed in
the colony room (26°-27° C, 50%-60% relative humidity,
13 hours light, 11 hours dark) and held IV3-IV3 days
before oviposition. Each shift, the cages are moved for¬
ward one step in a sequence that moves the cages around
the colony room to the oviposition room. At the Mission
mass-production facility, 7 cages are filled with eggs
(egged) each shift (21 shifts/week) for production of 200
million flies/week. More cages are used at Tuxtla Gutier¬
rez per shift for production of 300 million to 350 million
flies/week.

Mating occurs during the holding process; it begins about
1 day after emergence and is completed in about 5 days.
Oviposition occurs about 16 hours after the flies have
reached ovarian maturity (stage 10, according to Adams
and Reinecke 1979), which is about IV3-IV3 days after
emergence. The time required to reach maturity varies
slightly with each strain and usually shortens in the
laboratory. It would be useful in the mass-production
facility to egg the cages at the proper time after reaching
stage 10. But, under mass-production schedules, each
function must be performed with each shift; so, a group
of flies must be egged when eggs are needed whether or
not the flies are at the best time for oviposition.

For oviposition, the cages are equipped with oviposition
vats and boards and moved into a dark room (26°-27° C).
The oviposition vat, which is preheated, contains a foam-
rubber mat, egging attractant, egging boards, and a
lighting system. The aluminum vats (160 by 30 by 2.5
cm) are thermostatically controlled at 37°-38° C. The
foam-rubber mat is cut to fit and is saturated with the
egging attractant (6 liters/vat), which is a 50 : 50 mixture
of used liquid media and water with about 2 liters of
citrated whole blood added. The egging boards are a box¬
like arrangement of four l-by-4 boards (152 cm long) run¬
ning lengthwise and held together by a l-by-4 board (20
cm long) across each end. These boards are placed
edgewise on top of the rubber mat. The lighting arrange¬
ment (a hood reflector and three 7V2-watt light bulbs) is
placed above the egging boards. The lights and attract-
tant cause the flies to migrate to the boards, where
oviposition occurs. Flies are allowed to lay eggs for 4
hours. Then the cages and contents are placed into a cold-
room (3°-5° C) to inactivate the flies. Egging vats are
removed, and the flies are destroyed by freezing and
burning. Eggs are removed from the egging boards with
a spatula and prepared for incubation. Eggs are weighed
in 7-g batches (6 g at Tuxtla Gutierrez because the rear¬
ing vat is smaller), placed in moistened petri dishes, and
incubated at 25° C for 12 hours until hatched. After

194

hatch, the larvae are transferred to the starting room
where they are placed in starting pans for rearing.

Production

Mass production of screwworm flies has been described
by Graham and Dudley (1959), Smith (1960), and Baum-
hover et al. (1966). These descriptions were based on
technology used during the southeastern eradication pro¬
gram. In the current program, many of the techniques
are similar, but others have been changed because it is
larger and more complex than the earlier program. One of
these changes is the present use of liquid nutrient
medium for feeding the larvae. The liquid medium is
cheaper and more readily available than various types of
flesh. Both the southwestern and Mexican-American pro¬
grams use the liquid nutrient medium for larval rearing;
the only meat used in the rearing facilities is that used in
adult fly diets. The use of liquid medium has been
facilitated by the research of Gingrich (1964) and
Gingrich et al. (1971) and the modifications of the
medium resulting from the research done by H. E. Brown
and J. W. Snow (1978, 1979, and unpublished data). The
development of this medium simplified logistic problems
associated with acquiring and storing medium com¬
ponents, medium preparation and handling, feeding
schedules and procedures, and waste disposal.

Starting larvae

Eggs collected from oviposition cages are incubated in
petri dishes. After eclosion, the larvae are distributed
evenly in fiberglass pans (64 by 45.7 by 7.6 cm) rimmed
with dry whole egg containing about 2.5 cm of liquid
starting medium suspended on either cellulose acetate
fibers or cotton linters for support. The liquid starting
medium is suspended at a rate of 30 liters of medium to
0.92 kg of shredded cellulose acetate or 1.82 kg of cotton
linters. The starting medium used at Mission consists of
6% dried whole blood, 3% dried whole egg, 3% calf-milk
supplement, 87.8% water, and 0.2% Formalin (form¬
aldehyde) as a preservative. Starting medium used at
Tuxtla Gutierrez is the same as that used for general
rearing and consists of 8% dried whole blood, 3% dried
whole egg, 3% dried nonfat milk, 0.2% Formalin, and
85.8% water.

The starting pans are held in a room at 38° C and about
90% relative humidity for 40-44 hours. The larvae are
provided supplemental feed as needed. After the 40-44
hours, larval weights range from 5.5 to 7.5 mg, and lar¬
vae are ready for transfer to the mass-rearing floor. The
starting room provides a less hostile environment than
the mass-rearing floor, allows good early growth, and pro¬
vides for the production of larger larvae. Its use also
saves space on the rearing floor.

At Mission, the mass-rearing vats are covered with a
2.5-cm-thick cellulose acetate mat (0.92 kg) and saturated
with liquid nutrient medium (about 30 liters). The
medium (85.8% water, 8% dried whole blood, 3% dried
whole eggs, 3% dried nonfat milk or 3% calf-milk
replacer, and 0.2% Formalin) is prepared in a medium¬
mixing room outside the rearing plant and piped to the
rearing floor. The medium is added to the vats through
hoses with radiator-filler nozzles. The hoses lead from the
overhead medium-supply system. Rearing vats are pre¬
heated to about 37° C. The contents of the starting pans
(one pan per rearing vat) are distributed evenly on the
surface, and the larvae are allowed to work into the
medium. Vat temperature is maintained near 38° C. Dur¬
ing the first 16-20 hours, supplemental nutrient liquid is
provided as needed to prevent drying and excess larval
crawling and to provide added nutrients. Twenty hours
after larvae are put in the vats (corresponding to a larval
age of 60-64 hours) and at subsequent 4-hour intervals,
the spent medium is removed from the vats with a
specially designed vacuum system.

During the rearing process, larvae that leave the rearing
vats prematurely are swept up and returned for further
feeding. Those larvae that leave 44 hours or more after
being introduced into the vats are collected for pupation.
Larvae are collected by a flowing-water system that
transports them to a sump where they are pumped
through a separator that shakes water and larvae apart.
Larval feeding is ended after the larvae have been on the
rearing floor for 72 hours, and the vats are flooded with
water to make the last larvae crawl off. At Mission, the
rearing vats are moved over the water grates; at Tuxtla
Gutierrez, since the vats are fixed in place, the water is
turned on through a series of valves. Larval collection
continues until most larvae have left the vat; this occurs
about 80 hours after their introduction. The cellulose
acetate mats used at Mission are vacuumed dry and hung
over rails to collect the final few larvae; at Tuxtla Gutier¬
rez, the cotton linters are pressed dry. In both cases, the
support media are finally disposed of by heat treating or
burning. Larvae collected at the water-larvae separator
are placed in pans (same as starting pans) containing
about 3.5 cm of hardwood sawdust at a rate of 3
liters/pan for pupation. These pans are placed on a
monorail and conveyed to the pupation room. The larvae
and sawdust remain in the pupation room for 16 hours at
26° C and 75% relative humidity.

Pupal collection and handling

Pupae being removed from the pupation room are
separated from the sawdust by a shaker-screen separator.
The sawdust is reused, and the pupae are transported to
the pupal holding area by a slow-moving (1 m/min) con¬
veyor belt (0.5 by 20 m). The pupae and residual larvae

195

are passed under a lighted passageway, and the larvae
exit through the belt for a further pupation period of 4
hours and are then screened again. Pupae leaving the end
of the belt are collected in 2.5-liter batches in screened-
bottom metal trays and placed in the pupal holding room.
Those pupae to be used for colony maintenance are col¬
lected at this point. Those pupae to be irradiated are
placed in the pupal holding room at 26° C and 75%
relative humidity for 5 days before irradiation.

Irradiation

and packaging of pupae

Pupae are irradiated as near emergence as possible to pre¬
vent damage to the flies. The pupae are placed in a
5.1-liter perforated metal cylindrical canister (12.7 by 50.2
cm) and are irradiated in the Husman irradiator (Iso-
medesx, Parsippany, N.J.), which uses Ce 137 as the
radioactive source. The pupae receive a minimum dose of
5 krads to insure sterility. After irradiation, pupae are
transported to the field-operation section for packaging
before dispersal. At this point, samples of each canister
are taken for quality control of the irradiation process.

Controlling Product Quality
and Production Yield

To make sure that the screwworms produced will perform
as expected after they are released, quality is assessed in
several phases of the rearing program. The quality of the
insects being released is evaluated; as part of this prod¬
uct testing, the irradiation process is routinely assessed
to insure that the released insects are sterile. The quality
of the insects being used to maintain the stock colony is
also tested. And the quality and quantity of larvae pro¬
duced in the starting room is evaluated to make sure that
this vital part of the mass-rearing process is working
properly.

Quality of mass-produced

screwworm flies

Various evaluations are routinely performed to assess the
quality of mass-produced screwworm flies. Results from
these evaluations are used to adjust production handling
and distribution procedures to insure that high-quality
sterile flies are released. Pupal samples from each shift
(taken during the irradiation process) are weighed. The
pupal cases are removed to reveal the state of biological
development of the flies. This test indicates whether the
pupae have developed to within 24 hours of emergence at
the time of irradiation. If so, and if the emergence span
(hours taken for 90% of the flies to emerge) is known, the
proper time of irradiation can be set. Early irradiation
reduces emergence and longevity and affects flies in other

undesirable ways. Late irradiation does not allow enough
time for packaging, handling, and transportation to the
various areas for release and results in reduced
emergence and lifespan. An emergence span that requires
more than 48 hours means the flies are not uniform and
requires changing of the irradiation schedule.

Longevity indicates how long the flies will survive when
released. Longevity after release can be affected by ir¬
radiation (particularly if not timed properly) and length of
time in colonization as well as natural conditions such as
temperature, humidity, and availability of food and
water. Longevity is determined by hatching the flies at
27o_28o C in a cage containing food and water and then
computing the percentage of flies still surviving after 14
days (normally 70% -80% for mass-produced sterile flies).

The quality of the flies subjected to the release pro¬
cedures is evaluated by sampling the flies (one box per
planeload) before and after the distributing plane returns.
The emergence percentage and the percentage that fly
provide data for evaluating the effects of handling,
storage, and dispensing. Agility tests have also been used
to assess fly quality. In these tests, the flies are allowed
to emerge in a confined space, and the rate at which they
disperse is determined. Results from these tests have
been highly variable, and efforts are being made to im¬
prove the techniques to provide reliable data on the
dispersion ability of sterile flies.

Trap-back studies similar to that described by Ahrens et
al. (1976) have been useful in evaluating fly quality.

These evaluations are particularly valuable in assessing
new strains or when serious problems occur in the erad¬
ication program. These tests involve releasing marked
flies and determining migration, survival, and other
quality attributes by computing the percentage trapped
at various distances from the release site. These tests
provide data on movement and survival of mass-produced
screwworm flies and candidate strains to be released.

Such tests require much personnel and time for adequate
execution, so they are not routinely performed.

A good test of the quality of released sterile flies is ob¬
tained by sterility evaluations similar to that reported by
Coppedge et al. (1980). These tests are conducted on pro¬
duction and candidate strains to assess their migration,
survival, and mating capabilities. The tests determine
how much the fertility of egg masses collected in test
areas has been reduced. Strains are released in areas of
high occurrence of screwworm infestations. Traps are
deployed and trap-catch data are collected to determine
the migration and survival of the released flies and occur¬
rence of wild flies. Sentinel sheep pens, containing two to
three sheep with artificially inflicted wounds, are main¬
tained; eggs laid on the wounds are collected; and the per-

196

centage of fertility is determined. At the same time, egg
masses are collected from animals in the area that have
been wounded during ranching operations; the percentage
of fertility of these eggs is compared to the percentage
determined for eggs taken from the artificially wounded
sheep. Fly quality can be measured by comparing the re¬
sults from the test area to those obtained in control areas
or to those obtained from a different group or strain of
flies released in the same area. Changes in patterns of fly
catches and sterility percentages also indicate changes in
fly quality. This type of evaluation is being used in the
Sinaloa State of Mexico to assess the quality of flies be¬
ing mass-produced and released.

Whitten (1980) has used the isozyme technique to assess
the quality of various candidate strains of flies to be used
in mass production. Bush et al. (1976) suggested that the
electrophoretic examination of the isozymes ofa-
glycerophosphate dehydrogenase could be used to deter¬
mine changes in the genetics of mass-reared flies. This
technique has been used in the Mission laboratory to
assess the quality of flies and particularly to monitor the
changes occurring during long-term colonization. Present¬
ly, this technique is being used in the mass-rearing fa¬
cilities as another criterion for assessing fly quality.

Goodenough et al. (1978) used an electroretinographic
technique to evaluate visual sensitivity of four strains of
screwworm flies. They found that the visual sensitivity
and the distribution of visual responses of newly colonized
strains were nearer than those of older strains to those of
native wild flies. So, visual responses can be used to
assess the quality of mass-reared flies and to evaluate the
degradation of the flies during long-term colonization.
These evaluations have been started by the mass-
production facilities to assess fly quality.

Quality of the irradiation process

Assessment of the irradiation process is made from 15
pupae randomly selected from each group being ir¬
radiated. After adult eclosion, the flies are allowed to
mature and mate; any later sign of oviposition or ovarian
development indicates the irradiation dosage was not am¬
ple for female sterilization. But the female requires about
twice the amount of irradiation for sterilization as the
male. Male screwworm flies require about 2.5 krads for
sterilization (Bushland and Hopkins 1953); so the min¬
imum dosage from the irradiators is set at 5 krads.
Although evaluating ovarian development is a possible
measure for quality control, it is a time-consuming pro¬
cedure and requires skill in distinguishing various stages
of ovarian development. The technique does not provide
on-the-spot assessment of irradiation quality.

Quality of the adult fly colony

Quality of the brood colony is assessed by the volume
and percentage of hatch of eggs harvested from each col¬
ony cage. When problems occur with oviposition, the rate
of sexual maturation in the colony is determined by
dissecting samples of females at various stages. But this
procedure is limited by available personnel and the quan¬
tity of flies that need to be examined.

Larval quality and yield

The quality of the larvae from the starting room is
assessed by larval weights, which indicate the quality of
the starting-room techniques. A larval weight of 6 mg or
more is desired at the end of the starting period to insure
proper growth under mass-rearing conditions. Alterations
in larval density in the starting pans and modifications of
the larval diets affect larval weight. Larvae resulting
from 7 g of eggs per pan reach a weight of 6-7 mg at the
end of 40 hours of growth after hatch. The weight can be
increased to 7-8 mg by reducing the quantity of eggs per
pan to 5 g and to 10-12 mg by also increasing the blood
level in the diet to 8% and using 3% dried nonfat milk in¬
stead of calf-milk replacer (H. E. Brown, unpublished
data).

The quality of the larvae produced in the mass-rearing
process is also assessed by larval weights coupled with
larval yields. These measurements give a good indication
of the overall success of the mass-rearing process. At¬
tempts are made to keep larval weight above 60 mg at
the end of the rearing process. Alley and Hightower
(1966), and Hightower et al. (1972) showed that larger
male flies have a higher mating frequency and that flies
resulting from larvae weighing less than 56-60 mg are in¬
ferior and cannot compete with larger flies. This
minimum has been maintained as a standard over the
years of production. When a strain has been mass-
produced for several months, the larval weights decrease;
but, after introduction of new strains, larval weights in¬
crease in most instances. The DE-9 strain (a strain pro¬
duced from egg masses collected near Aldama, Tamauli-
pas, Mexico), which was recently introduced into the rear¬
ing program, produces weekly average larval weights of
72.8 mg each (D. D. Wilson, personal communication).
This is the largest average larval weight that has been
produced in the program’s existence. Larval weights from
the previously produced Arieruz strain were averaging
slightly above the 60-mg minimum just before the intro¬
duction of the DE-9 strain.

Larval weight is also used as the criterion for evaluating
various experimental conditions and diets for possible use
in the mass-production program. If the new technique or
component being tested produces larvae as large as the

197

standard, then it is considered suitable for mass produc¬
tion of screwworms. Although larval weight may not be
the best criterion for assessing quality, it is the best
within the limitations of the current program’s facilities.

Another way of insuring that the production system is
working properly is to assess yields of larvae and pupae.
Predicted mass-production levels are based on experience,
which has shown that 7 g of eggs should yield 12-13
liters of larvae, assuming that proper techniques are
followed and that egg hatch is 95% or better. A reduction
in yield may indicate low fertility, problems with rearing
techniques, or low quality of dietary ingredients used in
the nutrient medium. The cause of the reduction should
be found and remedied.

References

Adams, T. S., and Reinecke, J. P.

1979. The reproductive physiology of the screwworm,
Cochliomyia hominivorax. I. Oogenesis. J. Med.
Entomol. 15: 472-483.

Ahrens, E. H.; Hofmann, H. C.; Goodenough, J. L.; and
Petersen, H. D.

1976. A field comparison of two strains of sterilized
screwworm flies. J. Med. Entomol. 12: 691-694.
Alley, D. A., and Hightower, B. G.

1966. Mating behavior of the screwworm fly as af¬
fected by differences in strain and size. J. Econ.
Entomol. 59: 1499-1502.

Baumhover, A. H.; Graham, A. J.; Bitter, B. A.; Hopkins,
D. E.; New, W. D.; Dudley, F. H.; and Bushland, R. C.

1955. Screwworm control through release of sterilized
flies. J. Econ. Entomol. 48: 462-466.

Baumhover, A. H.; Husman, C. N.; and Graham, A. J.

1966. Screwworms. In C. N. Smith (ed.). Insect Col¬
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Academic Press, New York.

Brown, H. E., and Snow, J. W.

1978. Protein utilization by screwworm larvae
(Diptera : Calliphoridae) reared on liquid
medium. J. Med. Entomol. 14: 531-533.

1979. Screwworms (Diptera : Calliphoridae): a new
liquid medium for rearing screwworm larvae. J.
Med. Entomol. 16: 29-32.

Bush, G. L.; Neck, R. W.; and Kitto, G. B.

1976. Screwworm eradication: inadvertent selection of
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Science 193: 491-493.

1975. Screwworm research and eradication. Bull. En¬
tomol. Soc. Am. 21: 23-26.

Bushland, R. C., and Hopkins, D. E.

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1964. Nutritional studies on screwworm larvae with
chemically defined media. Ann. Entomol. Soc.
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Gingrich, R. E.; Graham, A. J.; and Hightower, B. G.

1971. Media containing liquified nutrients for mass¬
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Goodenough, J. L.; Wilson, D. D.; and Agee, H. R.

1977. Electroretinographic measurements for com¬
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reared screwworm flies, Cochliomyia
hominivorax (Diptera : Calliphoridae). J. Med.
Entomol. 14: 309-312.

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1978. Visual sensitivity of four strains of screwworm
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Graham, A. J., and Dudley, F. H.

1959. Culture methods for mass rearing of screw¬
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1972. Growth and critical size at pupation for larvae
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1955. Possibilities of insect control or eradication
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Improved Techniques

for Mass Rearing Anopheles albimanus

By Donald L. Bailey and J. A. Seawright1

Introduction

Anopheles albimanus Wiedemann is an important vector
of human malaria over most of Central America and part
of South America. Rozeboom (1936) was the first to
colonize An. albimanus, and few improvements to his sys¬
tem were reported until Ford and Green (1972) published
techniques developed during their 2-year study. In the
early 1970’s, the U.S. Agricultural Research Service’s In¬
sects Affecting Man and Animals Research Laboratory at
Gainesville, Fla., began intensive studies on the feasibil¬
ity of releasing sterile males for the control of An.
albimanus in El Salvador. Not until then was there a
need for further improvement of the techniques for mass
producing this species. This paper describes all the im¬
provements made since that program began.

Colonization

When Rozeboom (1936) established a colony from field-
collected larvae supplemented with field-collected adults,
he had problems with high adult mortality and low ovi-
position rates. And Dame et al. (1974) reported a very
slow rate of colonization in a colony of An. albimanus
from about 11,000 adult females collected from stables in
El Salvador. They attributed this slow colonization to a
low oviposition rate caused by a low level of insemination
of females in filial generations.

During the recent pilot test of sterile-male release in El
Salvador, the field performance of mass-reared sterile
males (first colonized in 1975) had to be assessed
periodically by comparison with a recently colonized field
strain. So, to avoid the colonization problems reported by
these previous researchers, Bailey, Lowe, and Kaiser
(1980) developed an improved system for rapid coloniza¬
tion of An. albimanus. This new system required a
minimum of effort and material.

Researchers collected adult females from a stable for 10
days. Each day they placed 50 of the collected females in
5-dram plastic vials containing 5 ml of water infused with

'Research entomologists, Insects Affecting Man and Animals
Research Laboratory, Agricultural Research Service, U.S. De¬
partment of Agriculture, P.O. Box 14565, Gainesville, Fla.
32604.

a small amount of liver powder and dried yeast. The vials
were held for 4 days; then the eggs produced in each vial
were counted. This system was compared with one where
50 females were placed in a cage (61 by 61 by 61 cm) each
day for 10 days; the number of eggs produced was
counted. With both systems, 500 adult female mosquitoes
were used. The eggs collected from the vials and from the
cages were reared to the pupal stage and placed in
separate adult cages. Eggs were collected from the F,
adults that emerged, and the numbers were calculated.
The 500 parent females in vials produced 37,438 eggs
with 67% hatch. The F, adults produced 207,359 eggs
with 63% hatch. This was a 550% increase and indicated
that more than 1 million F3 eggs would have been pos¬
sible. The 500 parent females in cages produced only 334
eggs, and the F, adults did not oviposit.

Genetic Sexing System

The success of the sterile-male method depends on the ef¬
ficient distribution of sufficient numbers of competitive,
sterile males into the habitat of the target species. A
sound system must be available for the mass production
of sterile males. For the pilot test with An. albimanus in
El Salvador, an efficient method of separating males and
females was also needed. (Since females of An. albimanus
are potential malaria vectors, their release must be
limited even though they are sterilized).

Dame et al. (1974) reported a mechanical method of
separating males and females based on pupal size. Lowe
et al. (1981) tried to use a membrane filled with
malathion-laden blood for the preferential killing of adult
females. Neither method was satisfactory for a large pilot
project. The mechanical method caused significant losses
(up to 40%) of males, and the membrane method was a
problem because of the damage done to the insects dur¬
ing the handling of large numbers (about 1 million/day) of
adult mosquitoes. So, we developed a genetic method for
the preferential killing of females. We knew that if such a
genetic method were effective during the egg or first lar¬
val stage, the cost of producing sterile males could be re¬
duced by about one-half because twice as many males
could be produced with the same space, food, and
personnel.

The genetic sexing system that we synthesized used pro-
poxur susceptibility (prs ) as a conditional lethal (dies

200

when treated with propoxur2) trait, a T(Y;2R ) transloca¬
tion, and an In(2R ) inversion. The locus for propoxur
resistance (pr), which is dominant, is on the right arm of
chromosome two; this allele was linked to the Y
chromosome via a radiation-induced translocation (an in¬
terchange of chromosomal pieces between nonhomologous
chromosomes). We induced six different T{Y;2R) trans¬
locations. Since crossing over occurs in both sexes of An.
albimanus, recombinant, resistant females were present in
each generation. To suppress recombination, we ir¬
radiated males of the translocation stocks to induce an in¬
version (which reverses a section of a chromosome). Nine
genetic sexing strains with <2.5% resistant, recombinant
females were detected (Kaiser et al. 1978, Seawright et al.
1978). One of these strains, MACHO, was then produced
at the mass-rearing plant in El Salvador. After a few
problems in the early phase of implementation (Bailey,
Lowe, et al. 1980), MACHO was a complete success. We
were able to kill the females of the strain by treating the
eggs.

Before the MACHO strain was available, we were releas¬
ing an average of 170,000 sterile males per day in El
Salvador. The average number of males produced was
much greater than 170,000, but inefficient separation
methods drastically reduced the number available for
sterilization and release. With the MACHO strain, the
average release during the last year of the study was
954,400 males per day, and these males were produced
with about the same resources as the 170,000 daily
average for the regular strain. Also, 99.9% of the mos¬
quitoes produced for release were males.

The synthesis of the MACHO strain is a good example of
the way genetic principles can be used in the mass pro¬
duction of insects. In our case, we were attempting to
maximize the production of sterile males, but selective
breeding schemes can be used to enhance the mass pro¬
duction of insects for any purpose. In fact, many of the
problems encountered in mass-production facilities can be
solved by genetic approaches.

Production
Production of adults

For the pilot test in El Salvador, we produced about 1
million MACHO sterile males each day. The adult cages
and the stocking techniques used to mass-rear An.
albimanus were those reported by Bailey, Lowe, et al.
(1980). They used 126 cages (61 by 61 by 61 cm), made of
aluminum window-screen framing and covered on four
sides with nylon screen (8 mesh/cm). The inside surface of

2o-I sopropoxyphenyl methylcarbamate.

the top and bottom of the cage was made of white For¬
mica for an easily cleanable surface. Access into the cage
was provided by a sleeve made of a 75-cm length of
25-cm-diameter surgical tubing with one end attached to
a 30- by 60-cm opening in the front. At first, the cages
were stocked with about 6,000 pupae; then, about 4,000
pupae were added every 2 days for 1 month. After a
month in production, each cage was removed, cleaned,
and restocked. Four cages were cleaned and restocked
each day to minimize the number out of production at
one time.

The most important recent advance in maintenance of
adult colonies has been the development of a system for
feeding of preserved bovine blood through natural animal
membranes (Bailey et al. 1978). For our colony mainten¬
ance, 61 liters of fresh blood were collected 3 days a
week, defibrinated mechanically, and stored in a
refrigerator at 5°±2° C. Blood that was to be used for
feeding was removed from the refrigerator and placed in
condoms made of sheep intestinal membrane (150
ml/membrane). The membranes were closed with
clothespins and heated to 44° C in a water bath. A mem¬
brane with heated blood was then placed in each of two
feeding ports made of 4-inch polyvinyl chloride plastic
pipe and tube gauze. The blood was removed, reheated,
and returned to the ports twice during the day for a total
of three feedings (at 0800, 1100, and 1400 hours). Then
the membranes were discarded. Cotton pads saturated
with 10% sugar water were provided at all times in two
other feeding ports. These ports eliminated the need for
entering the cages for feeding, thus minimizing the
escape of adults. The use of membranes to feed adult An.
albimanus has eliminated the need to maintain about 40
rabbits for feeding the adult colonies. At first, a reduc¬
tion in egg production was associated with the preserved
blood; but, in time, egg production returned to a normal
level.

Production of eggs

Eggs deposited overnight in plastic pans (15 cm diameter
and 8 cm deep) containing 2 cm of water (one pan per
cage) were collected by pouring the water containing the
eggs through nylon screen (12 mesh/cm) to remove dead
mosquitoes and other debris. About 90% of the eggs were
then poured into a 0.01% propoxur solution and held for
24 hours to kill the females. The remaining 10% were left
untreated and used for colony replacement.

In earlier rearing of An. albimanus, the eggs were held an
additional 24 hours to hatch, and numbers of the hatched
larvae were estimated and the larvae placed in rearing
trays. Dame et al. (1978) described an improvement in
which the eggs were dried after the first 24 hours and
samples were measured volumetricaily to obtain accurate

201

numbers of larvae in the rearing trays. The eggs were
dried in plastic trays (25 by 18 by 7.5 cm high) with a
bottom of white nylon cloth by drawing air through the
tray with a fan for 30 minutes. Dried eggs were sifted
through stainless steel screen (40 mesh/cm) to break up
clumps. Then a vibrating device was used to measure
them volumetrically into plastic microcentrifuge tubes
cut to length to hold the desired quantity.

With the standard strain of An. albimanus, we needed
0.085 ml of eggs (average of 6,779 eggs) per rearing tray.
But the MACHO strain is naturally 50% sterile (eggs do
not hatch) because of the translocation; so we needed 0.17
ml of untreated eggs to achieve equivalent production.
Since the females (99.9%) had been killed with the egg
treatment, we needed 0.34 ml of treated eggs per sample.
These samples were then placed in individual 350-ml
Styrofoam cups containing 75 ml water and 1.4 ml of a
2% liver and yeast suspension (1 : 1). These cups were
held for 24 hours at 29°±0.5° C for the eggs to hatch.

Excess eggs were stored in 100-ml plastic bottles at 10° C
(Bailey, Thomas, et al. 1979), so a stockpile was available
for use in case of an unexpected reduction in egg produc¬
tion. Since untreated eggs can be stored for 7-10 days
with little reduction in hatch, they can be used for colony
maintenance or treated later with propoxur to rear stock
for field release. Treated eggs can only be stored for 2-3
days without reduced hatch.

Production of larvae

The larval rearing trays used for mass production were
made of ABS plastic (56 by 43 by 7.5 cm high), and each
contained 3 liters of water. A precise temperature-control
system using electrical heating tapes and electronic pro¬
portional controllers (Dame et al. 1978) was devised. The
shelves holding the rearing trays were 15 m long and ar¬
ranged in banks of nine. Each shelf had one 30-m heating
tape that ran the full length of the shelf and doubled
back again; the two strands were 30 cm apart. One con¬
troller regulated the tapes on three shelves; so three con¬
trollers were used for each bank. With this system, the
water temperature in the rearing trays could be main¬
tained at 29°±0.5° C.

The trays containing water (300 containing propoxur-
treated eggs and 104 containing untreated eggs) were
placed on the shelves for 24 hours so the water
temperature would stabilize at 29°±0.5° C before the
newly hatched larvae from the hatch cups were poured in.
Just before the larvae were introduced, 150 ml of a liver
powder, yeast, and 40% protein hog supplement (1:1:1)
food suspension (2.25 g dry ingredients) was added to
each tray. The larvae received an identical feeding 72
hours later. On the next 2 days, each tray received 150

ml (3 g dry ingredients) of a suspension of hog supple¬
ment alone.

Production of pupae

On the day after the last larval feeding (day 6 after the
larvae were placed in trays), pupae were removed. The re¬
maining larvae were consolidated (four trays in one) and
returned to the rearing shelves with the original water
(Fowler et al. 1980). The next day, pupae were again
removed, but this time any remaining larvae were
discarded. The pupae were removed from the rearing
trays by a system, described by Bailey, Lowe, et al.
(1980), adapted from Weathersby (1963). A mixture of lar¬
vae and pupae was placed in a large plastic funnel that
contained cold water (10° C) and had a shutoff valve at¬
tached to the outlet. Larvae sank to the bottom in the
cold water, and the pupae remained on the surface. When
the valve was opened, the larvae passed through first and
were collected on a screen; then they could be returned to
the rearing trays or discarded. The pupae were then col¬
lected on a screen and used to restock rearing cages or
were sterilized for field release depending on whether
they had been treated with propoxur. The consolidation
of trays of larvae greatly increased efficiency by increas¬
ing the number of trays that could be set in a given
space and by decreasing the number of trays handled
during the second harvest. The quality of the insects was
not reduced by this system.

Production Management

In our rearing program, we had to monitor environment,
maintain specified production levels, and insure that our
product insects met the standards set for their use in col¬
ony maintenance and sterile-male release. The laboratory
staff in El Salvador consisted of two entomologists, a pro¬
gram assistant (who collected, compiled, and organized
the data), an administrative assistant (who organized
work schedules and coordinated responsibilities of the
technical staff), and 26 technicians. The technical staff
was divided among five sections responsible for adults,
eggs, larvae, pupae, and special research. Each section
had a supervisor and an assistant supervisor and the
necessary personnel to perform the duties. Work
schedules were arranged so production was possible for a
full 7-day week. The supervisor of each section was
responsible for daily reports on that section’s function.

Environmental monitoring

A hygrothermograph was operated constantly in the
adult colony. Each day, the supervisor of the adult sec¬
tion reported the temperature and humidity at 2400,

0800, 1200, and 1600 hours. He also reported the
temperature of the refrigerator where the preserved blood

202

for feeding the adults was stored. The supervisor of the
egg section reported the temperature of the water in
which the eggs were incubated and also the temperature
in the refrigeration unit where the extra dried eggs were
stored. The supervisor of the larval section reported the
water temperature in the rearing trays at each level for
all the rearing shelves at 1000 hours each day. The super¬
visor of the pupal section periodically checked and
reported the temperature of the cold water for pupal
separation. These daily reports were absolutely essential
to alert the entomologist in charge of any malfunction in
the environmental control equipment. If corrections were
made soon enough, there was usually little or no loss in
production.

Maintaining production levels

The supervisor in each section was also responsible for
reporting the production levels in his section each day.
For the adult section, this report included the total
number of cages of adults, number of cages stocked with
pupae, number of pupae per cage, and number of cages
cleaned and restocked. The egg section reported the
number of cages producing eggs, total egg production
(ml), average egg production per cage (ml), number of
hatch cups set, amount of extra eggs stored (ml), and
number of rearing trays set. The supervisor of the larval
feeding section reported the number of trays receiving
food (first through fourth feedings) and the total number
of bad trays (those trays with total or almost total mor¬
tality by the fourth day). He was also responsible for col¬
lecting random samples of the food mixture during
feeding and for running sedimentation tests on the
samples to determine whether the food was being for¬
mulated and mixed properly. The supervisor of the pupal-
handling section reported the total number of trays
harvested (first and second harvests), the average pupal
production per tray (based on 3 randomly selected trays
from each harvest each day), the relative size of the
pupae (average number of pupae in 3 samples of 1 ml
each), and the sex ratio from each harvest (based on 3
samples of 100 pupae each).

Quality Control

High yields are useless if the insect does not survive, or
if it does not mate once released. In the El Salvador proj¬
ect, quality was judged with tests done in the laboratory
and in the field.

Laboratory tests

Because a genetically altered strain was being used, cer¬
tain checks were made to insure the integrity of the
genetic sexing system. For example, the percentage of
egg hatch (treated and untreated) was checked daily. The

expected untreated hatch was 50% (because of the trans¬
location) and the expected treated hatch was 25%

(because of deaths among females). Another indicator was
sex ratio, and we made a daily check of the sex ratio of
the pupae reared from propoxur-treated eggs. If the
system was working properly, the pupae from these trays
were about 99.9% males. But about 0.1% resistant
females were produced through genetic recombination in
each generation in the MACHO strain, so the number of
these females was expected to increase over time. Also,
at weekly intervals, adult males and females that had not
been treated in the egg stage were exposed to residues of
0.1% propoxur on filter paper for 1 hour, and mortality
was recorded. The data were used to monitor levels of
resistance and susceptibility in the males and females.
When the frequency of resistant, recombinant females
became excessive, they had to be purged by crossing
propoxur-resistant translocation males with females that
were totally susceptible. This nucleus colony was then in¬
creased for three or four generations until it was large
enough to be used to replace the old colony. Also, if the
MACHO strain was accidentally contaminated with nor¬
mally susceptible stock, the increase of susceptible males
accelerated rapidly.

Each day, a sample of 300 pupae from the rearing trays
was placed in a cage, adults were allowed to emerge for
48 hours, and the percentage emerging was recorded. The
adults were then constantly provided with 10% sugar
water on cotton pads and held at 27°±2° C, and the
percentage of the males and females surviving after 7
days was reported. This system indicated whether the in¬
sects being produced were healthy. Also, some males that
had been sterilized for field release and some nonsterile
males were crossed with normal females each day. The
percentage of egg hatch from those crosses was calcu¬
lated to determine how effective sterilization techniques
had been.

Field tests

Adult emergence in the field was monitored daily. But
the most important test— the determination of the com¬
petitiveness of the laboratory-reared, genetically altered,
chemically sterilized male— could only be made by using
comparative field releases. For this, we selected part of
the 30-km2 test area that had an extremely small natural
population of An. albimanus (Kaiser et al. 1979, 1981).
The comparison had the sterile MACHO males and males
of a recently colonized field strain, CAMPO, compete for
released CAMPO females. The releases consisted of two
replicates, each at a ratio of about five sterile MACHO
males to one CAMPO male to one CAMPO female. About
10,000 females were released per replicate. Pupae were
packaged for transport to the release site according to

203

the method described by Bailey, Lowe, et al. (1979). Calf-
baited traps (Lowe and Bailey 1981) were placed about
125 m north and south of the release site. A calf was
placed in each trap at 1800 hours and removed the next
morning. Adult An. albimanus were collected each day
from each trap, and the females were returned to the
laboratory. Males were excluded to prevent further
matings in the cages.

Females were held in the laboratory for 2 days and then
immobilized in a coldroom and placed in 5-dram plastic
vials, one female per vial. Five ml of water was added,
and the females allowed to oviposit. After 5 days, the
eggs were checked for hatch to determine whether the
mating had been with a sterile or fertile male. Com¬
petitiveness was determined by the formula (Fried 1971)
that

C=S//V(calculated)— S/A/lactual),

where S/7V(calculated) = the ratio of irradiated males to

normal males that will give an ex¬
pected percentage of sterility if
mating NQ X Nd gives >5%
hatch and if mating N 9 X Sd
gives 0%-5% hatch;

and S/TViactual) = the actual ratio used experi¬

mentally.

The average mating competitiveness of the MACHO
males was 78.5% of that of the CAMPO males; their
average dispersal ability ws 74%. So the males of the
genetic-sexing strain could disperse and could induce
high levels of sterility in indigenous populations.

Conclusion

The rearing techniques described in this paper have been
refined through a great amount of research effort. The re¬
quirements for each life stage in the mass production of
An. albimanus have narrow limits, and each is critical to
the successful rearing of this species. One factor is not
necessarily more critical than another, because any single
weakness or deficiency in the system can cause the entire
system to fail.

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Seawright, J. A.; Kaiser, P. E.; Dame, D. A.; and
Lofgren, C. S.

1978. Genetic method for the preferential elimination
of females of Anopheles albimanus. Science
200: 1303-1304.

Weathersby, A. B.

1963. Harvesting mosquito pupae with cold water.
Mosq. News 23: 249-251.

205

Some Systems for Production
of Eight Entomophagous Arthropods

By E. G. King1 and R. K. Morrison2

Introduction

Arthropod rearing is basic to most entomological endeav¬
ors, especially biological control. For example, im¬
porting entomophagous arthropods (predators and
parasites, the “natural enemies”) for establishment often
requires that these organisms be reproduced for several
generations in quarantine before release. Even after
establishment, large-scale production in insectaries may
be necessary to expand the entomophage’s geographic
distribution. Likewise, conduct of today’s integrated pest
management programs requires detailed knowledge of the
entomophage’s biology, behavior, and effectiveness.
Availability of these organisms for research often
depends on our ability to maintain colonies of them in
the laboratory. Finally, control of pest arthropods by
augmenting their predators and parasites through
periodic releases also requires large-scale production. So
specialized equipment and unique techniques have been
developed for producing hosts, because artificial diets
and in vitro rearing methods have generally not been
developed for large-scale production of predators and
parasites.

Methods for rearing many kinds of entomophagous ar¬
thropods have been reviewed by Finney and Fisher (1964)
and Morrison and King (1977), as have facilities (in¬
cluding quarantine) and specialized equipment for rearing
entomophages and their hosts by Fisher and Finney
(1964) and Leppla and Ashley (1978). But evaluating a
rearing program requires consideration of many factors
besides production capability, especially product quality
(Boiler and Chambers 1977).

Here we examine several systems currently being used
for producing entomophagous arthropods, and we com¬
ment on the strengths and weaknesses of each system.
We will mainly discuss these predator and parasite
species: Aphytis melinus DeBach; the common green
lacewing, Chrysopa camea Stephens; Cryptolaemus mon-
trouzieri Mulsant; Encarsia formosa Cahan; Lixophaga

'Research entomologist, Southern Field Crop Insect Manage¬
ment Laboratory, Agricultural Research Service, U.S. Depart¬
ment of Agriculture, P.O. Box 225, Stoneville, Miss. 38776.

2Research entomologist, Cotton Insects Research Unit,
Agricultural Research Service, U.S. Department of Agriculture,
P.O. Box DG, College Station, Tex. 77841.

diatraeae (Townsend); Spalangia endius Walker;
Phytoseiulus persimilis Athias-Henriot; and Tricho gram¬
ma sp. These are being mass-reared and used for control
of their natural hosts by augmentative releases in various
parts of the world. We also briefly discuss the use of
natural or unnatural hosts (those not normally attacked
in nature, but suitable in the insectary) for laboratory
rearing of entomophages and survey recent accomplish¬
ments in the development of artificial diets for them.

Systems for Producing
Entomophages

Production of Aphytis melinus

A. melinus is reared continually and released regularly
for control of the California red scale, Aonidiella aurantii
(Maskell), in the citrus-growing areas of California (Penn¬
ington 1975). DeBach et al. (1950) initially demonstrated
the potential of controlling California red scale by
augmentative releases of A. chrysomphali, and DeBach et
al. (1955) further defined it. After the field demonstration,
procedures were developed for mass producing A.
chrysomphali on California red scale fed potato tubers.
But the method developed by DeBach and White (1960)
that uses the unnatural host— oleander scale, Aspidiotus
nerii Bouche— for production of the California red scale
parasite, A. lingnanensis Compere, was better and is still
used by at least two commercial insectaries in California.
The imported species A. melinus is now reared and re¬
leased instead of A. linganensis, but the procedures re¬
main essentially the same as those described by DeBach
and White (1960).

A. melinus is produced (fig. 1) on a uniparental strain of
the oleander scale fed banana squash, Cucurbita maxima
Dend. Each day, crawlers (the immature mobile stage of
oleander scale) are implanted on fresh squash. These
crawlers are collected from squash that have been in¬
fested for 58-73 days. Infested squash about 73 days old
are offered to the parasite in oviposition-collection units
supplied with honey, the food for the adult parasites. A
specified number of freshly emerged and collected adult
parasites is placed on the unparasitized squash, and the
oviposition-collection unit is closed. After 24 hours for
oviposition, the parasites in the unit are anesthetized,
removed, and prepared for field release. The now-
parasitized squash are moved from the unit to storage
racks for parasite development. Removing the squash
also allows continual use of the oviposition-collection

206

Figure 1.— System for producing A. melinus
(parasite) on A. nerii (host) fed banana squash.

unit. About 2 or 3 days before emergence of adult
parasites, the squash are placed back into a prepared
unit, and the parasites are collected each 24 hours
throughout the emergence cycle.

Extensive tests were conducted in selecting the host
(oleander scale) and its food source (banana squash) by
DeBach and White (1960) and Finney and Fisher (1964).
No special problems remained after this host was chosen,
but careful attention has to be given in the selection,
storage, and handling of the squash. DeBach and White’s
(1960) production program is sound and functional,
though rearing procedures could undoubtedly be auto¬

mated, particularly in handling of materials. The system
has had no major engineering input, probably because it
has production facilities at each of the major citrus¬
growing areas instead of one large facility at a central
location.

Apparently, rigid procedures have not been developed
and implemented for monitoring and maintaining the
quality of A. melinus. But genetic variability in the genus
Aphytis is fully recognized in California and has been ex¬
ploited by researchers in establishing species of Aphytis
effective against the California red scale. And DeBach
and Hagen (1964) reported that strains of A. lingnanensis
had been selected for increased tolerance to cold and
heat. Use of these strains could improve their effec¬
tiveness in augmentation programs. Basic measurements
of production efficiency, which can warn of possible
breakdowns in insect quality, are built into the produc¬
tion system. And the sex ratio is routinely measured.

Production of Chrysopa carnea

Rearing and periodically releasing C. carnea to prey on
many insect pests has been very effective (Beglyarov and
Smetnik 1977, Ridgway et al. 1977). Its effectiveness
may be improved by use of supplementary foods contain¬
ing materials that attract, retain, and stimulate the
adults to oviposit (Hagen and Hale 1974). Mass produc¬
tion of C. carnea is limited by its costliness, though
several artificial diets have been developed for rearing the
larvae (Singh 1977), and methods for encapsulating some
of these diets have been developed by Hagen and Tassan
(1965) and Martin et al. (1978; see also Agricultural
Research 1971). But eggs of the Angoumois grain moth,
Sitotroga cerealella (Olivier), or eggs of other hosts, re¬
main the preferred larval food source (Beglyarov and
Smetnik 1977, Morrison and King 1977) because they
provide the best balanced and most available diet. An in¬
expensive, highly acceptable adult diet that induces and
maintains high rates of fecundity has been developed by
Hagen and Tassan (1970). But the Wheast in this diet is
no longer available and has been replaced with a com¬
parable product— Formula 57 available from CRS Co., St.
Paul, Minn. (K. S. Hagen, personal communication).

The basic technique for production of adult C. carnea
from eggs (fig. 2) evolved from the system first reported
by Finney (1948), later improved on by Rincon-Vitova In¬
sectaries, and further developed by Morrison and
Ridgway (1976) and Morrison (1977). Food (frozen
Angoumois grain moth eggs) is mixed with fully em-
bryonated C. carnea eggs and distributed into multicell
rearing units. The cells are covered on both sides with
organdie, which the hatched larvae can feed through dur¬
ing three later feedings. Morrison (1977) reported coating
glass, cut the same size as the unit, with a honey-water

207

L

PHOOUCTIO * AMO
OUAIITY COMTPOl

at comps

— *1 Y"t oviaosmotJ

,rs] •"¥'«»« *«u»/

ADO OVIPOS- 1 ioviaosi
ITIOMPAPta | "\(FttDA

OVIPOSITION
HOLD)

/-ssfesr-N

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Figure 2.— System for producing C. camea (predator) on frozen Angoumois grain moth (AGM) eggs.

mixture (1:1) and Angoumois grain moth eggs for
feeding the larvae. This plate is inverted over the cloth.

After pupation, the organdie is removed from the cells,
and the pupae are placed together for emergence. As the
adults emerge, they are collected daily, held in preoviposi-
tion units, and supplied adult diet and water for 4 days.
Thereafter, the units are lined with paper on which the
adults lay their eggs. This paper is coated with the adult
diet, a combination of Formula 57, sugar, and water. (A
suitable diet must be continuously available to maintain
high fecundity.) When 0-to-24-hour-old eggs are present,
the paper liners are then removed and replaced daily.
Morrison and Ridgway (1976) used a vacuum to im¬
mobilize the adults and then transfer them to clean units
for each change of oviposition paper. The kraft-paper

liners and attached eggs are removed from the oviposi¬
tion units and held overnight in room conditions (about
22° C) so the eggs will harden. When a hand-held ball of
nylon net is wiped over the paper oviposition sheets, the
hard nylon threads of the net easily break the egg stalk,
and the eggs come off. The collected eggs can be used for
maintenance of the colony or for rearing larvae to
distribute in the field. For field distribution, the C. camea
eggs are mixed with sawdust and frozen Angoumois
grain moth eggs; the mixture is held until the resulting
larvae are 1-3 days old; and then the larvae are
mechanically distributed (Agricultural Research 1972).

Production efficiency is continually monitored, but
methods for measuring quality have not been defined.
Tests have shown that adult survival and fecundity does

208

Figure 3.— System for producing C. mon-
trouzieri (predator) on P. citrii (prey) fed potato
sprouts.

209

decrease with increasing time in culture. Also, Jones et
al. (1978) found indications that developmental time of
immature stages increases, and egg viability, food con¬
sumption, and searching ability of C. camea decreases
with increasing time in culture. So they recommended
that C. camea intended to augment natural populations
should not be held in mass culture for more than six
generations before release.

Production

of Cryptolaemus montrouzieri

Armitage (1919) first reported on rearing and release of
C. montrouzieri for control of mealybugs in citrus. (A
complete documentation of the historical development of
the use of C. montrouzieri is given by DeBach and Hagen
1964.) Beglyarov and Smetnik (1977) reported that C.
montrouzieri is mass-reared in the U.S.S.R. and “hun¬
dreds of thousands” are released annually in plantations
of citrus, grapes, tea, and other plants for control of
mealybug and scale insects.

C. montrouzieri has been reared on various hosts feeding
on several food sources. But rearing methodology (fig. 3),
regardless of host and plant-food source, is usually like
the system reported by Fisher (1963). The citrus
mealybug, Planococcus citri (Russo), is mass-reared on
potato sprouts grown in subdued light on soil held in
wooden trays. After the sprouts are about 4.7 cm tall,
mealybug crawlers are allowed to crawl onto freshly cut
leafy terminals of Pittosporum undulatum Vent, or
Schinus molle L., which are placed in the trays for about
6 hours. These terminals are then placed among the
potato sprouts in darkened rooms, and the crawlers move
onto the sprouts as the terminals begin to dry. After 8
days, C. montrouzieri adults are placed in the trays where
they lay eggs on the potato sprouts and trays for 12
days. Then these adults are collected at opened windows
screened with cotton muslin and released in the citrus or¬
chards. Meanwhile, burlap bands are attached to the
front of the racks holding the trays as a substrate for
pupating beetle larvae, and the holding room is again
darkened for 6 days, the window shutter is once more
raised, and emerging beetles are collected for release at
the cloth-covered window opening.

Since C. montrouzieri was imported into the United
States in 1892 (DeBach and Hagen 1964), there has been
no success in either locating or selecting a strain that can
withstand temperatures below 0° C (Beglyarov and Smet¬
nik 1977). We found no reports on how laboratory rearing
affects genetic processes, or maintenance of desirable
characteristics in laboratory-reared beetles.

Fisher (1963) discussed in detail how to select the most
suitable potato variety for rearing the citrus mealybug.

Figure 4.— System for producing E. formosa
(parasite) on whiteflies (host) feeding on tobac¬
co plants.

Apparently, it feeds on sprouts of some potato varieties
more readily than on others, certain varieties are more
readily available than others, and some potatoes bruise
more easily than others (so they have a shorter storage
time). Watering the potato sprouts and implanting
crawlers on them must be done carefully. Beglyarov and
Smetnik (1977) reported that an artificial diet had been
developed for rearing C. montrouzieri; this diet might
eliminate many problems associated with maintaining the
host and its food source. Fisher (1963) did not mention
any pathogens affecting the beetle colony; however, con¬
taminants did affect storage time of the potatoes. Many
rearing procedures could be mechanized; facilities for

210

maintenance of the proper environmental conditions
could be built; and the handling of materials could be
simplified.

Production of Encarsia formosa

Hussey and Bravenboer (1971) reported that E. formosa
is widely used in Europe, particularly England and the
Netherlands, for control of the greenhouse whitefly,
Trialeurodes vaporariorum (Westwood). There are two
basic techniques for host (whitefly) production; one uses
cucumber plants, Cucumus sativus L. (Morrison and King
1977) and the other uses tobacco plants, Nicotiana
tabacum L. (Glasshouse Crops Research Institute 1975,
Hussey and Scopes 1977). In the system using tobacco
(fig. 4), the tobacco plants are exposed for 8-24 hours to
whitefly adults, which lay about 100 eggs/6.5 cm2. Then
the plants are shaken by hand and fumigated with
dichlorvos3 to eliminate the adults; and the egg-infested
plants are held until the whitefly reaches its third stage
(“ceases to be flat disc and develops elevated sides”). At
that time, adult parasites are placed on the plants. Ten
days later the parasitized and unparasitized whitefly
pupae are brushed from the leaves into suitable con¬
tainers and held for emergence of the unparasitized
whiteflies. (They emerge before the parasites; so they can
be easily removed from the parasite culture.) The remain¬
ing parasites are used for release in commercial
glasshouses and for the reproductive colony. Other
methods are available for producing the parasite on
tomato plants while the whiteflies are produced on tobac¬
co (Glasshouse Crops Research Institute 1975).

Cucumber plants for producing hosts and parasites are
used mainly in the Netherlands. There, plants grown ver¬
tically on string are the hosts. At first, about 100 white¬
fly adults are implanted on the young plants. Then, about
2 weeks later, E. formosa is introduced as a mature pupa
at about 8 parasites to 10 whiteflies. After 2 more weeks,
harvest of the bottom leaves (containing only mature E.
formosa pupae and emerging adults) begins after inspec¬
tion shows that some parasites have emerged. Both pest
and parasite move up the plant to newly emerged and in¬
fested leaves, and this production unit can sustain itself
for several months.

None of these studies gave methods for measuring the
behavioral quality of the mass-produced parasites. Pro¬
duction is apparently closely monitored. And raising or
lowering the temperature can influence synchronization of
host and parasite development.

32,2-Dichlorovinyl dimethyl phosphate.

Production of Lixophaga diatraeae

Ridgway et al. (1977) report that L. diatraeae is reared
and periodically released in several Western Hemisphere
countries for biological control, mainly of the sugarcane
borer, Diatraea saccharalis (Fabricius). It can be reared
on many lepidopterous larval species (Bennett 1969), but
King et al. (1979) reported that a large-scale production
system has recently been developed on an unnatural host,
the greater wax moth, Galleria mellonella (Linnaeus). The
method discussed here maintains the parasite on its
natural host, the sugarcane borer.

In the system for producing L. diatraeae (fig. 5), the adult
flies are held in cages covered with gauze. The cages con¬
tain food and water. When they are 12-14 days old, the
flies are aspirated into a collecting jar containing 1%
NaOCl. After the NaOCl is rinsed from the flies, they are
blended at 8,500 r/min (revolutions per minute) for 9
seconds in a 0.7% Formalin (formaldehyde) solution so
the maggots can be extracted. Passing what is left
through a screen separates the maggots from the remain¬
ing fly particles. Then the maggots are rinsed, suspended
in a 0.15% agar-water solution, and placed in a gridded
petri plate where the total maggot number is determined.
Afterwards, more agar-water solution can be added to
produce the desired maggot density in a given solution.
For either the sugarcane borer or the greater wax moth,
the larvae are exposed to the parasite maggots during
the last stage. The agar-water solution containing the
maggots is metered into 30-ml cups containing sugarcane
borer larvae and diet. When L. diatraeae are reared on the
greater wax moth, larvae are placed in trays containing
the parasite maggots, which have been poured or air-
brushed. Then a screen is placed over the trays; after
about 1 hour, corncob grits are added to the trays to ab¬
sorb excess moisture. On either host, the maggots com¬
plete development in 6-9 days, and both hosts and
parasite puparia are harvested on the 11th day after
parasitization. Puparia are removed with forceps from
cups containing sugarcane borer larvae. Puparia formed
outside greater wax moth cocoons are brushed from the
trays; those retained in the host cocoon are collected by
flotation after the cocoon is dissolved in 1% NaOCl.
Puparia to be shipped are packed between layers of cot¬
ton in cartons and placed in styrofoam boxes containing
icepacks that will maintain low temperatures (18°-25° C)
in transit. The flies are released in sugarcane fields 4-6
days after emergence (prelarviposition period).

In the system discussed by King et al. (1979) for produc¬
ing L. diatraeae, production was routinely monitored by
measuring of adult survival; mating; sex ratio; maggot
production; and percentage of parasitization of host lar¬
vae, of puparia production, and of fly emergence. No
routine methods were developed for monitoring parasite

211

Figure 5.— System for producing L. diatraeae (parasite) on its
natural host, the sugarcane borer, or an unnatural host, the
greater wax moth.

212

Figure 6.— System for producing S. endius
(parasite) on house fly pupae (host).

behavior. But King et al. (1978) demonstrated strain dif¬
ferences between L. diatraeae populations, and Morrison
and King (1977) reported that L. diatraeae quality
(puparium size, fly emergence from puparia, and adult
survival) was affected by host larval diet. Host larval diet
has also been shown to affect parasite larviposition rate
(J. P. Roth and E. G. King, unpublished data).

Production of Spalangia endius

S. endius has been. very effective in suppressing house
flies, Musca domestica Linnaeus, near poultry farms
(Legner and Brydon 1966, Legner and Dietrick 1974,
Weidhaas and Morgan 1977). And rearing facilities
(Morgan and Patterson 1977) and methods (Morgan et al.
1978) have been well developed. Partly because of these
developments, S. endius and similar microhymenopterous

parasites are being produced commercially and sold to
farmers (DeBach 1974).

In the system reported by Morgan et al. (1978) for mass
producing S. endius on house fly pupae (fig. 6), 2-day-old
house fly pupae (200,000) are transferred to Plexiglas
parasitization cages containing about 60,000 S. endius
adults. Since the parasite sex ratio is two females to one
male, 40,000 females are assumed to be available for
oviposition. The house fly pupae are put in the cages on
aluminum trays (6 trays/cage). The parasites disperse
among the pupae to lay eggs and also to feed on
hemolymph flowing from host oviposition wounds. After
the 1-day exposure, most of the wasps, which are
positively phototactic, are separated from the pupae by
being attracted to another part of the cage with a bright
light. The trays of host pupae are then emptied onto a
screen, and live parasites not attracted to the light are
sifted from the pupae. So most of the surviving female
parasites are retained in the cage. Adding 20,000
parasites to each cage each day compensates for the
average daily loss of 33% of the parasites and keeps the
number of females at 40,000 per cage. Also, when the
host pupae are removed, another lot of 200,000 pupae is
placed in the cage. The host pupae removed from the
parasite cage are divided into equal lots, placed in paper
containers, and held for 5 days at 27.8° C. During this
time, adult house flies emerge from unparasitized pupae
and can be eliminated. The remaining parasitized pupae
are then used for maintaining the parasite colony or for
release. Although production records are kept, Morgan et
al. (1978) did not report any routines for controlling
behavioral quality.

Production

of Phytoseiulus persimilis

P. persimilis is now being used in many countries as a
predator of tetranychid species in glasshouses (Markkula
et al. 1972, Beglyarov and Smetnik 1977, Hussey and
Scopes 1977, Ridgway et al. 1977). And Mori (1974)
shows its potential for controlling tetranychids attacking
field crops. Because P. persimilis cannot yet be produced
at a cost competitive with acaricides, it is not being used
more widely.

Morrison and King (1977) and Glasshouse Crops Re¬
search Institute (1975) have reported that P. persimilis is
generally produced in three phases— production of plant
material (bean); production of the host, Tetranychus ur-
ticae Koch, on the bean plants; and production of P. per¬
similis on T. urticae. The bean plant Phaseolus vulgaris
L. is used in the Netherlands and England for rearing T.
urticae. In England, the T. urticae population is first
started on another kind of bean, Vida faba L., 1 week
after planting. Three weeks later, the now heavily in-

213

fested V. faba plants are laid on pots of mature P.
vulgaris that will be a food source and substrate after im¬
plantation with P. persimilis. In the Netherlands, T. ur-
ticae are implanted after appearance of the first true
leaves (fig. 7). When the second true leaves appear, P.
persimilis is introduced. Once P. persimilis has multiplied
and has eliminated the T. urticae, the leaves can be
harvested and stored at low temperatures (7.3°-12.9° C)
or delivered to release spots. Although production records
are kept, there are apparently no routines for monitoring
the quality of the predator or parasite produced.

Production

of Trichogramma species

Species of Trichogramma are mass-produced and used to
control caterpillar pests in the U.S.S.R. (Shcheptil’nikova
et al. 1974), China (National Academy of Sciences 1979),
and Mexico (Gomez 1975), and also, but not as much, in
Western Europe and the United States (Starler and
Ridgway 1977). The parasites are typically mass-
produced on unnatural hosts and not the target or
natural host.

The Angoumois grain moth, Sitotroga cerealella (Olivier),
is used worldwide, except China, as an insect host for
Trichogramma because it can be easily and inexpensively
mass-produced. In China, most large-scale Trichogramma
production is on eggs from a silkworm, Antheraea pemiyi
Guerin-Meneville, reared outside on oak trees, Quercus
spp., grown in field plots. Other insect hosts include the
Mediterranean flour moth, Anagasta kuehniella (Zeller), in
Western Europe and another silkworm, Sarnia
(=Philosamia) cynthia ricini (Boisduval), and a grain
moth, Corcyra cephalonica, in China. Interestingly, as a
byproduct of parasite production, the silkworm cocoons
can be used for sericulture. Nevertheless, not all impor¬
tant Trichogramma spp. can be reared on silkworm
eggs —7! ostrinaeae for example— so C. cephalonica is also
needed in China.

In China, the silkworm reared on oak foliage is harvested
from field plots after the larvae have formed cocoons in
late summer. The cocoons are stored at low temperatures
during the winter and until needed the next year. Co¬
coons containing female pupae are mechanically separa¬
ted from those containing njale pupae by size and weight
(the female is larger and heavier). The female moths are
collected as they emerge and passed through a grinder, a
process that extracts the unfertile eggs. Screening and
decanting the water used in the grinding process sep¬
arates the eggs from other body parts; the eggs are then
centrifuged and dried. Since the eggs are not fertile, they
can be stored at low temperatures for several weeks after
extraction without embryonation occurring. For
parasitization, the eggs are glued to cards or sheets of

Figure 7.— System used in the Netherlands for
producing P. persimilis (predator) on T. urticae
(prey) feeding on bean plants.

paper and exposed to the parasites for 2 or more hours.
Typically, unparasitized eggs are placed near a light
source 1 or more meters from the reproductive stock of
parasites. The parasites are positively phototactic; so
they will find the eggs after flying to the light source.
After enough parasites are on the host eggs (about one
parasite per egg) they are moved to a darkened area to
complete parasitization. The parasitized eggs can be
stored at low temperatures (5° C) until time for field
release.

214

I o* ^

215

The basic system (fig. 8) used for producing Trichogram¬
ma in the United States (Morrison et al. 1978) and in the
U.S.S.R. (Beglyarov and Smetnik 1977) requires exposing
Angoumois grain moth eggs to Tricho gramma adults in a
closely confined area. The host eggs are attached to
plastic sheets with a fine mist of water, or they are glued
to paper. After the desired exposure time has elapsed, the
now-parasitized host eggs are removed from the parasite
chamber and held for parasite development. Parasitized
host eggs can be broadcast on fields if they are attached
to plates or point-released if they are glued to paper.

Specifically, after host eggs are stuck to plastic or glass
sheets, the sheets are exposed to parasites in a glass¬
sided, illuminated oviposition cage for 24 hours (Morrison
et al. 1978). The sheets containing the now-parasitized
eggs are moved to the back half of the oviposition cage,
which does not have glass sides and is dark. At the same
time, freshly prepared eggs are introduced. After 20
hours, the parasitized egg sheets in the back are removed
through a door at the dark end of the oviposition cage.
And, while the door is open, specific amounts of parasite
pupae due to emerge within 24 hours are placed in the
oviposition cage, and similar material placed in the cage 4
days earlier is removed. So a consistent, specific number
of ovipositing adults is in the cage. Since Trichogramma
is positively phototactic, those adults remaining on the
host eggs at the dark end of the unit and those emerging
from pupae move to the light at the glass-sided end of
the unit. Light intensity and diffusion must be adjusted
so the parasite will be attracted to the lit end of the cage
but not so strongly attracted that it remains on the glass
exclusively. Also, glass sides of the cage are alternately
lit and darkened so that phototactic parasites will move
back and forth across the host eggs. This movement
results in a consistent parasitization rate.

The parasitized eggs on the sheets removed from the
cage are brushed onto organdie-covered frames for fur¬
ther development at 26.8° C. After enough oviposition
stock is reserved, the remaining parasitized eggs are
placed in cold storage (R. K. Morrison, unpublished data)
or they are cold-programed for aerial release (Bouse et al.
1978). The temperature-programing system used requires
holding the eggs at 26.7° C for 6 days and transferring
them on the seventh day to a chamber where they are
held at 16.7° C for 6 more days. Over 75% of the
Trichogramma will emerge from these eggs within 4
hours after application in the field.

Regardless of the system, quality-control procedures are
typically production oriented and consist of keeping
records on production, parasitization rate, emergence, and
sex ratio. Each system does stress the need for replacing
the colony every year with field-collected material to
avoid release of a parasite that may have reduced effi¬

ciency in host searching and parasitization. The Chinese
system is unique because it requires the parasites to fly
several feet in search of host eggs, so it continually
eliminates weak, inactive individuals. Also, in China,
laboratory colonies of Trichogramma spp. are typically
replaced with field-collected parasites after about 15
generations because of deterioration (J. Z. Bao, personal
communication). In the United States, selection of a
parasite strain that will attack the target host is ac¬
complished by annual colony establishment with para¬
sites collected only from the target host and affected
crop. Also, large numbers ( > 2,000) of the parasites are
collected to insure a broad genetic base.

Other Considerations
in Production of Entomophages

Host production

In the United States, production of entomophagous ar¬
thropods to augment natural supplies is confined to a few
private firms, cooperatives, and government agencies.

But production and use of predators and parasites for
control of arthropods is much more widespread
worldwide. Nevertheless, without exception these en¬
tomophages are being mass-produced on live hosts or
host products. The necessity of rearing the host and
sometimes the host’s food plant often more than doubles
the cost and complexity of rearing the parasite or
predator, restricting wider-scale usage of this technique
in biological control.

In China, production of Trichogramma on oak silkworms
mass-produced in the field on oak trees does circumvent
many of the problems that come with producing host
material in the laboratory insectary. And the oak
silkworm cocoon can still be used for silk producton.
Others have extended this technique by producing the en¬
tomophages on hosts reared in the field. For example,
Halfhill and Featherston (1973) reported a study where
the parasite Aphidius smithi Sharma and Subba Rao was
reared in field cages containing the host pea aphid, Acyr-
thosiphon pisum (Harris), and allowed to escape into sur¬
rounding alfalfa, Medicago sativa L., fields through
temperature-controlled vents in the cage roofs. And
Stevens et al. (1975) reported that the Mexican bean bee¬
tle, Epilachna varivestis Mulsant was controlled in soy¬
beans, Glycine Max (L.), by release of the eulophid
Pediobius foveolatus (Crawford) in nurse crops (snap
beans, Phaseololus rulgaris L.) where they multiplied on
Mexican bean beetle larvae and spread into adjacent soy¬
bean fields. Another technique for rearing entomophages
in the field is the application of supplementary foods that
induce oogenesis (Hagen and Hale 1974).

216

Artificial diets
and in vitro rearing

Excellent artificial diets are available for rearing many
phytophagous arthropods but are generally lacking for
entomophagous arthropods (Singh 1977; see House 1977
for the most recent review of nutrition of entomophages).
Development of suitable artificial diets could enable mass
production of entomophagous arthropods for augmen¬
tative purposes at a cost competitive with other methods
of insect pest control, and could insure production of an
organism of consistent quality.

For predators, the most progress has been made in
developing artificial diets for rearing C. camea. Signifi¬
cant advances include development of an artificial diet for
rearing larvae (Hagen and Tassan 1965; Vanderzant 1969,
1973), encapsulation of the larval diet (Martin et al. 1978),
and development of a nutritious and practical adult diet
(Hagen and Tassan 1970). Some progress has been made
in artificial diets for rearing some of the coccinellids— for
example, Coleomegilla maculata DeGeer (Atallah and
Newson 1966).

The state of our knowledge on nutrition of parasitic
Hymenoptera and Diptera is poor; however, some species
have been reared in vitro. Bronskill and House (1957)
reported limited success at rearing the ichneumonid endo-
parasite Pimpla turionella (Linnaeus) on pork liver. And
Coppel et al. (1959) reported that pork liver was an ex¬
cellent diet for rearing the sarcophagid Agria housei
Shewell [= Agria affinis (Fallen) or Pseudosarcophage af-
finis (Fallen)]. House (1954) developed a diet for rearing
A. housei and continually refined it (House 1972).
Bouletreau (1972) was successful at rearing the pupal en-
doparasite Pteromalus puparum Linnaeus in host
hemolymph; and Hoffman and Ignoffo (1974) were able to
rear this parasite on media containing yeast hydrolysate
and bovine serum. Recently, Yazgan and House (1970)
described a successful, chemically defined artificial diet
for rearing the ichneumonid pupal endoparasite Itoplectis
conquisitor (Say). This diet was greatly improved by
Yazgan (1972), so parasite mortality decreased; and
House (1978) developed a method for encapsulating it.

The hymenopterous ectoparasite Exeristes roborator
(Fabricius) was also reared successfully on a chemically
defined diet by Thompson (1975), and he continued these
studies (Thompson 1977) to expand our knowledge on
nutrition of hymenopterous larval parasites. Successes in
rearing of the egg parasite Trichogramma spp. (Hoffman
et al. 1975, Guan et al. 1978, Liu et al. 1979) show that a
suitable artificial diet for rearing this insect is feasible.
Likewise, recent successes in rearing tachinids in vitro
(Grenier et al. 1975, 1978; Nettles et al. 1980) have ex¬
panded our knowledge on their nutrition. In fact, Net¬
tles et al. (1980) demonstrated that the parasitic maggot

of Eucelatoria bryani Sabrosky does not have to at¬
tach itself to the host tracheal system to complete
development.

Use of unnatural hosts

Many large-scale rearing programs have used unnatural
hosts for production of entomophagous arthropods. Un¬
natural hosts are typically used in place of natural hosts
because of costs, convenience, and ease of handling. In
fact, four of the programs discussed above used un¬
natural hosts for mass production— A melinus is reared
on oleander scale fed squash, C. camea is reared on
Angoumois grain moth eggs from wheat-fed moths, L.
distraeae is reared on greater wax moth larvae fed a
cereal diet, and Trichogramma spp. are reared on
Angoumois grain eggs from wheat-fed moths.

There has been some concern that entomophagous ar¬
thropods reared on unnatural hosts may change their
host preference because of preimaginal conditioning and
would therefore be less effective when released for control
of the natural host (National Academy of Sciences 1969).
It has been shown that rearing a parasite on an un¬
natural host increases a parasite’s acceptance of that
host. But Thorpe and Jones (1937) found no evidence that
a parasite prefers the unnatural host to the natural after
only a few generations. In fact, more recent research (Ar¬
thur 1965, Bryan et al. 1968, Legner and Thompson 1977)
has failed to demonstrate this phenomenon. Selection for
genotypes that respond more rapidly to unnatural host
and survive better on it may explain some reported cases
of preimaginal conditioning. In any event, parasites
reared for a short time on an unnatural host will probably
respond as strongly to the natural host as those reared
on it.

Reduced vigor of entomophages because of exposure to
laboratory hosts that supply inadequate nutrition to the
attacker is probably the most critical problem in use of
unnatural hosts for rearing (Morrison and King 1977).

For example, Trichogramma spp. reared on unnatural
hosts have been shown to have reduced size, fecundity,
longevity, and general robustness (Lewis et al. 1976). And
host diet has also been shown to affect the suitability of
a host for parasite development. For example, Etienne
(1974) reported that L. diatraeae could not be continuous¬
ly reared on greater wax moth larvae fed beeswax and
pollen, unless the host diet was supplemented with
vitamin E or wheat germ. And vigor of L. diatraeae flies
was improved when a cereal diet fed to greater wax moth
larvae was supplemented with wheat germ and the pro¬
tein content increased (King et al. 1979). Reduced vigor
has also been demonstrated in parasites that have been
reared on the natural host when it is fed different foods
(Kajita 1973, Sato 1975, Altahtawy et al. 1976). So the

217

host’s nutritive value to the parasite can be significantly
modified by the food it eats. Finally, suitability cannot be
determined merely by screening hosts, but host nutrition
and other factors must also be considered; and com¬
promises between entomophage quality and quantity
may have to be made because of cost and the numbers
required.

Storage

Most predators and parasites used in augmentation pro¬
grams must be mass-produced as needed and released at
a specific time in relation to development of the pest
host. The ability to store the entomophage or host for
long periods would greatly reduce costs and waste caused
by necessary seasonal increases and decreases in produc¬
tion. Morrison and King (1977) reviewed techniques used
in storing various species of entomophages and found
that holding at low temperatures to reduce developmental
rates is typically the major component in almost all
reported storage techniques.

Quality control

Apparently, no one routinely measures traits or at¬
tributes such as genetic variation, diurnal rhythmicity,
flight propensity, and flight ability to insure that essen¬
tial behavioral characteristics are maintained. Most
measurements now done assess production. These
measurements include percentage of parasitization and of
adult emergence, size or weight, sex ratio, and amount of
egg production. These production measurements are, of
course, essential for a consistent, physically standard
product but do not relate directly to field performance.

Several researchers have divided quality into various
components and then attached relative values to them
based on the proposed use of the predator or parasite (see
Boiler 1972, Boiler and Chambers 1976, Hoy 1976, and
Huettel 1976). We have consolidated these into four com¬
ponents— adaptability, sexual activity, host selection, and
motility. Increasingly, rearing programs require monitor¬
ing of traits that manifest these components; but few
instances exist where these have been used to control
quality in entomophagous arthropods (Huettel 1976).

Occurrence of the genetic bottleneck that insects go
through during colonization is a well-established
phenomenon (Boiler 1972, Mackauer 1972). Production is
usually low during a new colony’s first few generations in
the laboratory, but it increases after five to seven genera¬
tions. Obviously, the colony is undergoing intense selec¬
tion pressure for individuals having traits that enable
them to best survive and reproduce under insectary con¬
ditions. But, because genetic material is often lost in the
selection process, the organism’s ability to survive and

reproduce in the field may be reduced. And genetic
deterioration of the colony often becomes apparent only
after it is irreversible, because behavioral traits are not
routinely monitored. Periodically replacing the laboratory
colony with field-collected material has partly solved this
problem, but this is a myopic and costly solution. Ex¬
cellent opportunities exist for implementing techniques to
monitor and maintain essential characteristics and even
select for desirable traits in mass-produced entomophages,
but the necessary researchers have to be directly as¬
signed these responsibilities.

Engineering

Engineering efforts such as development of flow charts
combined with time-and-motion studies often lead to im¬
proved production and reduced costs. The trained
engineer can often visualize existing production problems
and their solutions better than an entomologist. So, a tru¬
ly interdisciplinary approach involving engineers, en¬
tomologists, insect behaviorists, nutritionists, geneticists,
and chemists is essential to production of entomophagous
arthropods.

Acknowledgments

Information reported in this paper was developed in
cooperation with the Mississippi Agricultural and For¬
estry Experiment Station and the Texas Agricultural Ex¬
periment Station. We acknowledge the assistance of J. L.
Moore, formerly a U.S. Agricultural Research Service
computer specialist, and T. C. Lockley, biological techni¬
cian, in preparation of flow charts.

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pests of farm crops. Kolos Publishing House,
Moscow. [In Russian.]

Singh, P.

1977. Artificial diets for insects, mites, and spiders.
594 pp. Plenum Press, New York.

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York.

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4: 947-952.

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1975. Defined meridic and holidic diets and aseptic
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ectoparasitoid Exeristes roborator (Fabricius).
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1937. Olfactory conditioning in a parasitic insect and

221

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1969. An artificial diet for larvae and adults of
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222

A Laboratory Method for Mass
Rearing the Eastern Spruce
Budworm, Choristoneura fumiferana

By D. G. Grisdale1

Introduction

The eastern spruce budworm, Choristoneura fumiferana
(Clemens), is the most widely distributed forest insect
pest in North America. Its range includes the Eastern
States from Virginia to Minnesota and all of the forested
regions of Canada from Newfoundland to Alberta, north¬
eastern British Columbia, the southern part of the Yukon
Territory, and the southern half of the MacKenzie River
basin of the Northwest Territories (Prebble 1975). It at¬
tacks balsam fir, Abies balsamea (L.) Mill.; alpine fir, A.
lasiocarpa (Hook.) Nutt.; white spruce, Picea glauca
(Moench) Voss; red spruce, P. rubens Sarg.; and black
spruce, P. mariana (Mill.) B.S.P. At the start of eastern
spruce budworm ’s life cycle in July and early August,
moths deposit eggs in masses on host trees (Prebble
1975). The larvae hatch in about 10 days and spin silken
hibernation shelters in crevices of bark, under bud scales
or lichens, and in the cups of old staminate flowers. First
larval molt occurs in late August in the hibernation
shelter; the second-instar larvae remain there without
feeding until the next spring. Overwintering larvae
emerge in late April or early May and bore into old tree
needles or into the unopened buds; or they feed on early
opening staminate flowers when these are available. After
a week or more, larvae move to opening vegetative buds
and feed on the flaring needles under a protective silken
shelter. Full-grown larvae pupate in the feeding webs in
late June or early July. The moths emerge from the pupal
cases about 2 weeks later, completing the annual one-
generation cycle.

In the 1920’s and 1930’s, aerial insecticide dusting was
used to control spruce budworms in small areas. In 1944,
aerial spraying with chemical insecticides began, and
such operations have been conducted in Canada every
year since then (Prebble 1975). In that time, intensive
research efforts have been made to develop more effective
methods of managing spruce budworm populations.
Before 1965, investigations were hampered by an inabili¬
ty to rear enough larvae for laboratory experimentation
and propagation of pathogens such as viruses and
microsporidia.

‘Research technician. Forest Pest Management Institute, Cana¬
dian Forestry Service, Sault Ste. Marie, Ontario.

Stehr (1954) described a method for rearing spruce bud-
worms throughout the year on shoots of balsam fir that
had been preserved by freezing. But not until a synthetic
diet was developed (McMorran 1965) was the rearing pro¬
gram able to expand. As the demand for budworm larvae
increased, methods were developed for mass producing
them (Grisdale 1970, 1972, 1973). This paper describes
further modifications in rearing methods. Presently, at
the Canadian Forestry Service’s Forest Pest Management
Institute in Sault Ste. Marie, Ontario, we are able to pro¬
duce about 250,000 spruce budworm larvae weekly. We
also mass-rear western spruce budworm, Choristoneura
occidentalis Freeman, by the same method.

Colony Maintenance

Artificial diet

The artificial diet that we use (table 1) is like McMorran ’s
(1965), though we substitute granulated agar for nutrient
agar, use a veterinary grade of Aureomycin (chlortetra-
cycline), and increase the amount of wheat germ. No For-

Table 1. — Composition of the spruce budworm diet and amounts
of ingredients for about 3,600 ml of diet (mixed in 1-gallon blender)

Ingredient Quantity

Granulated agar . g . . 60

Water (distilled), in blender . ml.. 792

Water (distilled), to dissolve agar . ml . . 2,232

Casein, vitamin free . g . . 126

4 M Potassium hydroxide . ml.. 18

Alphacel . g. 18

Salt mixture, Wesson . g . 36

Sucrose . g 126

Wheat germ . g 160

Choline chloride . g 3.6

Vitamin solution1 . ml . . 36

Ascorbic acid . g 14.4

Formalin (37% formaldehyde) . ml 1.8

Methylparaben . g 5.4

Aureomycin powder (chlortetracycline
hydrochloride, 55 mg/g) . g 20

‘100 ml contains 100.0 mg niacin, 100.0 mg calcium pantothenate,
50.0 mg riboflavin, 25.0 mg thiamine hydrochloride, 25.0 mg
pyridoxin hydrochloride, 25.0 mg folic acid, 2.0 mg biotin, and
0.2 mg vitamin B-12.

223

Figure 1.— Electric kettle (left) and vertical
cutter mixer (right) used in preparing casein
and wheat germ diet for rearing the eastern
spruce budworm.

Figure 2.— Disposable plastic cups and trays
used as rearing units for the eastern spruce
budworm.

malin (formaldehyde) is added to diet used in the multi¬
plication of virus because it may suppress the progress of
virus infection in the host (Vail et al. 1968). The diet is
prepared in 30-liter batches. An electrically heated steam-
jacketed kettle (capacity 75 liters) is used to liquify the
agar. The kettle (fig. 1) is fitted with a bridge-mounted
anchor-shaped mixer with nylon scrapers driven by a
1 -horsepower, variable-speed motor. Water is placed in
the kettle. With the mixer operating at low speed, the
agar is added and the heaters turned on.

While the agar is dissolving, water is added to a 40-quart
vertical cutter-mixer, a Hobart VCM-40 with two
speeds— 1,750 r/min (revolutions per minute) and 3,500
r/min— and equipped with standard narrow knives and a
hand-operated mixing baffle (fig. 1). The VCM is turned
on at low speed and the remaining ingredients added in
the order given in table 1 . A large funnel, custom made
to fit the largest opening in the inspection cover, is used
for adding the ingredients. The funnel allows uninter¬
rupted operation of the VCM and reduces splashing. The
VCM is left on for about 1 minute after the last ingre¬
dient has been added, and the mixing action is sup¬
plemented by the hand-operated mixing baffle. During
this primary mixing, some of the ingredients may stick to
the underside of the cover; so the VCM is stopped, and
this material is scraped off to insure that all ingredients

224

are in the diet and well mixed. The VCM is again
operated at low speed and the liquified agar added
through the funnel; the inspection cover is closed, and the
VCM speed is increased to 3,500 r/min for about 3
minutes. The diet is now ready to pour.

About 15 ml of diet is hand-poured into 3A-oz semitrans¬
parent ribbed plastic cups (Portion Packaging, Rexdale,
Ontario). This container was selected because each
second-instar larva can establish its own feeding site be¬
tween the ribs and be relatively undisturbed by others in
the cups until the fourth instar. Another advantage of
these cups is that larval development can be seen directly
through the cup, so the lid need not be removed. Dispos¬
able pressboard cafeteria trays (46 cm by 36 cm) are
used to hold the cups. About 90 cups can be set up in each
tray, which also serves as a unit for rearing the insects
(fig. 2). Trays are light and inexpensive and withstand
autoclaving once or twice before being discarded. After
the diet has cooled, it is sprayed with an antifungal solu¬
tion that is a modification of the one described by Chawla
et al. (1967). It contains 1.5 g sorbic acid and 0.6 g
methylparaben (methyl p-hydroxybenzoate) in 100 ml of
95% ethyl alcohol. The alcohol solvent evaporates rapid¬
ly, leaving a thin film of antifungal agents on all exposed
surfaces inside the cup. Trays of prepared diet are placed
in plastic bags (six trays per bag) that are sealed and
refrigerated at 0°-l° C until ready for use.

Larvae

Second-instar larvae in hibernacula and with diapause re¬
quirements satisfied are removed from cold storage and
held at 20°±1° C for 24 hours. Then they are placed in a
lighted rearing cabinet at 22°±1° C and 70% relative
humidity. After 1 additional day, or before larvae emerge
from hibernacula, a cheesecloth-Parafilm roll is cut into
strips on scored lines. These strips are cut into appro¬
priately sized patches (with 25-40 larvae each) and placed
in diet-filled cups. These are then capped with lids made
of unwaxed paper. To reduce the incidence of fungus in¬
fection, the cups are inverted and tapped to insure that
the gauze patch (fig. 3) falls onto the lid and away from
the surface of the diet.

When a specific number of larvae per cup is required, a
large gauze patch is placed in a glass dish (19 cm in
diameter) sealed with Parafilm. After emergence, larvae
are transferred individually to the diet by a camel’s-hair
brush. The dish must be moistened often; if emerging lar¬
vae are held overnight, the dish should be kept in a cool
(19°±1° C) darkened area to reduce larval activity and
mortality. Emergence time of second-instar larvae may be
shortened by at least 24 hours if patches are held in con¬
stant light at 24°± 1 0 C and kept moist and the container is
opened often for airing.

Colonized cups are exposed to 18 hours of light,
temperature of 24°±1° C, and relative humidity of 60%
until the larvae reach the fourth or fifth instar. This is
the point in the Institute’s program when most spruce
budworms are distributed for testing and virus
multiplication. Larvae to be used for the maintenance of
rearing stock or for physiological studies are transferred
to cups (six or seven larvae each) containing 10 ml of
diet. If the cups are allowed to remain crowded, can¬
nibalism becomes excessive and pupal and adult sizes are
sharply reduced. Larvae are easily sexed; so, for special
requirements, they can be reared separately from this
time. (Paired testes of male larvae are readily visible in
third and later larval instars.) When larvae are in the
sixth instar and near pupation, trays of cups are inverted
so that lids are upright. Because most larvae pupate near
the lid, pupae are more easily harvested with the cups in
this position. Larvae spend about 4 days in each of the
second, third, and fourth instars, and 5 days in the fifth.
Male larvae require about 8 days in the sixth instar
before pupating; females need 2 or 3 more days.

Diapause-free strains of spruce budworm may be
developed as a standard laboratory culture. A few non¬
diapause larvae (commonly called wandering seconds) are
often observed in families produced from field-collected
material. Shortly after molting into the second-instar,
these larvae leave the hibernacula. Selection over six
generations may yield a strain that is nearly free from
diapause. (See Harvey 1957 for a description of diapause-
free development and of rearing conditions necessary to
produce a reliable laboratory strain.) But, because of con¬
stantly fluctuating research demands, we have found that
removing second-instar larvae from cold storage when
and if required serves our needs more efficiently than
developing diapause-free strains.

Pupae

After pupation, pupae are sexed by the method of Jen¬
nings and Houseweart (1978). Because we sex so many
pupae, we find that the best method for general rearing
purposes is to count the abdominal segments visible ven-
trally and posterially to the wing pads (females have
three segments; males have four). Experienced personnel
observing these characteristics and the general shape of
the pupae rarely err. For studies requiring segregated
virgin moths, sexing by the location and shape of the
genital opening may be used. Sexed pupae are placed in
well- ventilated plastic crisper trays (27 by 20 by 10 cm)
for adult emergence (fig. 4). To facilitate eclosion, the bot¬
tom of the tray is carefully lined with paper toweling
fastened with tape to prevent the liner from shifting
when adults are removed. To synchronize adult emer¬
gence, male pupae are held at a temperature 2° C lower
than female pupae. The pupal stage lasts about 8 days.

225

Figure 4.— Sexed eastern spruce budworm
pupae and emerged adults in ventilated plastic
crisper trays.

1

Figure 6.— Eastern spruce budworm eggs in
black-headed stage attached to balsam fir
foliage distributed evenly near gauze patch on
aluminum pan bottom.

Figure 5.— Eastern spruce budworm mating
and oviposition cage. Note plastic shield on
cage door.

Mating and oviposition

A screened cage (35 by 35 by 25 cm) is used for mating.
The cage (fig. 5) has a sliding glass door; a paper towel
covers the bottom. Four or five small balsam fir branches
about 35 cm long are placed on the bottom of the cage as
oviposition sites. This amount of foliage seems to reduce
the disturbance of females when they are laying eggs and
results in larger and more uniform egg masses. Normally,
200 pairs of adults are transferred to a cage (after a short

time in a refrigerator) in a 12- by 15-mm shell vial. A
movable plastic shield is placed behind the sliding cage
door and is most helpful in preventing adults from escap¬
ing when they are first placed in the cage and later when
branches are removed. Up to 300 pairs of adults may be
introduced into cages with good results. Tests showed
that 150 pairs produced 28,653 eggs, or an average of
191.0 eggs per female; 200 pairs produced 41,293 eggs, or
206.5 eggs per female; 250 pairs produced 52,507 eggs, or
210.0 eggs per female; and 300 pairs produced 61,025
eggs, or 203.4 eggs per female (C. B. Budar and D. G.
Grisdale, unpublished data). Adults are held at 20°±1° C
and relative humidity of 80%-90% with 12 hours of light.
Cages are sprayed generously with distilled water two or
three times daily. Egg laying is largely completed about
a week after mating. Adults are killed in an oven and
discarded on the eighth or ninth day after introduction
into cages.

Eggs

Branches with egg clusters attached on needles are
removed from the mating cages every second day. To pre¬
vent excessive adult movement, cages are sprayed with
water just before branches are changed. Needles bearing
egg clusters are removed from branches by surgical
scissors or forceps. To reduce the amount of foliage
material and for a better estimate of how many eggs are
placed in hatching pans, the needles are cut just below
the egg cluster. Eggs are placed on uncovered disposable
cardboard trays and exposed to 18 hours illumination,
temperature of 24°± 1° C, and relative humidity of
55%-60% until they reach the black-headed stage and are
about to hatch. Holding egg masses of the same age until
they reach this stage insures uniform egg hatching and

226

Figure 7.— Sealed pan with attached cheese¬
cloth on Parafilm for hatching and establish¬
ment by eastern spruce budworm second-instar
larvae.

larval spinning and shortens the time larvae spend in the
hatching and spinning unit. Also, the water content of
the needles is reduced and practically eliminates any
problems with fungus. Larvae hatch about 8 days after
oviposition.

Hatching and spinning

An aluminum roasting pan (Supreme Aluminum of
Canada, No. 4910) is used as the hatching and spinning
unit. This pan is particularly well suited because of its
convenient size (45 by 30 by 6.5 cm). Its rounded edges
permit a perfect seal with Parafilm; it is easily cleaned
and sterilized; and unlike glass or plastic, it is extremely
durable. (Smaller pans of the same type are used when
there are not enough eggs for efficient use of the larger
pan.) A double layer of cheesecloth (30 by 15 cm) is at¬
tached to a slightly larger piece of Parafilm, which is
then pressed into the bottom of the pan. Egg masses in
the black-headed stage (enough to produce about 16,000
second-instar larvae) are distributed evenly over the bot¬
tom of the pan but away from the gauze (fig. 6). The pan
is then tightly sealed with Parafilm. On the underside of
the Parafilm is a patch (40 by 25 cm) of cheesecloth. The
Parafilm is attached to the pan with strips of masking
tape pulled downward and away from the rounded edge
to insure an escapeproof seal (fig. 7). Care should be
taken during the sealing operation to prevent the pan
from tilting so that eggs do not shift from their original
position.

Stocks of cheesecloth on Parafilm of various sizes are
prepared in advance. Cloth is readily attached to Parafilm
by rolling firmly with a cardboard roller (such as the
Parafilm core). The cheesecloth is next scored (to reduce

Figure 8.— An opened cheesecloth-Parafilm
roll, strips and patches of second-instar
eastern spruce budworm larvae, and the roller
used in the preparation of strips.

larval deaths when Parafilm is cut with scissors at the
time of patching) with a brass roller (fig. 8). The roller
was made on a lathe from a piece of brass stock; it is 14
cm long, and its rounded circumferential ridges are 1.4
cm apart and 1.5 cm high. The scoring is done on a sur¬
face covered with heavy paper to prevent perforation of
the Parafilm by the roller. When it is rolled with very
firm pressure, the roller’s ridges score the surface of the
cheesecloth and embed the fibers into the Parafilm. The
first-instar larvae cannot spin hibernacula in these scored
areas, so the spun-up larvae appear as regular strips on
the cheesecloth.

Hatching pans are held at a temperature of 22°±1° C
under continuous lighting. Once large numbers of first-
instar larvae are observed crawling on the upper gauze,
the pan may be turned over and rotated periodically dur¬
ing spinning. This rotation insures a uniform distribution
of larvae throughout both pieces of gauze. Silk laid down
by hatching larvae cements needles to the bottom of the
pan where they remain firmly attached even when the
pan is turned over. As soon as second-instar larvae are
observed in hibernacula, the sheets of Parafilm with at¬
tached gauze are removed, rolled loosely, sealed in¬
dividually in small plastic bags, and placed in a darkened
area at an incubator temperature of 19°±1° C. Larvae are
held at this temperature for about 3 weeks from the date
of hatching and are then ready for cold storage. This
prestorage treatment of second-instar larvae is a critical
part of the rearing technique because it results in

227

significantly higher storage survival (McMorran 1973).
(See Harvey 1957 for a description of the behavior pat¬
tern of the laboratory-reared spruce budworm from hatch
to building a hibemaculum where it molts and enters
diapause.)

Hibernation

Plastic bags of young larvae in hibernacula are placed in
appropriately sized brown paper bags to facilitate record¬
ing and shelf storage. Insects are placed in a coldroom, at
a temperature of 1° C, where they remain for 18-35
weeks. Moisture is not added to the plastic bags before
storage, because it may promote the growth of fungus.

Production Management

Trouble-free budworm rearing depends on several factors.
Probably the most important is having dedicated, compe¬
tent rearing personnel. Strict adherence to rearing tech¬
niques, sanitation practices, quarantine regulations, and
the use of rearing units with adequate environmental con¬
trol are also important factors. As anyone involved in in¬
sect rearing knows, hardly a day goes by without a minor
problem, and occasionally major ones occur. Problems
happen most typically with artificial diet, fungal infec¬
tion, and worker health and safety.

Artificial diet

Artificial diet of consistently good quality is an impor¬
tant factor influencing the quality of laboratory-reared
spruce budworms. At first, diet was prepared in an elec¬
trically heated kettle fitted with a bridge-mounted mixer
and dispensed into cups by an automatic filler (Grisdale
1973). Diet prepared in this manner was of good quality
and met our requirements, but comparative tests of lar¬
val development showed that larvae reared on diet
prepared in smaller, more easily mixed amounts in a
1 -gallon blender consistently developed at a faster rate.
And occasionally a batch of diet prepared in the kettle
produced budworms that developed slowly and irregular¬
ly; when many thousands of larvae were involved, produc¬
tion schedules were disrupted. Inadequate mixing and the
length of time diet remained in the kettle while being
dispensed were primary suspects. So the Hobard VCM
was put into operation in 1976. Since that time, all
prepared diet has consistently been of high quality except
in a few instances when the ingredients (wheat germ in
one case, salt mix in another) were of poor quality. But,
in both cases, we were able to trace the source of defec¬
tive ingredients quickly by examining records on diet
preparation.

Records are kept of the dates when new units of diet in¬
gredients are first opened, of the temperature of the liq¬

uified agar, of the diet temperature when it is dispensed,
and of other pertinent information on each lot of diet.
Diet ingredients are purchased from firms that have pro¬
vided us with quality products in the past. Most diet
ingredients are refrigerated as recommended by the sup¬
plier; wheat germ that will not be used within 3 weeks is
stored in a freezer. We have tried to lower cost of produc¬
ing diet during times of austerity, but we have found
that less expensive products are often of unreliable quali¬
ty and not suitable for budworm rearings. Of the several
insect species that can be reared on the same lot of diet
(Grisdale 1973), only spruce budworm is noticeably af¬
fected by poor diet.

Control of fungi

Fungal development on synthetic diets and infection of
second-instar larvae both before and after storage can
create serious rearing problems unless careful sanitation
practices and rearing techniques are closely followed.
Other infection sources are present in our laboratory
because we rear as many as 12 different insect species at
any one time in the same rearing facilities. Some
species— such as silkworm, Bombyx mori (Linnaeus);
hemlock looper, Lambdina fiscellaria fiscellaria (Guenee);
and the oleander hawk moth, Deilephila nerii
(Linnaeus)— are reared on foliage; if trays are not cleaned
frequently, fungus develops on frass and uneaten leaves.
Handling of all such fungus-infected material is carried
out in a biological containment hood. Other measures to
combat fungi are use of antifungal agents in the diet, sur¬
face spraying, and environmental controls. The unwaxed
cup lid significantly reduces fungus contamination. But
the inverted cups should not be incubated on a smooth
surface, which stops air transfer within the cup and so
permits the incidence of fungus infection to increase
greatly.

Since balsam fir foliage used as an oviposition site is con¬
sidered a principal source of infection, branches are
placed in plastic bags in a 1% solution of household
bleach (Javex, 6% available sodium hypochlorite),
agitated until the foliage is thoroughly wet, and then
refrigerated for use as required during 1 week. Other
steps for preventing fungal contamination are removing
young larvae from hatching pans shortly after hatch,
keeping prestorage treatment of second-instar larvae
brief, and storing larvae without adding moisture. Follow¬
ing these procedures should eliminate fungal infection
from budworm rearings.

Worker protection

Worker protection during the production of spruce bud¬
worm has received considerable attention in recent years,
particularly since transient allergic responses do occur

228

and can sometimes be incapacitating. At the Institute,
the problem is further complicated because we rear
several species and because we produce large numbers of
insects such as Orgyia spp. that have both wing scales
and urticating larval hairs. Workers were required to
wear protective face masks and gloves when handling
adults or obnoxious materials. Work was carried out
under a fumehood; but, because our building design did
not allow adequate venting, fumehood efficiency was
reduced. And the allergens continued to cause problems
despite these precautions. The hazard was severe enough
to justify the deletion of some species from rearing even
though considerable financial investment was involved.

In a further effort to reduce the problem with scales, we
installed a biological containment hood in the main rear¬
ing room. The hood, designed for worker protection only,
proved very effective in protecting staff from obnoxious
materials. Two more hoods were installed in other areas
of high contamination in the building, and the health
hazard has been reduced to a very low level.

Quality Control

We have not yet greatly emphasized inline testing for in¬
sect quality except for regular sampling of laboratory
adults for the microsporidian parasite Nosema
fumiferanae (Thomson). Also, efforts are made to obtain
disease-free field insects for yearly introduction into
laboratory rearing stock. The principal requirement at the
Institute has been for a vigorous larva free of microsporid¬
ian spores for general experimentation and propagation of
pathogens. Even with the microsporidian present, the use
of an antifungal agent, benomyl,2 allowed the production of
a standardized larva.

Rearing stocks that are free

of microsporidian infection

One of the most important factors limiting production of
healthy spruce budworm larvae is the microsporidian
parasite N. fumiferanae. In the past, as production of bud-
worm larvae increased, we experienced a recurring infec¬
tion of laboratory stocks by this pathogen. Wilson (1980)
describes N. fumiferanae as a debilitating agent, under
natural conditions, that affects host vigor, longevity, and
fecundity. Rearing larvae on synthetic diet in the
laboratory appears to mitigate the effects on host vigor
and longevity; but fecundity is reduced. And, unless
measures are taken to reduce levels of infection, larvae are
unsuitable for research purposes because the presence of
the parasite interferes with interpretation of experiments

2Methyl l-(butylcarbamoyl)-2-benzimidazolecarbamate as
Benlate, a wettable powder with 50% available benomyl.

involving other pathogens. Also, in virus propagation pro¬
grams, yields are greatly reduced.

Microsporidia are widespread in nature, affecting all larval
instars, pupae, and adults of the spruce budworm (Nielson
1963). Wilson (1973) found that there is a general buildup
in the N. fumiferanae infection as the budworm infestation
ages over 2 or more years. It had been our practice to col¬
lect field pupae each year from areas of new budworm in¬
festations where the incidence of parasitism and disease
was generally low (Grisdale 1970). Mating and handling of
all field stock was done individually. Then both male and
female adults from successful matings were examined
microscopically for spores. Progeny of moths free of
disease were kept for laboratory rearing stock; infected
progeny were discarded or used in virus-multiplication
programs. The field stocks considered free of disease were
crossed with existing laboratory stock to increase vigor
and maintain genetic variability. Because we handled
so many insects, we made no further attempt to diagnose
adults from multiple matings. In spite of our precautions,
we were never able to completely eliminate micro¬
sporidia from our rearings, probably because spores
in microscopically inspected adults may be overlooked
even after two generations of individual rearings (G. G.
Wilson, personal communication). A few missed spores can
quickly build up, particularly under mass-rearing condi¬
tions. In the laboratory, spores in frass and regurgitate are
infectious to healthy larvae. And Wilson (1972) found that
infected female but not male adults readily transmit N.
fumiferanae to offspring.

So, in 1978 we began a determined and successful effort to
eliminate microsporidia from our rearings. Field-collected
insects from several Ontario infestations were reared in a
room separated physically from the main rearing room.

The procedures we used for individual mating, spinning,
and storage were those described by McMorran (1965).
Progeny of adults diagnosed as free of infection after two
generations of controlled rearings were introduced to the
general rearing operation. Small, representative samples
from each geographical location were held for one more
generation of controlled rearing. We began sampling of
adults from all multiple matings. Cage number and origin
of stock are recorded; progeny are assigned this identifica¬
tion; and, after diagnosis, the presence or absence of
microsporidian spores is recorded. Because as many as
20,000 adults may be processed weekly, only a represen¬
tative sample of 50 adults per cage can be conveniently
examined.

Rearing stocks

with microsporidian infection

Each year, several million larvae of acceptable quality and
enough overwintering larvae to implement these rearings

229

have been produced from stocks infected with N.
fumiferanae. Benomyl has been used routinely in this
laboratory since 1975 to reduce the incidence of
microsporidian infection in larval rearings. At 100 p/m
(parts per million) incorporated into the synthetic diet at
time of preparation, benomyl effectively limits microsporid¬
ian multiplication during the period budworm larvae are
exposed to it. Larvae so treated are satisfactory for most
experimentation and for virus-multiplication programs.
And, because benomyl is such an effective fungicide, diet
in cups rarely becomes contaminated. Of course, larvae
reared on diet containing benomyl are not suitable for
studies involving microsporidian parasites or fungi. Larvae
reared on this treated diet have reduced mating success, so
they should not be used for stock maintenance.

Also Harvey and Gaudet (1977) discussing the effects of
benomyl in varying concentrations, showed that it effec¬
tively limits microsporidian infection in spruce budworm
larval stages; but its effect does not carry over into the
adult stage. Microsporidia multiply vigorously at the end
of the sixth instar and during the pupal stage. Benomyl in
concentrations of 75 p/m and above reduced budworm
growth and fertility. The most notable effect was the
reduction in fertile matings and in percentage of eclosion
from eggs. Males were more sensitive than females to
benomyl.

With these findings in mind, we rear infected budworms to
be used for stock maintenance on benomyl diet until they
reach the late stages of the fifth instar. At that time, the
number of larvae per cup is reduced to six, and they are
placed on diet containing no benomyl. Feeding on the
benomyl-free diet throughout the last instar diminishes
some of benomyl’s undesirable effects. And, though
microsporidian infection is still present, egg production
and egg hatch appear near normal. Handling of insects
during research, which is sometimes done in less than ideal
conditions, may result in fungus contamination. So we are
often asked to rear even disease-free insects on the
benomyl diet.

Field tests of adults

reared in the laboratory

Field trials using laboratory adults have been few. And we
have not been asked to add to this sparse information on
the differences between laboratory and wild insects.

Testing to date has shown little apparent difference be¬
tween laboratory and wild females in their ability to at¬
tract wild males under field conditions (C. J. Sanders,
personal communication). But Ennis and Charlebois
(1979) reported that field tests showed that some inbred
laboratory strains do not perform well in the field.
Laboratory-reared males from our randomly outbred stock,
however, responded like field males to pheromone

traps— released males were recaptured after 3 days even
when field populations were very high. And, though
recovery in pheromone traps of released males from an
orange-eyed mutant strain (Ennis 1978) that had been in¬
bred for 6 years was very poor, it was near normal for
orange-eyed males previously crossed with field stock to
introduce genetic background from wild insects (Ennis and
Charlebois 1979).

Acknowledgments

I thank Barry Hurtubise who prepared the photographs.

References

Chawla, S.; Howell, J.; and Harwood, R.

1967. Surface treatment to control fungi on wheat
germ diets. J. Econ. Entomol. 60: 307-308.

Ennis, T. J.

1978. An orange-eye mutant of the spruce budworm,
Choristoneura fumiferana (Lepidoptera-
Tortricidae). Can. J. Genet. Cytol. 20: 427-429.

Ennis, T. J., and Charlebois, N.

1979. A release-capture experiment with normal and
irradiated spruce budworm males. Can. For.
Serv. Bi-Monthly Res. Notes 35: 9-10.

Grisdale, D.

1970. An improved laboratory method for rearing
large numbers of spruce budworm,
Choristoneura fumiferana (Lepidoptera-
Tortricidae). Can. Entomol. 102: 1111-1117.

1972. An improved method for producing large
numbers of second-instar spruce budworm lar¬
vae, Choristoneura fumiferana (Lepidoptera-
Tortricidae). Can. Entomol. 104: 1955-1957.

1973. Large volume preparation and processing of a
synthetic diet for insect rearing. Can. Entomol.
105: 1553-1557.

Harvey, G. T.

1957. The occurrence and nature of diapause-free
development in the spruce budworm,
Choristoneura fumiferana (Clem.), (Lepidoptera-
Tortricidae). Can. J. Zool. 35: 549-572.

Harvey, G. T., and Gaudet, P. M.

1977. The effects of benomyl on the incidence of
microsporidia and the developmental perfor¬
mance of eastern spruce budworm (Lepidoptera-
Tortricidae). Can. Entomol. 109: 987-993.

Jennings, D. T., and Houseweart, M. W.

1978. Sexing spruce budworm pupae. U.S. For. Serv.
Res. Note NE-255, 2 pp.

McMorran, A.

1965. A synthetic diet for spruce budworm,

Choristoneura fumiferana (Clem.), (Lepidoptera-
Tortricidae). Can. Entomol. 97: 58-62.

230

1973. Effects of prestorage treatment on survival of
diapausing larvae of the spruce budworm,
Choristoneura fumiferana (Clem.), (Lepidoptera-
Tortricidae). Can. Entomol. 105: 1005-1009.

Neilson, M. M.

1963. The dynamics of epidemic spruce budworm

populations. R. F. Morris (ed.). Mem. Entomol.
Soc. Can., pp. 272-287.

Prebble, M. L.

1975. Aerial control of forest insects in Canada. 350
pp. Department of the Environment, Ottawa,
Canada.

Stehr, G.

1954. A laboratory method for rearing spruce bud¬
worm, Choristoneura fumiferana (Clem.),
(Lepidoptera-Torticidae). Can. Entomol.

86: 423-428.

Vail, P. V.; Henneberry, T. J.; Kishaba, A. N.; and
Arakawa, K. Y.

1968. Sodium hypochlorite and Formalin as antiviral
agents against nuclear-polyhedrosis virus in lar¬
vae of the cabbage looper. J. Invertebr. Pathol.
10: 89-93.

Wilson, G. G.

1972. Studies on Nosema fumiferana, a microsporid-
ian parasite of Choristoneura fumiferana (Clem.)
(Lepidoptera : Tortricidae). 108 pp. Ph. D.
thesis, Cornell University, Ithaca, New York.

1973. Incidence of microsporida in a field population
of spruce budworm. Can. For. Serv. Bi-Monthly
Res. Notes 29: 35-36.

1981. Protozoa— Nosema fumiferana, a natural

parasite of a forest pest and its potential for
use in pest management. In H. D. Burges (ed.),
Microbial Control of Pests and Plant Diseases.
A cademic Press, New York.

231

Fractional Colony Propagation

A New Insect-Rearing System

A. Dickerson2

By J. David Hoffman, C. M. Ignoffo, Paula Peters,1 and W.

Introduction

Improvements in performance of insect colonies result
mainly from advances in nutrition, rearing facilities, pro¬
cedures, equipment, and environmental control. But little
emphasis has been placed on one of the most important
factors in insect rearing— selection to improve perform¬
ance of the colonized insect. Fractional colony propaga¬
tion is an insect-rearing concept that involves separate
colonization of many subcolonies started from one parent
colony. Its objectives are to increase management control
and monitoring of the colonies, increase their productivi¬
ty, reduce extremes in population fluctuations, and con¬
trol spread of pathogens and microbial contaminants.

Fractional Colony Propagation

The system using fractional colony propagation was first
applied in 1978 to a colony of cabbage loopers,

Trichoplusia ni (Hiibner), that produced 500 pupae per
day. A new subcolony of T. ni was established each day
for 26 days (26 subcolonies). Each subcolony was
numbered consecutively and reared separately from other
subcolonies. To maintain the genetic diversity of the sub¬
colonies, eggs were obtained from the parent colony on
different days. A rating system, based on one major
criterion (fecundity) and five secondary criteria (hatch,
larval development, pupation, emergence, and disease),
was used to evaluate each subcolony.

All egg sheets were visually rated as either high (80
eggs/6.5 cm2), medium (50 eggs/6.5 cm2), or low (20
eggs/6.5 cm2). These densities were based on the fecundi¬
ty of the parent colony during the 12 months before sub¬
colonization. The number of eggs on sheets accumulated
for each subcolony in a generation was summed to
estimate total fecundity.

Hatch, larval development, pupation, and emergence were
monitored on specific days to evaluate the performance of

'Research entomologist, research leader, and supervisory
biological technician, Biological Control of Insects Research
Laboratory, Agricultural Research Service, U.S. Department of
Agriculture, P.O. Box A, Columbia, Mo. 65205.

2Research entomologist, Boll Weevil Eradication Research Unit,
Agricultural Research Service, U.S. Department of Agriculture,
4116 Reedy Creek Road, Raleigh, N.C. 27607.

the subcolony. Disease and microbial contamination in
the subcolonies were monitored continually. Eggs from
each subcolony were collected and refrigerated (at 12° C,
about 6 days) until percentage of hatch could be
estimated. An egg hatch of more than 90% (based on a
sample from each egg sheet) was acceptable. Rate of lar¬
val development was evaluated on the sixth day after egg
hatch because this stage can be visually estimated more
rapidly and accurately than earlier stages of larval
development. At least 80% of the larvae reared 6 days at
30° C should be in the fourth instar. Rate of pupation
was determined on the 11th day after egg hatch. At least
90% of the population reared at 30° C should have
pupated by this time. A count of dead and living pupae
19 days after egg hatch was used to determine percent¬
age of adult emergence. Adult emergence of less than
90% was considered abnormal. Actual measurements of
the first four secondary criteria were recorded only when
a visually estimated value fell below the expected per¬
formance level. Subcolonies rated for two consecutive
generations below the expected performance level for
hatch, larval development, pupation, or emergence were
replaced with higher-rated subcolonies.

A subcolony with any sign of disease was immediately
discarded and replaced with eggs from a subcolony as far
removed in time as possible from the diseased subcolony
(for example, refrigerated eggs collected 1 week earlier).

As a precautionary measure, any subcolony showing
questionable signs of disease (abnormal growth because
of nutrition, microbial contaminants, or environmental
factors) was also replaced with better subcolonies.

Converting the single-colony propagation system for T. ni
to fractional colony propagation did not require addi¬
tional labor, space, or facilities; nor did it increase rearing
complexity. The workflow for both systems was essential¬
ly identical except that a subcolony was started each day,
and progeny from different subcolonies were never mixed
(table 1).

Effects of the System

The system using colony propagation greatly improved
productivity and stability of the T. ni colony. And the
subcolony system allowed monitoring and manipulating
of the colony from generation to generation. Such
management control was not possible with the single¬
colony rearing system. So the productivity of the colony

232

Table 1. — Comparative flow of work for single- and fractional-colony propagation of Trichoplusia
ni (Hiibner)

Day Work common to both rearing systems Work specific to fractional colony propagation

0 Place eggs with diet; mark with date set up

8 Estimate percentage of 4th-instar larvae
(6 days after hatch).

13 Harvest pupae; estimate percentage of

pupation.

14 Surface-sterilize pupae .

15 Place pupae in emergence-oviposition cage

and date.

18 Initial emergence (females); feed 10% honey
solution daily.

21 Estimate percentage of emergence based on

number of live and dead pupae; collect 1st
egg sheet; date egg sheet; estimate egg
density— high, medium, low; mark egg-
density estimate on egg sheet; record egg-
density rating for each egg sheet; surface-
sterilize egg sheet.

22 Cut egg sample from egg sheet and incubate

at 30° C; refrigerate egg sheet at 1 2° C.

26 Stop collecting egg sheets from each cage
when egg sheet is rated low; estimate per¬
centage of hatch of egg sample, and record.

Mark with subcolony number; never mix eggs
from different subcolonies.

Count larvae if estimate for 4th instar is less
than 80%; record value if below 80%.

Count pupae if estimate for pupation is less
than 90%; record value if below 90%;
never mix pupae from different subcolonies.

Mark with subcolony number.

Never mix moths from different subcolonies.

Count pupae if estimate for emergence is less
than 90%; mark egg sheet with subcolony
number; never allow egg sheets from dif¬
ferent subcolonies to mix.

Determine percentage of hatch by counting a
sample of eggs if estimate of hatch in egg
sample is less than 90%; sum all egg sheets to
estimate total fecundity, and record.

(based on fecundity) increased more than 30% above the
mean within six generations (subcolony selection began
at the third generation). Higher fecundity may possibly
be gained with more selection.

Production stability, measured as fluctuations in fecundi¬
ty, improved rapidly. Before the colony was subcoionized,
average fecundity fluctuated unpredictably (about 30%).
After subcolonization, fluctuation in the average number
of eggs was reduced sixfold within six generations.

Pathogens introduced at low levels usually spread
throughout an entire colony before they are detected and
often cause lower productivity, slower rates of develop¬
ment, and reduced egg hatch. So, subcolonies with these
conditions can be replaced, even before the disease symp¬

toms appear. Fractional colonization minimizes contact
between subcolonies; so it should reduce the spread of a
pathogen. Subcolonies that are contaminated are easily
replaced with another subcolony.

Subcolonies will probably become more homogeneous by
continuous inbreeding. But pooling of progeny (eggs, lar¬
vae, pupae, or adults) from each subcolony for laboratory
or field experiments should help maintain heterogeneity.

Although the full impact of fractional colony propagation
has not yet been fully realized, the increases in manage¬
ment control, productivity, and production stability were
excellent. This system should be applicable to both small
and large rearing programs with two or more setups per
generation period.

233

Production of Insects for Industry

The Dow Chemical Rearing Program

By W. R. Fisher1

Introduction

In the mid-1970’s, Dow Chemical U.S.A. recognized that
its research program for insecticide development would
be improved by increasing the uniformity and quality of
the test insects. The company decided to construct a new
rearing facility at its agricultural chemical research
center in Walnut Creek, Calif. This paper describes the
development of their new program designed for rearing
the western spotted cucumber beetle, Diabrotica undecim-
punctata undecimpunctata Mannerheim, on natural corn
diet and for rearing four lepidopteran species on artificial
diet: codling moth, Laspeyresia pomonella (Linnaeus);
tobacco budworm, Heliothis virescens (Fabricius); beet
armyworm, Spodoptera exigua (Hiibner); and black cut¬
worm, Agrotis ipsilon (Hiifnagel).

The previous program had been beset by problems that
were impractical to solve in the existing facility.

Microbial contamination was the most critical problem.
Disease destroyed beet armyworm colonies and
significantly reduced yields of tobacco budworm. The
spread of disease among colonies was always a threat in
the small holding rooms that contained more than two
species. A colony devastated by disease had to be rees¬
tablished from clean stock, a process that took a month
or more. Sublethal infections reduced colony vigor and
resulted in test organisms that were poor in quality and
abnormally susceptible to chemical challenge. Contami¬
nants, such as mold, thrived on the artificial diets used
to rear tobacco budworm and codling moth. The fungi
covered much of the diet surface, competed with larvae
for nutrients, and caused relative humidity in rearing con¬
tainers to be above optimal levels for the insects, thereby
reducing their yields and quality.

All microbial contaminants significantly increased the
time, labor, and money needed to satisfy colony and
research requirements. Contamination was difficult to

’Formerly a biologist with Dow Chemical U.S.A., Walnut Creek,
Calif. Now a graduate research assistant with the Department
of Entomology and Nematology, University of Florida,
employed through a cooperative agreement with the Insect At-
tractants, Behavior, and Basic Biology Research Laboratory,
Agricultural Research Service, U.S. Department of Agriculture,
Gainesville, Fla. 32604.

control in the old facility for several reasons. Most impor¬
tantly, artificial diet was prepared in an area that was
not adequately isolated from larval holding rooms. In
fact, containers with pupae from these rooms were taken
through this area to be harvested, exposing freshly
poured diet to contaminants. Inadequate filtration of con¬
ditioned air, unrestricted movement of employees and
other persons, poor sanitary procedures, and an inability
to sterilize reusable containers also contributed to the
problem.

Health hazards, inherent in any rearing program, were
not adequately controlled in the old facility. For example,
fungi growing on artificial diets produced potentially in¬
fectious spores that were inhaled by workers harvesting
pupae or washing containers. Wing scales from lepidop¬
teran oviposition cages and airborne dusts generated dur¬
ing mixing of artificial diets caused allergic symptoms in
some employees.

Finally, procedures for rearing certain species were not
compatible with an efficient and sanitary program. For
example, codling moth larvae were reared in plastic con¬
tainers covered by loose-fitting glass lids. The plastic was
heat labile and could not be sterilized in a steam auto¬
clave. The lids allowed late-instar larvae to escape into
the holding room where they would pupate in walls and
shelves, making sanitation difficult. Furthermore, to
reduce desiccation, the surface of the artificial diet used
for the codling moth larvae was coated with melted paraf¬
fin. But, in the process, wax would build up on the inside
of the square-cornered containers and make sterilization
by immersion in sodium hypochlorite impossible. So mold
was common, and yields were low, thus necessitating
more containers and labor to satisfy research needs.

These experiences with the previous program provided a
basis for development of the new rearing facility. Five
major factors were considered during its design: isolation
of critical work areas, the flow of materials and products,
room characteristics, equipment, and regulations and pro¬
cedures.

234

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Solid circles— dirty work area. Cross-hatching— isolated room for holding insects. PT— passthrough.
SA— steam autoclave. HR— insect holding room. MR— mechanical room containing motor-control
center, water heater, boiler, telephone panel.

Development of the Dow
Rearing Facility

Isolation of critical work areas

Isolation to maintain cleanliness in a rearing facility re¬
quires separation of operations, products, and materials.
It also requires establishment of barriers to prevent
spread of contaminants throughout the building. To get
such isolation, the Dow facility was divided into three
major sections. The first and most critical, the clean work
area, consists of clean-storage, diet-preparation, and

surface-sterilization rooms (fig. 1). Only uncontaminated
materials and insect eggs and pupae sealed in clean con¬
tainers are allowed into this area. Sterilized, reusable con¬
tainers and cages and other materials are kept in the
clean-storage room. In the diet-preparation room, dietary
ingredients are weighed and mixed, and diets are dis¬
pensed into containers and planted with eggs. Also in
this room, surface-sterilized pupae are distributed into
oviposition cages. Insect eggs and pupae are surface-
sterilized with sodium hypochlorite in the room next to
the diet-preparation room.

235

The next level of isolation consists of eight holding rooms
where insects are kept during larval development and
adult oviposition. Rooms 4 and 5, the most isolated, are
used to maintain parasites or predators for studies of
their susceptibility to insecticides. This area also serves
to isolate potentially diseased insects collected from wild
populations before they are infused into production col¬
onies. Rooms 1-3 and 8 are used to hold adults and
developing larvae reared on artificial diets. The western
spotted cucumber beetle, the only species reared on
natural diet, is maintained in holding rooms 6 and 7.

The final area of isolation is the room for pupal harvest
and container sterilization. Because the operations here
involve dirty containers, a high potential exists for the

release of contaminants into the rest of the facility. So
four ultraviolet-light passthroughs and a double-door,
steam autoclave were installed to allow movement of
products and materials in and out of this room without
spreading contaminants. All tasks that liberate con¬
taminants, such as opening larval containers and
harvesting pupae, are done inside a fume hood.

Flow plan of materials and products

The flow plan (fig. 2) provides a logical, one-way move¬
ment of materials and products that links isolation areas
in the facility. Containers from clean storage are taken to
the diet-preparation room, filled with diet, and planted
with eggs. They are then distributed to holding rooms

236

where they remain until completion of larval develop¬
ment. Next, containers are passed to the harvest room,
where pupae are removed from the diet. Reusable con¬
tainers are washed, sterilized, and transferred into the
clean-storage room. Pupae are placed in clean cups with
tight-fitting lids and taken to the surface-sterilization
room where they are washed in sodium hypochlorite,
placed in cages, and taken to the appropriate holding
room for adult eclosion and oviposition. Sheets containing
eggs are collected from these cages, placed in clean
plastic bags, and taken to the surface-sterilization room.
Rooms 2 and 3 are next to the pupal harvest area and
hold larval stages that are susceptible to pathogens and
that have the greatest potential for facility contamina¬
tion. Direct access to the harvest room via passthroughs
with germicidal lamps eliminates spread of contaminants
during transfer of containers before pupal harvest.

Because of the unsanitary conditions associated with in¬
sects raised on natural diet, rooms 6 and 7, where
western spotted cucumber beetles are reared, are not part
of the internal flow plan. Instead, the inside doors have
been sealed, and the only access is from the outside. All
needs for this species, except for environmental control
and monitoring, are met independently of the artificial-
diet program. For example, washing larval containers and
preparing them for reuse is done under a terrace that
runs the length of the building from the mechanical room
to holding room 4. A sink in room 6 enables workers to
harvest and sanitize eggs without risking contamination
to other areas of the facility.

Room characteristics

Separation of species, and of different stages of the same
species, is achieved in eight larval and adult holding
rooms. Each room has individually controlled
temperature, relative humidity, and lighting. Light fix¬
tures are sealed to exclude insects. Each room has a hose
bib and drain for washing walls and floors. Floor covering
throughout the facility is seamless, troweled epoxy that
extends 6 inches up the walls to provide coved corners
for easy cleaning. Walls and ceilings are painted with an
epoxy paint that resists abrasion and that is inert to
sanitizing solutions. Portable racks for holding rearing
containers facilitate inspection of insects, sanitation, and
proper airflow.

Equipment

Equipment was selected to compliment facility design
and flow plan and to provide a safe and healthy working
environment. In the diet-preparation room, a 13-liter mix¬
er is operated inside a fume hood to protect workers from
allergenic dusts generated during preparation of diets.

For larger batches of diet, an 80-liter, steam-jacketed

mixer is used. A hood in the pupal-harvest room removes
spores from mold growing on diet and insect wing scales
released during cleanup of oviposition cages. A collector
in room 8 removes scales from oviposition cages of tobac¬
co budworm moths.

Conditioned air is moved in the facility by two air¬
handling units located on the roof. One unit supplies
high-humidity air (65%-80%) to adult oviposition rooms,
while the other provides lower humidity requirements for
larval holding rooms and work areas. All supply air is
filtered through an absolute filter that removes 99.9% of
all particles greater than 0.3 pm in size. Conditioned air
entering the harvest room is exhausted to the outside
and not returned to the rest of the facility because of the
contamination potential. The air-conditioning system is
pneumatically controlled from a panel in the hall. En¬
vironmental variables are monitored by a data logger/
recorder programed with high and low limits for each
room. Significant variation from these limits is indicated
by an audible signal, and detailed emergency procedures
are outlined for workers who respond to the alarm. Opera¬
tions of the air handlers, the chiller, the boiler, and the
pneumatic control system are also monitored. If a break¬
down occurs during nonworking hours, telephone equip¬
ment automatically dials an operator who identifies the
problem and notifies the insectary manager.

Sanitation equipment provides a firstline defense against
introduction of contaminants into insect colonies. Ultra¬
violet lamps, mounted on ceilings throughout the clean
isolation area, are automatically turned on during non¬
working hours to help reduce incidence of surface and air¬
borne contamination. The autoclave has double doors for
loading unsterile materials from the harvest-room side
and for removal on the clean-storage side after steriliza¬
tion is complete. The passthroughs were designed and
constructed at Dow. Each is 0.8 by 0.6 by 0.8 m and is
made of plywood covered with Fiberglas cloth and resin
for strength. Doors on both ends are made of safety glass
in an aluminum frame. The 15-watt germicidal-lamp fix¬
tures installed inside the passthroughs are turned on for
30 minutes with a timer switch. To protect employees
from skin or eye damage during illumination, spring-
loaded switches shut off the lamp when either door is
opened. The passthroughs allow containers to be moved
between isolation areas and reduce the spread of con¬
taminants. In fact, agar-plate counts have shown com¬
plete destruction of black-mold spores during operation of
the passthrough.

Regulations and procedures

Regulations have been established to insure that sanitary
conditions are maintained. No job is complete until all
materials used are put away inside drawers or cabinets

237

Figure 3.— Codling moth larval rearing con¬
tainer with diet, pupation sites, and surface-
sterilized egg sheets taped to the inner surface
of the lid.

and work surfaces are sprayed with disinfectant and
wiped clean. Clean work areas are vacuumed daily and
mopped with germicidal detergent twice a week. Access
by employees to the clean area is restricted to only those
who have duties there. Individuals exposed to disease
organisms, for example when conducting field trials, are
not allowed in the building. Lab coats and plastic aprons
used during work in the harvest room are not allowed to
be worn elsewhere in the insectary. Each isolation area
has the materials and equipment required to complete
jobs designed for those areas, so movement of items be¬
tween areas is reduced. For example, there are three lab¬
oratory carts in the facility to service the various rooms
in the major isolation areas. Materials or products are
removed from carts and placed in passthroughs when be¬
ing transported to another area. Larvae or adults that
have escaped from cages are immediately discarded. If
they are accidently stepped on, the area is sprayed with
disinfectant and thoroughly cleaned. Undesirable insects
like cockroaches are controlled with sticky traps or boric
acid, which do not affect colony insects.

Rearing procedures were developed to provide the most
efficient use of space and time. For example, a better pro¬
cedure for rearing the codling moth was developed. The
problems with the old procedure were largely related to
the square containers, loose lids, and paraffin-coating
method. The new procedure is based on a new larval con¬
tainer. The container (fig. 3) has a modified snap-on lid
that provides adequate gas and moisture exchange while
preventing larval escape. Wax-paper strips containing
eggs are taped to the underside of the lid, and newly
hatched larvae drop to the artificial diet in the container
and begin feeding. Prepupae leave the diet and pupate in
a sterile, corrugated cardboard strip lining the inside

Figure 4.— Racks for surface sterilization of
eggs laid on wax-paper substrates. Separation
of egg sheets within racks insures direct con¬
tact with sterilant.

perimeter of the container. The strip is then rolled up and
placed in a cage for eclosion and oviposition. The round
container is easier to clean and sterilize than the old con¬
tainers with square corners. Cleanup is also made easier
by a new way to coat the diet with melted paraffin. While
still hot, the diet is swirled 1-2 inches up the side of the
container where it cools in a thin sheet. Melted wax is
then applied to the diet surface by means of a soft
camel’s-hair brush, while the sheet of diet on the side
keeps wax from touching the container. This new pro¬
cedure has resulted in a fivefold increase in pupal yield
and a 75% reduction in cleanup time. Fungal contamina¬
tion of the diet has been totally eliminated.

Another procedure developed for the Dow program is a
surface-sterilization technique for codling moth and beet
armyworm eggs laid on wax paper. In the old procedure,
layers of egg-laden wax paper were stacked and placed in
a beaker containing a dilute bleach solution. But the
waxy surfaces resisted wetting, making the disinfection
process ineffective. The procedure also required frequent
handling that damaged eggs or knocked them off the
sheets. In the new procedure, egg sheets are placed in
specially designed racks (fig. 4) that effectively separate
sheets, insuring complete contact with the sterilizing
solution. After sterilization, the racks are placed in two
successive tanks containing clean water. Rinsing the
sheets in this manner requires no running water; so about
11,000 liters/year are saved. After rinsing, the rack is
removed, and the sheets are allowed to dry, a process
hastened by separation of the sheets. This procedure also
eliminates unnecessary handling of egg sheets.

238

Evaluation of the Dow
Rearing Program

Once the new program began, the quality of all insects
improved significantly, and yields became consistent and
predictable. Disease symptoms were eliminated. Reduc¬
tion in disease and dietary contamination allowed elimina¬
tion of fungicides, Formalin (formaldehyde), and other
antimicrobials from the diet. Symptoms of dietary defi¬
ciencies and inadequate environmental conditions that
had previously been masked by disease symptoms be¬
came apparent, and steps were taken to eliminate then-
causes. Yields increased, in some cases dramatically, and
have remained consistent from one generation to the
next. So the number of containers needed to meet colony
and research requirements was reduced for all five species
at a significant savings in man-hours and raw materials.
Because of more uniform development rates, testing

results became more reliable. Finally, health and safety
hazards were significantly reduced by the use of equip¬
ment and procedures designed to provide a cleaner work
environment.

The facility, equipment, and procedures in the new pro¬
gram allowed these improvements to be made. But the in¬
sectary manager made the program a success by coor¬
dinating activities, training personnel, anticipating and
solving problems, and regularly improving the program. I
discuss the role of an insectary manager in such a pro¬
gram in “The Insectary Manager.”

Acknowledgments

I thank Jeanne Wiegand, Department of Veterinary
Science, University of Florida, for her comments and
review of the final draft of this manuscript.

239

Industrial Insect Production
for Insecticide Screening

By R. E. Wheeler1

Introduction

The main objective of insect production in the pesticide
industry is to rear enough organisms to use in screening
for new insecticides. The insects may also be used to de¬
tect insect-growth regulators, sterilants, attractants, and
repellents. In comparison to large industrial or govern¬
ment facilities that are mass producing insects to use in
biological-control programs, pesticide-research laborato¬
ries usually have smaller rearing facilities, lower produc¬
tion quotas, and lower operating and equipment budgets.
But the pesticide-research laboratories usually rear a
broader spectrum of insect types. What types and num¬
ber of species are reared depend on economic and environ¬
mental concerns and on trends in pest resistance to
chemicals. And, whether a program is large or small, its
success depends on having a facility designed for insect
rearing and on having a management commitment to ade¬
quately staff and fund it. Quality insects can be produced
only if these requirements are met. And a successful
chemical screening program for insecticides depends on
having quality insects. This paper outlines the guidelines
and procedures we have developed at the Chevron Chemi¬
cal Co., Richmond, Calif., for species selection; colony
establishment, maintenance, and quality control; produc¬
tion and use; and product quality control. Following these
suggestions should help insure a successful rearing
program.

Selection of Species
for Rearing

Industrial pesticide-screening laboratories test many com¬
pounds in several chemical classes of unknown pesticidal
activity. Since insect species vary in their susceptibility
to any one insecticide class, it is desirable to screen
against many insect types to avoid missing a potentially
useful insecticide. Selection of species for rearing must
consider insecticide resistance and which target species
and index species are feasible to use in the particular pro¬
gram.

‘Lead scientist, entomology, Chevron Chemical Co., 940 Hensley
St., Richmond, Calif. 94804.

Insecticide resistance

The selection of the degree of insecticide resistance in the
insect population used to start a colony depends on the
test methods used for the initial screen. When hundreds
of unknown compounds are being screened for the first
time, it is more practical to test compounds at a single
high dose. If the insect population used to start a colony
is too susceptible, too many false leads are obtained. For
such single-dosage screening tests, it is desirable to select
insect populations with insecticide resistance similar to
that of the target-species populations encountered in agri¬
culture. When initial or advanced screening methods use
serial dilution dosages, the degree of insect resistance is
less critical since dosage-response curves can be plotted,
ranked, and compared to known standard insecticides.

Target species

When possible, the insects to be screened should include
medically and agriculturally important target species.
Selection can be based on the relative economic impact of
the pests. But certain species are routinely used as test
organisms because they produce a particular physiologi¬
cal, morphological, or behavioral response. For example,
the yellow mealworm, Tenebrio molitor Linnaeus, and the
reduviid, Rhodnius prolixus Linnaeus, produce highly
quantitative and qualitative responses to juvenile hor¬
mone mimetics (Bowers and Thompson 1963, Wiggles-
worth 1969); and the American cockroach, Periplaneta
americana Linnaeus, is useful for electrophysiological
measurements of insecticidal activity (Narahashi and
Yamasaki 1960a, 1960b, 1960c). Even parts of insects can
be used— for example, antennae for electroantennogram
studies in pheromone research (Schneider 1962, Davis
1973, Roelofs 1977).

Index species

In some instances, it is not feasible to rear the target
species, because of time and expense or because its avail¬
ability is limited by its distribution. So a substitute index
species may be needed. The term “index species” refers
to a species known to demonstrate susceptibility to stand¬
ard known insecticides like that of a phylogenetically
related target species. In California, because of quaran¬
tine restrictions, the western spotted cucumber beetle,
Diabrotica undecimpunctata undecimpunctata Manner-
heim, is commonly used as an index species for Dia¬
brotica com rootworms; these species have similar

240

Table 1. — Species reared and the developmental stages used for screening potential insecticides at
Chevron Chemical Co., Richmond, Calif.

Order and species _ _ Stages used

Acari: Twospotted spider mite, Tetranychus urticae Koch . Adult.

Coleoptera:

Confused flour beetle, Tribolium confusum Jacquelin du Val Adult.

Egyptian alfalfa weevil, Hypera brunneipennis (Boheman) . Larval, adult.

Granary weevil, Sitophilus granarius (Linnaeus) . Adult.

Red flour beetle, Tribolium castaneum (Herbst) . Adult.

Sawtoothed grain beetle, Oryzaephilus surinamensis (Linnaeus) . Adult.

Western spotted cucumber beetle, Diabrotica undecimpunctata undecimpunctata
Mannerheim. Egg.

Yellow mealworm, Tenebrio molitor Linnaeus Pupal.

Diptera:

House fly, Musca domestica Linnaeus Adult.

Yellowfever mosquito, Aedes aegypti (Linnaeus) Larval, adult.

Hemiptera: Lygus bug, Lygus hesperus (Knight) . Adult.

Homoptera: Cotton aphid. Aphis gossypii Glover . Adult.

Lepidoptera:

Beet armyworm, Spodoptera exigua {Hiibner) . Larval.

Cabbage looper, Trichoplusia ni (Hiibner) . Larval.

Tobacco budworm, Heliothis virescens (Fabricius) Larval.

Orthoptera:

American cockroach, Periplaneta americana (Linnaeus) . Nymph.

German cockroach, Blattella germanica (Linnaeus) Adult.

biology and similar susceptibility to standard rootworm
insecticides. The Egyptian alfalfa weevil, Hypera brun¬
neipennis (Boheman), serves as an index species for the
geographically restricted boll weevil, Anthonomus gran-
dis grandis Boheman, and as a target species in itself.

In most cases, industrial insecticide-screening labora¬
tories routinely rear at least seven insect species, in¬
cluding representatives of the orders Diptera, Coleoptera,
Lepidoptera, Homoptera, Hemiptera, and Orthoptera;
usually, one mite species is also reared (for examples, see
table 1). Often, small colonies of other species, such as
stored-product beetles and moths, are maintained and
then enlarged when special projects call for their use.

Colony Establishment

Before an insect species is colonized, its life history,
known rearing techniques, physical requirements, and
probable insecticide resistance should be thoroughly
studied. Important considerations are population source
and population size.

Population source

Insects used to start colonies of agriculturally and
medically important target species should be collected
from their preferred host crop or natural habitat. And the

amount and kinds of insecticides used in the collection
area should be ascertained. Often, other insectaries will
provide a subculture with documented background
information on insect origin, time in culture, and insec¬
ticide susceptibility (Dickerson et al. 1980). Use of a sub¬
culture should reduce the likelihood of introducing a
pathogen or other contaminant into the culture. Field-
collected material should always be reared in isolation to
insure freedom from contaminants before introduction to
the main insectary.

Population size

The sample size collected to establish a colony should be
large enough to meet the minimum sustainable popula¬
tion size that will yield, after three generations, enough
insects for preliminary standards testing. These pre¬
liminary data will indicate whether or not the selected
population possesses the desired level of susceptibility to
insecticides. The sample size is also guided by the fecun¬
dity of the species. As a general rule, 300 to 500 in¬
dividuals at a given stage are enough to start a colony.
To increase the chances of obtaining maximum homozy¬
gosity in this size of sample, only a single developmental
stage should be collected. For highly vagile insects, this
method minimizes chances of sampling two or more pop¬
ulations that have merged from diverse locations.

241

Rearing facilities

The insect-rearing system at the Chevron Chemical Co.
uses a series of six rooms isolated from a central hallway
by anterooms. Three of the rooms are used to rear several
species that have similar environmental requirements.
Even so, careful planning and procedures minimize the
chance of disease transmission.

The ‘‘public-health insect” room (kept at 31.8° C) contains
the German cockroach, Blattella germanica (Linnaeus);
the American cockroach; the house fly, Musca domestica
Linnaeus; and the yellowfever mosquito, Aedes aegypti
(Linnaeus). Colonies are serviced by being transferred to
an adjacent workroom through a small, sliding
passthrough door. This workroom is maintained at a
lower temperature than that in the ‘‘public-health insect”
room to provide a more comfortable environment for the
technicians.

The lepidopteran room contains cabbage loopers, Tri-
choplusia ni (Hiibner); tobacco budworms, Heliothis
virescens (Fabricius); and beet armyworms, Spodoptera
exigua (Hiibner). The larval stages of these species are
reared in the same room. Pupae are removed to another
isolated room for adult emergence and oviposition. This
room is equipped with exhaust hoods for removing adult
moth wing scales. The workroom for preparing artificial
diet is also isolated from the rearing rooms. Both the
diet-preparation and rearing rooms are designed for easy
cleanup.

Other multiuse rooms contain several grain pest species;
the Egyptian alfalfa weevil; and the lygus bug, Lygus
hesperus (Knight).

Each of the other rooms contains only one species be¬
cause of their space and environmental requirements and
to prevent cross contamination. Greenhouses and outside
beds provide space for raising host plants such as lima
beans for the twospotted spider mite, Tetranychus ur-
ticae Koch; cucumbers for the cotton aphid, Aphis
gossypii Glover; and alfalfa for the Egyptian alfalfa
weevil.

Colony management

How a particular colony is managed depends on which
stages will be used in testing. With rapidly reproducing
species such as the cotton aphid and twospotted spider
mite, schemes for colony and production management are
the same. Insects in all stages of development are trans¬
ferred to new host plants once a week after peak use
demands have been satisfied. For those species whose
adult stage is the one tested (table 1), a small part of the

adult yield is reserved to maintain the colony, and the
rest is used to meet the production quota. Similarly, for
those species whose immature stages are the ones tested
(table 1), part of the harvest is reserved to maintain the
colony.

Colony quality control

Colony homozygosity.— When the life cycle of the insect
is longer than 1 week and a particular stage is needed
once a week (as it is for house flies or lepidopterans), the
colony must be divided into several developmental shifts.
But dividing the colony creates a series of subcolonies.

To maintain homozygosity among the subcolonies, we
routinely reserve some eggs or pupae from one subcolony,
place it in suitable cold storage for 1 week, and then
backcross it with the next shift at a ratio of one stored to
four new.

Biometric measurements.— Biometric measurements on
egg production, length of development, size and weight of
test stages, sex ratios, and yield of adults are important
indicators of consistent rearing procedures. For the
dosage-mortality responses to an insecticide to be con¬
sidered accurate, these measurements must be within es¬
tablished ranges.

Environmental control

After many years of experience in designing environ¬
mental-control rooms for rearing insects, we have
developed several design guidelines that are often
overlooked in the basic design of commercial pre¬
fabricated environmental rooms. Equipment for
establishing air quality (cleanliness, temperature, and
relative humidity) should be housed outside the insectary.
Preconditioned air should be continuously introduced into
the insectary (not recirculated) so it creates a slight
positive pressure at all room-entry points. This pressure
reduces the possibility of airborne contaminants entering
the insectary from outside through cracks, doors, or win¬
dows. The air-temperature-conditioning system (for both
heating and cooling) should be one that gives the least
amount of Btu’s to achieve the desired temperature. To
regulate the Btu inputs, solid-state proportional ther¬
mostats can be used to regulate steam-heating valves,
electric-resistant load heaters, and refrigeration bypass
valves.

Because environmental conditions in an insectary are not
necessarily the same as the microclimate in the insect¬
rearing cage or container, sensing elements should be
placed inside the insect-rearing cage to determine actual
environmental conditions. And use of a programable data
logger to monitor important environmental conditions is

242

invaluable. Such systems provide computer-stored and
printed data records and can automatically signal con¬
ditions that may endanger or disrupt the insect colonies.

Diet quality control

Quality control of artificial diet is maintained by closely
following prescribed preparation methods and by using
materials of known origin and consistent quality. If re¬
quested, most manufacturers will inform consumers of
significant changes in composition or quality of their
product. It is important to observe the stability of the
shelf life for perishable components of diets, such as
vitamin mixes and wheat germ. These items, particularly,
should be placed in cold, dry storage to maintain max¬
imum potency.

Natural diets consist mainly of host plants grown in
greenhouses or outside beds. To obtain plants of con¬
sistent maturity, planting schedules should be shifted
periodically to adjust for seasonal changes in growth
rates. Perennial host plants such as alfalfa may require
protection from insect and mite pests and from parasites
and predators. The Chevron Chemical insecticide Dibrom2
is very useful for cleaning up or eradicating pests from
host plants. Dibrom is a broad-spectrum, contact insec¬
ticide exhibiting less than 2-day residual life on plants.

Contamination control

Facility design and material flow is especially important
in keeping the insects as free from contamination as pos¬
sible (see “Production of Insects for Industry. The Dow
Chemical Rearing Program,” by W. R. Fisher). Insect¬
rearing cages are also critical, and disposable ones should
be used whenever possible. Disposable cages are par¬
ticularly important for insects that are susceptible to
pathogens and where rearing procedures are used that
allow contamination of the insects, diet, and containers.
Permanent or complexly designed cages should be
constructed of stainless steel or polycarbonate plastics,
which can withstand rigorous cleaning procedures such
as autoclaving or steam cleaning.

Production and Use

Most industrial laboratories operate a complex initial
screening schedule for herbicidal, fungicidal, insecticidal,
and plant-growth regulator activity. At Chevron Chem¬
ical Co., 7 of the 18 insect species routinely reared are
used in the initial screening that requires 2,000-4,000 in¬
dividuals of a certain stage and age for each species on a

2l,2-Dibromo-2,2-dichloroethyl dimethyl phosphate.

specified day of the week. Though the schedules and pro¬
cedures for colony and production maintenance in a 5-day
workweek often overlap considerably, the schedule for
production maintenance is more flexible and can be
changed according to how many insects are needed. The
biology of the insect must be thoroughly understood if in¬
sect production and testing schedules are to be success¬
fully synchronized. Those exogenous factors, such as
temperature, humidity, and food, that regulate the rate of
insect development and its quality must be identified and
controlled.

The egg stage is a convenient point to synchronize pro¬
duction and utilization schedules. In our insectary, we
use this stage to synchronize the production of yellow-
fever mosquito larvae, Egyptian alfalfa weevils, cabbage
loopers, and American cockroaches. The cotton aphid,
western spotted cucumber beetle, and twospotted spider
mite colonies provide a continuous supply of all stages of
development. With colony maintenance on a 5-day
workweek, pupae harvested from the house fly colony,
which is maintained at 32° C, normally emerge on a Sun¬
day morning. To provide 2-day-old adults for testing on
Wednesday, some of the pupae are placed in adult emer¬
gence cages and incubated at 22° C. So adult emergence
comes 1 day later than it does in the stock colony.

Product Quality Control

Quality control is embodied in every aspect of insect pro¬
duction from the physical design of the facility and pro¬
duction procedures to the insects themselves. For us, the
most important quality characteristics are those showing
that the insects have the required susceptibility to insec¬
ticides. The dosage-mortality response of the test insect
to standard insecticides is the primary reference point for
determining insect quality. Accordingly, one or two
standard insecticides are routinely included in all tests on
candidate insecticides. At regular intervals, a colony is
monitored by tests of dosage-mortality response to a
broad spectrum of insecticide types and classes. Through¬
out the entire period that a colony is maintained, a data
bank containing standard data on insecticide dosage-
mortality is kept and periodically analyzed for drifts in
insecticide susceptibility. (Figures 1-3 illustrate how
much colony susceptibility can fluctuate.)

The baseline LD50/90 values are first established after the
data from the initial series of tests indicate that a new
colony’s standard insecticide response has stabilized.
When a long-established colony’s LD50/90 tends to shift
beyond the standard deviation, close examination of the
rearing procedure and biometric measurements may
sometimes indicate reasons for the shift, particularly if
there are changes in insect body mass and sex ratios. A
trend toward increased susceptibility indicates genetic

243

changes caused by lack of insecticide pressure on the col¬
ony or accidental introduction of susceptible insects into
the colony. There are several ways to counteract such
changes: apply insecticide pressure to the colony, infuse
the colony with resistant field-collected specimens, or
establish a new colony with the desired level of resist¬
ance. The method we use depends on the species and the
seasonal availability of field specimens.

Occasionally, colony survival may be threatened by fail¬
ure of environmental-control equipment, disease, invasion
by other arthropods, or contamination. For example, in
1972 our cotton aphid colony was infested by a hymenop-
terous parasite. But we were able to reestablish this col¬
ony from a few (300) unparasitized aphids. Afterwards,
dosage-mortality data from Dibrom tests with progeny
from these individuals indicated an increase in tolerance.
This increase continued for about 6 months; then the
tolerance began to return to its previous level (fig. 1).

It is important that all colonies of a given species re¬
spond to chemical effects in the same way. For example,
over the last 15 years, Chevron Chemical Co. has main¬
tained a strain of twospotted spider mites resistant to
parathion.3 When a change in the LD50 response to
parathion indicated increased susceptibility, the entire
colony was subjected to parathion treatments until the
dosage response stabilized (fig. 2). Likewise, how diet
changes affect insect quality should always be examined.
Several years ago, at Chevron Chemical, the agar compo¬
nent of the caterpillar diet was replaced by another gel¬
ling agent— Gelcarin HWG (Marine Colloids Division,
FMC Corp., Springfield, N. J.). Examination of dosage-
mortality data from caterpillars reared on the new diet in¬
dicated no changes in insecticide susceptibility (fig. 3).

Acknowledgments

I wish to express my appreciation to Jed Harrison of
Chevron Chemical Co. for preparing the illustrations.

References

Bowers, W. S., and Thompson, M. J.

1963. Juvenile hormone activity: effects of

isoprenoid and straight-chain alcohols on in¬
sects. Science 142: 1469-1470.

Davis, E. E.

1973. An electrophysiological approach to mosquito
control. Res. /Dev. October 1973, pp. 32-36.

Dickerson, W. A.; Hoffman, J. D.; King, E. G.; Leppla,

N. C.; and ODell, T. M.

1980. Arthropod species in culture in the United
States and other countries. 93 pp. Entomo¬
logical Society of America, College Park, Md.

Narahashi, T., and Yamasaki, T.

1960a. Mechanism of the after-potential production
in the giant axons of the cockroach. J.
Physiol. (London) 151: 75.

1960b. Mechanism of increase in negative after¬
potential by dicophane (DDT) in the giant ax¬
ons of the cockroach. J. Physiol. (London)

152: 122.

1960c. Behaviors of membrane potential in the

cockroach giant axons poisoned by DDT. J.
Cell Physiol. 55: 131.

Roelofs, W. L.

1977. The scope and limitations of the electroan ten-
nogram technique in identifying pheromone
components. In N. E. McFarlane (ed.), Crop
Protection Agents— Their Biological Evalua¬
tion, pp. 14-165. Academic Press, New York.

Schneider, D.

1962. Electrophysiological investigation on the
olfactory specificity of sexual attracting
substances in different species of moths. J.
Insect Physiol. 8: 15-30.

Wigglesworth, V. B.

1969. Chemical structure and juvenile hormone ac¬
tivity: comparative tests on Rhodnius pro-
lixus. J. Insect Physiol. 15: 73-94.

Figures 1-3 follow on pages 245-247.

*0, 0-Diethyl 0-(p-nitrophenyl) phosphorothioate.

244

1972 1973 1974 1975 1976 1977 1978 1979

I I I - 1 - 1 - 1 - 1 - 1 —

E

o

■Q

b

Figure 1.— 24-hour LD50 values for standard insecticides against various insects from 1972 through
1979. Applications: house flies— dosages (p/m ai in 70 ul acetone) sprayed on each of 3 replicates of
20 adult insects (150 tests); American cockroaches— dosages (p/m ai in 70 pi acetone) sprayed on
each of 3 replicates of 10 fifth-stage insects (200 tests); German cockroaches and alfalfa
weevils— dosages (p/m ai in 70 pi acetone) sprayed on each of 3 replicates of 10 adult insects (200
tests); cotton aphids— aphid-infested cucumber leaves dipped in aqueous formulation (150 tests).
Dotted lines fall between upper and lower 95% confidence limits. X=time when the aphid colony
was parasitized by hymenopterous wasps and reestablished from 300 mother-aphid isolates.

245

1970 1971 1972 1973 1974 1975 1 976 1977 19 7 8 1979

r— — i i 5f x 1 x 1 1 1 jT-1 r—

Figure 2.— 24-hour LD60 values for standard insecticides against adult twospotted spider mites from
1970 through 1979. Applications: mite-infested lima bean leaves dipped in aqueous formulations.
Dotted lines fall between upper and lower 95% confidence limits. X (on horizontal axis) = time when
entire mite colony was treated with a parathion spray (80 p/m).

246

1974 1975 1976 1977 1978 1979

“I 1 1 T - x - 1 - I —

Figure 3.— 24-hour LDS0 values for methyl parathion against third-stage larvae of various insects
from 1974 through 1979. Applications: cabbage loopers— cucumber leaves were dipped in an aqueous
formulation and then implanted with cabbage looper larvae (150 tests); tobacco budworms and beet
armyworms— cotton leaves were dipped in an aqueous formulation and then implanted with tobacco
budworm or beet army worm larvae (150 tests). Dotted lines fall between upper and lower 95% con¬
fidence limits. X (on horizontal axis)=time when Gelcarin HWG was substituted for agar agar in the
lepidoperous larval diet.

247

Mass Rearing the Cabbage
Looper, Trichoplusia ni

By N. C. Leppla,1 P. V. Vail,2 and J. R. Rye3

Introduction

Laboratory-reared cabbage looper, Trichoplusia ni
(Hubner), moths are needed to perform specific functions
(as, for example, in the sterile-insect technique); and im-
matures are used as hosts for the production of parasites,
parasitoids, predators, and pathogens (viruses, bacteria,
fungi, protozoa, etc.). All stages are sources of biological¬
ly active compounds (hormones, pheromones, enzymes,
etc.). The capability of producing thousands of adult cab¬
bage loopers per day has been developed (Henneberry
and Kishaba 1966 described this technology). Currently,
however, more work is needed to devise better rearing
methods that will produce moths that are qualitatively
uniform and behaviorally effective. Developing better
methods depends on a thorough analysis of past efforts,
application of available technology, and expansion of
research to determine the usefulness of alternative rear¬
ing methods. This paper discusses innovations now in use
and some of the promising ideas currently being studied.

Rearing Techniques

Diets

Originally, diets for rearing cabbage looper larvae were
adapted from some that had been developed for other
species; these were based on agar and plant material (for
example, the diets described by Bottger 1942, Adkisson
et al. 1960, Berger 1963, and Shorey and Hale 1965).
Which specific ingredients to use varies with what
materials are locally available, the kind of mixing equip¬
ment, whether the diet ingredients can be used for other
purposes (so that bulk orders would be appropriate), and
personal preference of the scientists as limited by the
rearing system. So products such as macerated host
leaves, wheat germ, softened lima beans, ground pinto
beans, and alfalfa meal have all been used in the basic

‘Research entomologist, Insect Attractants, Behavior, and
Basic Biology Research Laboratory, Agricultural Research Serv¬
ice, U.S. Department of Agriculture, P.O. Box 14565,

Gainesville, Fla. 32604.

“Research entomologist, Stored-Product Insects Research
Laboratory, Agricultural Research Service, U.S. Department of
Agriculture, Fresno, Calif. 93727.

“Biological technician, Insect Attractants, Behavior, and Basic
Biology Research Laboratory.

diet, alone or in various combinations. (See table 1 for
lists of two of the least expensive diet regimes.)

The basic technqiues of diet preparation have changed lit¬
tle. But procedures were simplified when the addition of
chemical preservatives to the diet was developed to
reduce the need for heat sterilization. Henneberry and
Kishaba (1966) reviewed the antimicrobials suitable for
use with cabbage loopers and presented detailed instruc¬
tions for incorporating 1,800-2,000 p/m (parts per million)
of sorbic acid and methyl p-hydroxybenzoate
(methylparaben) preservatives into a diet of agar and
alfalfa leaf meal. They also used Aureomycin
(chlortetracycline) and Formalin (formaldehyde).

Table 1. — Two of the least expensive diet regimes for rearing cab¬
bage looper larvae

Ingredients1

Diet

Pinto bean 1 Pinto bean/
Wheast2 wheat germ3

Water .

ml .

. 3,560

3,400

Pinto beans

420

275

Wheast .

128

Torula yeast .

125

Wheat germ .

200

Casein .

100

Carrageenan

34

46

Ascorbic acid .

15

13

Vitamin mixture .

ml

12

12

Sorbic acid .

3

4

Methylp-hydroxybenzoate .

5

8

Formalin .

4

15

Tetracycline .

Aureomycin

mg

. 250

(chlortetracycline) .

mg

500

‘Unless otherwise noted, amounts are in grams (±1 g).

“From R. Patana (Vail et al. 1972). Container— paper bag (16 by
19.5 by 32 cm deep). Temperature— 26.7° C. Relative
humidity— 62%. Yield— average 130 pupae/container.

“Modified from W. W. Thomas (Vail et al. 1973) for use at the
Insect Attractants, Behavior, and Basic Biology Research
Laboratory. Container— paper cup (6.5 cm high by 11 cm
diameter). Temperature— 28°±1° C. Relative humidity—
65%±5%. Yield— average 60 pupae/container.

248

To rear six lepidopteran species, Patana (1969) used lima
beans and carrageenan as principal dietary ingredients
and transferred the medium into suitable containers with
a pressurized diet dispenser. Burton (1969) used a pinto
bean diet for the corn earworm, Heliothis zea (Boddie),
and dispensed it into containers with an automated
packaging machine. Either of these systems is suitable
for rearing cabbage loopers.

Larvae

Since cabbage looper larvae are not especially can¬
nibalistic, they may be reared in any vessel that will con¬
tain them and their diet, restrict microbial contamination,
and maintain a suitable environment. Larval micro¬
environments are provided by balancing factors such as
ambient climate, buffering capacity of the rearing con¬
tainers, dietary constituents (particularly available
moisture), and larval density and feeding behavior.
Assuming that adequate environmental conditions are
provided, selection of a suitable container depends on
local availability, cost, amount of required handling, and
space allocated for larval rearing. Vail et al. (1973)
evaluated relatively inexpensive 16- by 19.5- by 32-cm-
deep paraffin-coated paper bags by using different
amounts of seven diets, varying the number of eggs, and
incubating them in a constant environment. They found
these to be successful containers, especially with the pin¬
to bean/wheat diet shown in table 1. For paraffin-coated
paper cups, the most popular larval rearing containers,
Henneberry and Kishaba (1966) determined the best diet
quantities, insect densities, and rearing conditions.

Pupae

Pupae are usually extracted from the rearing container
by hand, and debris-laden silk is removed by washing
them in a 1%-1.5% sodium hypochlorite solution (about
20% commercial bleach) for 10-15 minutes (Henneberry
and Kishaba 1966). This procedure can be expedited if a
portable washing machine is used for agitation (a sodium
hypochlorite solution about 0.14%, is used for the
10-minute operation) and for automatic rinsing in water.
After the pupae dry, a sorting machine may be used to
position them for visual determination of sex (Wolf et al.
1972).

Oviposition and egg handling

Several kinds of oviposition cages have been tested, but
the cylindrical type made of hardware cloth and wrapped
with paper toweling remains the most useful (Ignoffo
1963). Henneberry and Kishaba (1966) tested one made of
0.64-cm-mesh hardware cloth that was 15 cm in diameter
and 28 cm high. They found the best conditions to be
24°-29° C and 50% or higher relative humidity, with

48-60 moths per cage. Moths were fed a carbohydrate
solution (1 g methyl p-hydroxybenzoate plus 50 g sucrose
plus 5 cc unprocessed honey plus 0.1 g of ascorbic acid
per 100 ml demineralized water) administered in cotton-
filled cups. Precise ranges of tolerance have been
established for the environmental factors that affect
mating and oviposition in these cages (see, for example,
Shorey 1963 and Henneberry and Kishaba 1967). Leppla
and Turner (1975) demonstrated that light quality, par¬
ticularly low intensity illumination at night, is important
for maximum fecundity, and Petterson (1974) reported
the use of fluorescent plant lights to increase egg
production.

In large operations, the hardware-cloth cages release
potentially hazardous insect scales into the insectaries.
Carlyle et al. (1974) solved this problem by incorporating
Plexiglas boxes with removable paper-toweling substrate
into a scale-filtration system. A moth-collecting system
(Ridgway and Whittam 1970) developed for the pink
bollworm, Pectinophora gossypiella (Saunders), could also
be adapted for use with the cabbage looper. But, in
moderate-sized colonies, the oviposition cage made of
hardware cloth is still the most convenient; scales may be
controlled by placing the cages above a plenum connected
to an industrial dust collector (Leppla et al., in press).

Selection of an oviposition substrate and surface-
sterilization procedure for cabbage looper eggs depends
on the overall treatment system and whether dry or wet
eggs are placed on diet. Paper toweling is the most
popular substrate; sometimes waxed paper and Teflon-
coated or cornstarch-coated paper have been used because
eggs can be removed more easily from these than from
uncoated paper (Hoffman et al. 1968). In any operation,
surface sterilization of the eggs with 10% Formalin
(45-minute exposure, 45-minute rinse in water) or with
0.3% sodium hypochlorite (5-minute exposure, rinse with
10% sodium thiosulfate, and wash in water for several
minutes) is recommended for elimination of viral and
fungal contamination (Vail et al. 1968). The method using
paper toweling and sodium hypochlorite has been incor¬
porated into a liquid system using automatic egg collec¬
tion and surface sterilization (Leppla et al. 1973).

Sanitation and internal security

Sanitation and internal security are basic to the main¬
tenance of mass-rearing facilities for cabbage loopers and
other Lepidoptera. A facility where large numbers of
moths are reared should include at least three levels of
quarantine (fig. 1). At the first level, adequate reception
and decontamination areas must be available to insure
that only pure materials are accepted into internal
storage. At this level, females are screened for pathogens
before their eggs are added to the colony. Only clean

249

RECEPTION

DECONTAMINATION

GO

LU

L->

<3.

Od

<C

cd

a

L->

o

CO

CO

<3;

t

STORAGE
Ingredients
Containers
Materials
ADULT COLONIES

EGG TREATMENT
DIET PREPARATION

EGG PLACEMENT

INCUBATION

HARVEST

MAINTENANCE

<3

LU

Qd

<C

CQ

Cd
I —
CO

Q

WASTE DISPOSAL

Figure 1.— Traffic pattern and levels of quaran¬
tine for a cabbage looper mass-rearing facility.
Arrows indicate direction of exchange across
thresholds and within the facility. The wider
the boundary, the more internal security is en¬
forced.

dietary ingredients, containers, and eggs are transferred
to third-level areas where the diet is prepared and eggs
are placed on it. Eggs and diet are sealed in containers
before leaving this maximum security to begin incubation
at the second level of quarantine. Pupal harvest and
maintenance (purchasing, pupal sexing, packaging, minor
equipment repair, etc.) also occur at the second level. An¬
cillary facilities may be used to house additional adults,
incubate larvae, or provide general work and storage
space at the first level of security (Leppla and Ashley
1978). Separate areas also at this level should be
designated for product distribution and waste disposal.

Access is usually limited to rearing personnel to prevent
contamination and maintain continuity in the work en¬
vironment. Also, individuals should be physically isolated
in specific work areas; or, if this is impossible, they may
begin each day in places that require low levels of con¬
tamination and progress to less critical locations. For
example, a person could prepare diet and place surface-
sterilized eggs on it before harvesting pupae and han¬
dling adults. But routine disinfection is needed for effec¬
tive isolation, and antimicrobials should be used when ap¬
propriate (Sikorowski 1975). Nutrient-agar plates may be
exposed to the ambient air regularly to determine the ef¬
fectiveness of this practice and to identify areas and pro¬
cedures that become sources of contamination. If un¬
cleanliness persists, the infested areas must eventually be
fumigated with steam, formaldehyde, methyl bromide,
etc., or disinfected and quarantined.

The degree of prophylaxis, routine disinfection, and
decontamination depends on characteristics of the rearing
facility. Absolute quarantine requires internal support
systems (food, water, restrooms, communication) and traf¬
fic restrictions (locks, double doors, regulated air
pressure, internal storage) that encumber efficient in¬
teraction among various parts of the facility. Too much
exposed surface area and use of porous building materials
precludes effective disinfection. Also, temporary facilities
must be available when areas are vacated for decon¬
tamination. These measures reduce efficiency and in¬
crease building expenses; so they should be used only
when absolutely necessary.

Current Research on Mass Rearing

Currently, researchers intent on improving mass rearing
of the cabbage looper and other Lepidoptera are con¬
cerned mainly with quality control. The behavior and
genetics of mass-reared insects must be appropriate to
their particular function. To make certain that the prod¬
uct insects meet the requirements, techniques must be
developed to routinely monitor vigor, competitiveness,
irritability, durability, and overall responsiveness. An
effective quality-control program depends on having
standards of comparison for each variable. For the cab¬
bage looper, most developmental standards have been
established (for example, see Toba et al. 1970 for a rating
system for larval diets and Smilowitz and Smith 1970 for
a scheme to measure the growth of each larval instar).
Setting behavioral standards is more complex because
the appropriateness of behavior depends on the product
insect’s intended use (Boiler 1972; Mackauer 1972;
Chambers 1975, 1977; Boiler and Chambers 1977; Webb
et al. 1981). So only partial behavioral standards have
been established (see Leppla et al. 1980 and Leppla and
Guy 1980).

Other areas where research is improving techniques for

250

mass rearing moths include efficient preparation and
dispensing of diet. For example, Harrell et al. (1973) and
Sparks and Harrell (1976) have adapted machines de¬
veloped for the food-processing industry. A diet that does
not require cooking (for example, see Spencer et al. 1975,
Leppla 1976, and Howell 1977) and a procedure for inter¬
mittent antimicrobial treatment (for example, see Chawla
et al. 1967) could facilitate this automation. Progressive
substitution and simplification of ingredients in the larval
diet have been made possible by sophisticated nutritional
studies (for example, see Chippendale and Beck 1968,
Kishaba et al. 1968, Poitout and Bues 1972, Terriere and
Grau 1972, Toba and Kishaba 1972, Dadd 1973, and
Vanderzant 1974).

Researchers are also learning to control cabbage looper
quality by controlling the rearing environment. For exam¬
ple, in an effort to increase rearing efficiency, Grau and
Terriere (1967) studied how larval rearing temperature
and dietary lipids influence the physical condition of
resulting cabbage looper adults. They also quantified
some of the complex interactions of environmental fac¬
tors that affect larval development (Grau and Terriere
1971). Others (for example, Gothilf and Beck 1967 and
Beck and Chippendale 1968) have studied how rearing en¬
vironment affects larval behavior. The results of these
studies have emphasized that the processes that make up
the rearing environment are interrelated rather than in¬
dependent.

Another area of current research is the automation of
rearing procedures. For example, in a preliminary study
to determine whether certain procedures in cabbage
looper rearing could be automated, Stimmann et al. (1972)
found that pupae could probably withstand mechanical
harvesting. Other promising techniques have been de¬
veloped for transferring moths without actually handling
them (for example, see Seay et al. 1971 and Wolf and
Stimmann 1971).

Development of new methods and materials to improve
rearing efficiency has significantly reduced production
costs. But implementation of these potential advance¬
ments depends on what can be done locally and practical¬
ly. Automation and other advanced techniques are useful
only if they complement functional rearing programs.

Conclusions

Large-scale cabbage looper rearing has been done at five
locations (U.S. Department of Agriculture laboratories at
Phoenix and Mesa, Ariz.; Columbia, Mo.; and Quincy and
Gainesville, Fla.). Each of these locations has produced
25,000-100,000 pupae/week for at least 6 months. Effi¬
ciency of these facilities has varied with the quality of
the facility and the availability of trained personnel. The

facilities with accurate, independent control of de¬
velopmental environments, for example, have been able to
prevent decomposition of the diet, promote uniform larval
development, and therefore achieve greater pupal yields.
Also, the facilities that filter airborne scales have been
able to reduce the effort needed to care for adult colonies.
So, in each of these pioneering rearing programs, 200-800
pupae have been produced per man-hour in 139-180 m2 of
space modified for all essential operations.

The need to provide a continuous supply of insects, as
has been done in these facilities, has brought about an
unusual synthesis between science and industry. Mass¬
rearing procedures are developed by teams of scientists
(rearing specialists, other entomologists, and agricultural
engineers) whose cooperation increases efficiency and
decreases costs. They develop the procedures through
understanding the growth processes of each insect stage,
studying rearing ecology (climate, containers, competi¬
tion, pathology, etc.), and investigating alternative
materials and sources. Industry contributes the new
materials and equipment specifically required for the new
procedures.

The best way to organize these procedures, materials, and
equipment into a rearing program is to conduct a sys¬
tems analysis and implement its recommendations. A
systems analysis is called for because mass rearing of in¬
sects is mainly an ecological discipline, one that must ac¬
count for a host of interrelated variables. A systems
analysis will consider all the primary variables that con¬
tribute to the efficient production of a quality product.
And it will suggest the best ways to make use of the
available procedures, materials, and equipment to reach
that end. Of course, the system can only be as good as its
available parts. Currently, then, since much work needs
to be done on quality control in cabbage looper rearing,
even the best programs can be depended on only for
quantity.

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253

Section 6

Management of Insect-Rearing Systems

The purpose of this last section is to draw together and
selectively expand the conference topics. In many re¬
spects, then, it recapitulates and unifies the overall
program. So this section has two purposes: to consider
important aspects of insect rearing that have not been
elaborated earlier in the conference and to discuss con¬
cepts of management at levels ranging from small proj¬
ects to massive international programs. Management con¬
cepts for insect-rearing programs have not, so far as I
know, been discussed in such detail, so the authors were
forced to think creatively and generate novel ideas. I
encouraged them to do this not because they could pro¬
duce perfect concepts, but because they are uniquely
qualified to ask pertinent questions. Their answers, and
your insightful modifications, should provide new
testable alternatives for managing our science and
technology.

Important aspects of insect rearing that have seldom
been treated as separate topics are quality control,
academic training, health and safety, and systems
analysis. Derrell Chambers, well known for his previous
contributions to quality-control theory, has collaborated
with Tom Ashley to develop practical means for monitor¬
ing insect-rearing products and processes. Marian Brooks
considers academic training from her perspective as a
university professor; since curriculums specifically
designed to prepare students for careers in insect rearing
do not exist, her challenge has been to point out short¬
comings in formal course work and to describe general
skills that could be developed for handling problems in
this field. Bob Wirtz volunteered to prepare a manuscript
on human health and safety based on his recent national
questionnaire on the subject; his findings, though based

on a pilot survey, indicate a need for concern about and
documentation of the human pathogens and allergens en¬
countered in insectaries. And Dave Akey, Bob Jones, and
Tom Walton have invented the first comprehensive
systems-analysis program for perpetual colonies. During
the last 5 years, they have developed and implemented
this information system by using it to manage colonies of
biting gnats and mosquitoes.

Discussion of management principles and concepts begins
with Chuck Schwalbe’s and Tim Forrester’s experiences
in developing and operating the Gypsy Moth Project at
Otis Air Force Base. Next, Bill Fisher probes the con¬
troversial question: What constitutes an insect-rearing
specialist? He deals with the existing but previously
undefined roles of centralized insectaries and their
managers. My paper, based on a 10-year association with
all levels of insect rearing, is specifically concerned with
population-suppression programs. Our anchorman, Pat
Patton, is one of today’s most enterprising young
managers. He currently oversees several massive insect-
control programs and is eminently qualified to describe
how important international politics can be to the
successful conduct of insect-control programs.

Without the unifying influence of effective program
management, current investments in insect production
and utilization may be in jeopardy, if not wasted. We
hope our efforts will encourage continued development of
the expertise and technology necessary to effectively
manage all sizes of insect-rearing programs and the con¬
trol programs that they support.

N. C. Leppla,

Research entomologist,
Agricultural Research Service

255

Putting the Control

in Quality Control in Insect Rearing

By D. L. Chambers and T. R. Ashley1

Concepts of Quality Control

Many studies have been done of concepts of quality con¬
trol, including Feigenbaum (1961), Juran et al. (1974), Ott
(1975), and American Society for Testing and Materials
(1976). Quality control is essential to survival in today’s
competitive market, and insect rearing demands the same
attention to quality control as any other industry. The
concepts of quality control are also the same; these are:

1. “Quality Control” is a management procedure that
develops, maintains, and improves quality.

2. The steps required in this process are setting stand¬
ards, appraising conformance with those standards,
acting on appraisal information, and planning how to
improve the product.

3. The elements of quality control— “product control,”
“process control,” and “production control”— are not
the same, but they are closely related. They all develop
information, have feedback loops, and control quality.
But product control rejects faulty products, and pro¬
duction control maintains consistency of production
output. Product control tells how well the product is
conforming to specifications and standards of quality.
It gives feedback so that a product’s departure from
established specifications can be corrected, or it
eliminates substandard products. Process control tells
how the manufacturing processes are performing. It
controls these processes so that deviations from the
product specifications will not occur as a result of
variation in the processes. Production control
regulates the consistency of production output, the
numbers of items produced, and the timeliness of their
production.

Industry’s recognition of these concepts proceeded slow¬
ly; so did other quality-control functions such as product
improvement and special process studies (acting on
quality-control data to identify causes of defects). Prog¬
ress was slowed by the reign of various enthusiasms,
such as “statistical quality control,” and of variations in

nomenclature, such as “quality assurance” and “system
effectiveness.” Struggles arose within corporations over
the role, size, and management of quality-control divi¬
sions. But quality control has come a long way from an
era characterized by product checking (“this one’s good
enough, this one isn’t”) to today's embracing of total
quality control. For this concept, most experts on quality
control have adopted tools and procedures to regulate the
processes of production so that product quality will be in¬
sured through control of processes.

Entomologists often concentrate too much on production
control while at best only controlling production proc¬
esses and products. We still see quality control as an
alarm and inspection system that oversees and intimi¬
dates production personnel. Instead, quality control
should be used by production personnel to measure and
control their own activities. This regulatory capability re¬
quires accurate information, which must be routinely
gathered. This information must be used not only in deci¬
sions on whether to accept or reject the product but also
in managing the processes that produce the products. In
this paper, we will emphasize process control.

Process Control

Existing industrial quality-control techniques may be
used for obtaining control over the processes used in in¬
sect production. These techniques will also help in the
derivation of reasonable, attainable specifications for the
products of those processes. Methods that estimate how
well processes work include direct measurement of
segments of a process (which requires setting aside a
production unit for the term of its measurement), deter¬
mining how well the product does what it is supposed to
do (which is most meaningful but least quantifiable),
and measuring how well the product conforms to its spec¬
ifications (which lacks predictability, requires firm spec¬
ifications, and does not provide process control). Two
techniques, process-capability studies and process-control
charts, would seem the best and most useful measures
of our control over insect-production processes.

Process analysis

‘Laboratory director and research entomologist, Insect Attract-
ants, Behavior, and Basic Biology Research Laboratory, Agri¬
cultural Research Service, U.S. Department of Agriculture,
Gainesville, Fla. 32604.

Process control requires that process analysis be applied
to ongoing insect-production systems. Assume that the
quality of performance by the insects being produced is
acceptable; do you know why? Can you quantify how
your production procedures result in acceptable quality?

256

WEIGHT MEDIANS (mg)

Figure 1.— Frequency distribution of the
weights of 50 cabbage looper pupae. Such in¬
formation can be used to identify normal varia¬
tion.

Can you predict whether your processes will continue to
produce quality insects? The first value of process analy¬
sis is its ability to identify causes of variability and to
predict future performance. The second value is its ability
to specify measurements of critical performance traits.

For example, a process might be a combination of people,
equipment, supplies, and activities that results in a batch
of pupae (one of a series of processes in an insect-rearing
operation). The capability of that process is its ability to
achieve measurable success in pupal variables (number,
weight, diameter, color, etc.). The process capability of
the associated operations is expressed in the measurable
reproducibility of these variables. Variability is to be ex¬
pected in, for example, the weights of pupae. But, if the
production process is under control (not drifting or re¬
sponding to uncontrolled influences), that variation will
be random (not assignable to specific causes) and con¬
tained within limits. Quality-control engineering has
found that a controlled process will result in
measurements that fall within ±3 standard deviations.
This measure of dispersion is similar to familiar probabili¬
ty statistics.

Pupal weight is a variable of product quality. Measuring
it tells us something about the acceptability of those
pupae. Measuring it can also tell us something about the
processes that produced those pupae. A continuing series
of measurements will tell us about trends in the quality
of the product and trends in the capability of the process
to continue producing at that level of quality. Knowing
what the quality is and has been is important, but know¬
ing what it will be, knowing how much control we have
over the production processes is important also. This is
the process capability.

Process analysis begins with an examination of the
variability of the process. A method used to examine the
capability of a process is called the process-capability
study. Its object is to define how much and in what way
product quality varies. It accepts that a controlled proc¬
ess has a random variability (not assignable to specific
causes) that should lie within ±3 standard deviations. A
process-capability study requires a large sample size— in¬
dustrial practice suggests a sample size of 50. A
frequency-distribution graph is made from this sample
(fig. 1). If the sample is typical and the frequency
distribution is normal, then process-control charting can
safely be used. Figure 1 is a frequency analysis of the
weights of 50 cabbage looper pupae and indicates the nor¬
mality of these weights.

The process-capability study based on an analysis of fre¬
quency distribution describes how much and in what way
the process varied at the time when the measurements
were made. But it is more important to examine how
much each production process varies sequentially. Fre¬
quency analysis is not well suited to such a sequential ex¬
amination because multiple large samples would be too
expensive. Also, frequency distributions are not easily in¬
terpreted, especially by technical staff or production
crews.

Process-control charts

Process-control charts are an alternative to frequency
analysis. They are chronological graphical comparisons of
measured product characteristics. They can be used to
detect and identify assignable causes of variation, and
they can be used to warn of drift toward unacceptable
quality due to nonrandom or assignable causes— they can
predict developing problems.

Process-control charts represent the differences between
means of small samples and the grand mean of a large
sample and the differences between ranges of small
samples and an average range (a range is the difference
between the maximum and minimum values in a sample).
A chart of means_displays the central tendency of the
average value of X or R from a sampling of n observa-

257

Figure 2.— Chart based on a hypothetical mean
or range, illustrating acceptable and unaccep¬
table trends in a measured variable.

tions of the measurements of a process; a chart of ranges
displays the spread of these measurements. The two
charts rely initially on the large sample drawn for the fre¬
quency analysis. The grand mean and the average range
are placed on the control charts as the expected central
tendency and spread. The limits of normal variation es¬
tablished above and below the central line are typically
±3 standard deviations of the mean or range. The control
charts are then used to plot sequential measurements of
means and ranges made on small samples (usually 5 or
10) at regular intervals over a specific time. These
measurements show trends in process controllability, and
exceptional deviations can then be related to production
activities or mistakes. When mean or range values exceed
control limits, the process is probably out of control— a
major problem exists on the production line. If variation
is random and contained well within the control limits,
control charting is needed to help maintain control. Even
if the charted points fall within the control limits, non-
random drift toward one of the limits (fig. 2) must be
carefully interpreted.

For example, analysis of the frequency distribution of
cabbage looper, Trichoplusia ni (Hiibner), pupal weights
(fig. 1) indicated random variation, so use of control
charts was appropriate. We divided the sample of 50
pupae taken for the process-capability study into ten
5-pupae subsamples and determined a mean and range
for each subsample. Then we calculated a grand mean
and its standard deviation and an average range and its
standard deviation. We used these values to plot the
horizontal lines on the mean and range charts (fig. 3).

Figure 3.— Mean and range charts for weights of cabbage looper
pupae for 30 days. Horizontal lines were determined from sam¬
ple of 50 pupae divided into 10 subsamples. Data points were
calculated from samples of 5 pupae collected on 11 successive
dates. SD = standard deviation; range difference between the
maximum and minimum values in a sample. UCL =upper con¬
trol limit. LCL=lower control limit (±350 in this example).

Next we took samples of pupae from the rearing opera¬
tions at regular intervals. Although industry normally
assumes sample sizes of 5 or 10 to be adequate, we
measured 25 pupae and analyzed their weights in sub¬
divisions of 5, 10, 15, 20, and 25 pupae to determine the
best sample size. The analysis verified that a sample size
of 5 or 10 is adequate. So we used only 5 of the 25
weights for each sample in plotting the points on the
mean and range charts.

There were no alarming trends in either means or ranges,
though there was one prominent deviation (sample No. 4);
for mean pupal weight, the production process remained
in control and contained well within ±2 standard devia¬
tions. The deviation in means was seen by the production
staff as they processed the pupae; deviations in range
were not. Later, we analyzed variables influencing this
process and attributed deviations in both mean and range
values to larval crowding. This use of information from
process analysis to solve production problems is an exam¬
ple of closing the feedback loop (see “The Closed-Loop
System in Quality Control for Insect Rearing,’’ by J. C.
Webb). This use of the feedback loop was stimulating to
the rearing staff and resulted in renewed analysis of the
production process.

When it is not possible to calculate upper and lower con¬
trol limits (±3 standard deviations), use tables available

258

i

Table 1. — Comparison of control limits derived from data with

those derived from a table1

Control limit
(standard deviation)

Value +3 —3

Mean

Data 314.4 192.6

Table 290.0 217.0

Range

Data 136.7 (2)

Table 133.6 (2)

'Calculations based on the pupal weights (mg) of 50 cabbage
loopers in subsamples of 5 each; tables from Bicking and Gryna
(1974).

2Three standard deviations below the average range value was a
negative number.

(Bicking and Gryna 1974) for determining these values
for both means and ranges. The control limits calculated
from our data and those determined from a table were
similar (table 1), so such tables will be fairly reliable.

Charts of means and ranges are normally displayed to¬
gether, the mean chart above the range chart. In this
way, production staff can easily follow the progress of
their processes over time. This ease of preparation and
analysis by technical staff is one of the greatest merits of
process-control charting.

Conclusion

We advocate immediate adoption of control charting for
each of your routine measurements within each of your
processes. Examples in a fly-production plant might in¬
clude fecundity, egg fertility, larval maturation rate, lar¬
val production rate, pupal size, percentage of eclosion,
adult behavior, adult physiological variables, sterility
assays, and release and recovery data.

Texts, handbooks, and manuals are available that de¬
scribe many modifications of control charting. Our exam¬
ple is representative but simplified, and aspects such as
sample size, frequency and variability must be examined.
Based on an evaluation of how sample size affects a con¬
trol chart’s accuracy, we believe a sample size of 5 or 10
would be adequate for examining the process of pupal
production, with weight as the measure, and days or
batches as the frequency. But we set control limits at ±3
standard deviations because of quality-control procedures
established by industry. We feel that these limits may
permit too much sample variation and suggest that ± 2

standard deviations might be more appropriate for pupal
weights. The degree of variability permitted about an
established average must be related to the expected per¬
formance of the reared insects.

We also believe that use of process-capability studies will
aid in the difficult task of establishing useful standards,
specifications, and tolerances. This task is especially
challenging in programs of insect production and release
because of the difficulty in adjusting unknown or partly
known performance requirements to restricted flexibility
in production processes. Standards and specifications are
invariably compromises between these requirements and
restrictions. Process analysis can provide data for making
these compromises and feedback for regulating them.

Our experience with process analysis has encouraged us
to adopt it broadly. We believe it offers information
largely new to rearing facilities, information that will lead
to new management capabilities and, therefore, to new
levels in the professionalism of our efforts.

Acknowledgments

We thank N. C. Leppla, U.S. Agricultural Research Serv¬
ice, for providing the pupal weights used for the control
charts.

Glossary

Assignable cause— an identifiable factor that contributes
to variations in quality.

Central tendency— the expected or average value(s) of X
or R from sampling units of n observations.

Control chart— a chronological graphical comparison of
measured product characteristics with limits reflecting
the ability to produce, derived from past experience.
Control limits— limits on the spread from the central
tendency (often ±3 standard deviations), computed
from the variations within sampling units, established
to indicate the presence of an assignable cause of qual¬
ity variation.

Control, types of

Quality control— integrative procedure that develops,
maintains, and improves quality; it is the regula¬
tory process through which quality of performance
is measured and compared with standards and
which takes action on the difference. It contains,
but is not limited to, components of total quality
control including product inspection, production
management, and the tools, technologies, and
assays of quality measurement.

Process control— regulation of the performance
of production processes so that deviations from
product tolerances and specifications do not
occur.

Product control— regulation of the conformity of

259

/

the product to specifications and standards of
quality.

Production control— regulation of the consistency,
reliability, and timeliness of production output.

Feedback loop— returning output information to the be¬
ginning of a process for correcting discrepancies
between intended and actual performance or for the
maintenance of current process standards and pro¬
cedures.

Process-capability study— analysis of process components
(people, equipment, supplies, activities) to determine
the inherent reproducibility of the process.

Quality— fitness for use.

Random variability— naturally occurring variation, not
attributable to assignable causes in quality variation.

Specifications— measurable units and subunits of design.

Standards— design references.

Tolerances— acceptable limits of variation in specifica¬
tions.

References

American Society for Testing and Materials.

1976. ASTM manual on presentation of data and con¬
trol chart analysis. 162 pp. The Society,
Philadelphia, Pa.

Bicking, C. A., and Gryna, F. M., Jr.

1974. Process control by statistical methods. In J. M.
Juran, F. M. Gryna, Jr., and R. S. Bingham, Jr.
(eds.), Quality Control Handbook, pp. 23-1—
23-35. McGraw-Hill Book Co., New York.

Boiler, E. F., and Chambers, D. L. (eds.).

1977. Quality control: an idea book for fruit fly
workers. Int. Organ. Biol. Control. West
Palearctic Reg. Sect. Bull. 1977/5, 162 pp.

Feigenbaum, A. V.

1961. Total quality control. 627 pp. McGraw-Hill
Book Co., New York.

Juran, J. M.; Gryna, F. M., Jr.; and Bingham, R. S., Jr.
(eds.).

1974. Quality control handbook. 1780 pp. McGraw-
Hill Book Co., New York.

Ott, E. R.

1975. Process quality control. 379 pp. McGraw-Hill
Book Co., New York.

260

Academic Training for Insect

By Marion A. Brooks1

Introduction

The skills, training, and experience needed by people in¬
volved in insect rearing vary according to the kind of pro¬
gram-colonizing a new species, maintaining a small col¬
ony, or mass producing insects for specific uses— and the
level of responsibility. This paper concentrates on what
academic training will be useful to those who work, or in¬
tend to work, in insect rearing.

Training for Colonizing
New Species

What must one know to colonize an insect species that
has never before been successfully reared? The most
basic requirement is, of course, to be an entomologist.

But the courses an entomology student might take may
not be pertinent. To be successful at colonizing new
species, the entomologist should have courses that in¬
volve close observations of insects in their natural hab¬
itats. Knowledge of insect behavior in the field will help
the entomologist know whether insects are behaving
normally in the laboratory. The entomologist should be
trained in the use of environmental controls so that he
understands how to regulate light, temperature, and
humidity. He should have a course in insect morphology
so that he knows, for example, how to determine growth
rates by measuring body parts. Likewise, he should have
a course in statistical design and analysis that will teach
him how to handle data such as growth rates. He should
have a course in insect pathology so that he can recog¬
nize when the laboratory insects are diseased. He should
have training in preliminary diagnostic procedures so
that he can recognize when prophylactic measures are
needed and what kind to use. Final identification of the
disease organism may have to be done by a diagnostic
laboratory after the colony has been rescued, so the ento¬
mologist needs training in how to collect and ship
specimen samples. To be able to detect nutritional defi¬
ciencies, the entomologist should have training in insect
behavior and nutrition.

In colonizing a new species, the entomologist must be
versatile. He must be something of an engineer, a mor¬
phologist, a biometrician, a nutritionist, a pathologist,
and a behaviorist. In actual practice, rearing a new

‘Professor, Department of Entomology, Fisheries, and Wildlife,
University of Minnesota, 1980 Folwell Ave., St. Paul, Minn.
55108.

Rearing

species may simply be a matter of consulting the litera¬
ture on the rearing of similar species and proceeding by
trial and error. But the student anticipating a career in
insect rearing should follow a curriculum designed to sup¬
port that career: animal nutrition and biochemistry; in¬
sect physiology, pathology, neurophysiology, behavior,
ecology, and genetics; and comparative endocrinology.
And the time to begin this course of study is not when
starting a rearing program but early, even in the under¬
graduate years.

Training

for Maintenance Programs

A second kind of insect rearing is the maintenance of
stock cultures for support of small research or teaching
programs. In such a case, a technician or student is
trained by a professional entomologist to follow pro¬
cedures and sequences already established by a pioneer¬
ing investigator and improved by various modifications.
Regular, routine care is specified, and the technical work
demands strict adherence to a schedule. The need for a
technician to have a broad background in entomology is
lessened by the necessity of following instructions. But
even here the operator needs to observe carefully to
detect any deviations from normal and report them
promptly to the supervisor.

Diseases caused by fungi and viruses are the greatest
threats in small programs. On-the-job training should suf¬
fice to show the technician how to avoid contaminating
the food with saprophytic molds or bacteria. The sources
of viral infections are usually unknown, but again a few
demonstrations of the appearance of insects diseased by
viruses should be adequate to make this problem recog¬
nizable. The worker needs to be shown how to keep alert
for the presence of mites, fungus gnats, small weevils,
etc., all of which are potential troublemakers in an insect
colony.

Although extensive formal academic training is un¬
necessary for maintenance work, an introductory under¬
graduate course in general microbiology would be helpful
in teaching the beginner the principles of pure culture
and the techniques of avoiding contamination. These
comments pertain primarily to rearing Lepidoptera.
Suitable modifications may be made for rearing aquatic
insects or less fastidious orders such as Diptera or
Blattaria.

261

Training

for Mass-Rearing Programs

Another kind of insect rearing is the mass-production
program that is the subject of many of the papers pre¬
sented in this conference. In these cases, the pioneering
work has been done, the technicians have been trained,
the engineering staff has been recruited, and the facilities
have been built. Usually a single commodity is to be pro¬
tected through the release of a parasitoid or of a pest
species that has been altered in some way. The program’s
emphasis is on teamwork, engineering, and efficiency. Ex¬
isting technologies are modified through statistical
analyses, sound economic practices, and improved data
collection. Improvements at these levels depend on the
entomologists having advanced training and experience
in insect physiology in its broadest sense.

For the mass-reared insects to be competitive when they
are released into the field, their sensory physiology and
behavior must not be radically changed. In some pro¬
grams, it is desirable to maintain genetic variability; in
others, quality control may depend on suppression of
variability. Training in genetic measurements is a definite
advantage for such work, but there is little evidence that
such courses are available in entomology departments.
While insect pathology courses are offered at many
schools, students who did not make early plans to work
in microbial control are rarely given any training in infec¬
tious diseases. It is obvious that academic course work
should be broadened to include genetics and pathology in
the training of any student who may work eventually in
programs involving biological control through release of
mass-reared insects and other organisms.

Academic Curriculums

for Insect-Rearing

and Biological-Control Programs

A tabulation of courses taught in entomology depart¬
ments at 51 institutions (Pieters 1979) suggests that
there is probably no formal offering of a course called in¬
sect rearing. Nor are there courses in nutrition, genetics,
or endocrinology. These subjects are probably parts of
courses entitled “physiology,” “biology,” or “behavior.”
Some of the listed courses useful in rearing insects are
general entomology, biology, morphology, physiology,
ecology, behavior, pathology, and biological control.

In anticipation that rearing programs will become more
important in the future with increasing use of biological
controls, entomology departments should encourage
students to prepare for this kind of work. The feeding
habits and nutritive requirements of many insects should
be studied. Some students might have enough certainty
about their future to be able to predict that emphasis on
a given taxon— mosquitoes, for example— would be in¬
structive. Some students might concentrate on a par¬
ticular commodity, such as com and its associated fauna.
Specializations such as these should be supported by a
broad program of insect physiology that would include
many of the courses mentioned above.

Acknowledgment

This is paper No. 11,243, Scientific Journal Series,
University of Minnesota Agricultural Experiment
Station.

Reference

Pieters, E. P.

1979. Survey of entomology courses currently being
taught in the United States. Bull. Entomol.

Soc. Am. 25: 284-285.

262

Health and Safety in Arthropod Rearing

By Robert A. Wirtz1

Introduction

The colonization and rearing of insects and other arthro¬
pods are prerequisites for many studies in entomology
and related fields. The mass rearing of arthropods is cur¬
rently playing a critical role in the development and im¬
plementation of pest-control programs (Smith 1966).

There are, however, potential health hazards associated
with occupational exposure to arthropods. Extensive
documentation exists on allergic responses to some ar¬
thropod products or bites (Frazier 1969, James and
Harwood 1969, Ebeling 1975), especially to stinging Hy-
menoptera (American Academy of Allergy, Insect Allergy
Committee 1965; Barnard 1973). Food inspectors, food
handlers, and other workers have also developed allergies
and dermatitis from exposure to material infested with
arthropods. Little has been written, however, about
allergies associated with arthropod rearing. Concern
about this lack of documentation resulted in the forma¬
tion in 1979 of an Insect Allergy Committee2 sponsored
by the Entomological Society of America. An initial goal
of this committee was to identify and document the ex¬
tent of health hazards associated with occupational ex¬
posure to arthropods.

Insect-Allergy Survey

In the fall of 1979, the Insect Allergy Committee mailed
questionnaires to 135 educational, government, and
private institutions in the United States. To survey in¬
stitutions actively involved in arthropod rearing, the com¬
mittee used the 1979 mailing list of the Insect Rearing
Group’s newsletter, Frass. Of those contacted, 82
responded; 48 (58.5%) of these reported at least one per¬
son with an allergy that coincided with occupational ex¬
posure to arthropods, their diets, or host animals; 32
(39.0%) reported that they had no work-related allergies;

2 (2.4%) reported having no occupational exposure.
Allergies were reported by 113 people, 18 to 79 years old.
The number of arthropod species encountered on the job

'Entomologist/CPT U.S. Army, Department of Entomology,
Walter Reed Army Institute of Research, Washington, D.C.
20012. The opinions and assertions contained herein are the
private views of the author and are not to be construed as of¬
ficial or as reflecting the views of the Department of the Army
or the Department of Defense.

2Committee members: W. A. Brindley; J. R. Gorham; A. M.
Hammond; J. D. Hoffman; M. H. Weiden; and R. A. Wirtz,
chairman.

ranged from 1 to 76 with the mode being 2 (28.8%). Peo¬
ple reporting allergies used some type of protective equip¬
ment routinely 25% of the time and as needed 60% of the
time. The protective equipment used included gloves
(35%), respirators/face masks (37%), head nets (15%), and
others (clothing, exhaust hoods, etc., 13%). The complete
results of the survey appeared in the Entomological
Society of America Bulletin (Wirtz 1980).

The sources of reported allergies were divided into nine
general categories (table 1). The Lepidoptera caused 69
(61.1%) of the reported allergies; most were reactions to
moth or butterfly scales. The gypsy moth, Lymantria
dispar (Linnaeus), was the most frequently implicated of
the Lepidoptera, 32; then tobacco homworm, Manduca
sexta (Linnaeus), 8; pink bollworm, Pectinophora
gossypiella (Saunders), 4; tobacco budworm, Heliothis
virescens (Fabricius), 3; silkworm, Bombyx mori (Lin¬
naeus), 3; greater wax moth, Galleria mellonella (Lin¬
naeus), 3; unspecified moths, 4; and 1 or 2 each of 7 other
species, 12. Of the 13 Coleoptera that caused allergies, 5
were weevils; 9 of the 11 Orthoptera causing allergies
were cockroaches. Honey bees, Apis mellifera Linnaeus,
were reported in 9 of the 12 cases for Hymenoptera. Mos¬
quitoes caused five of nine Diptera allergies; most were
reactions to adult scales. Eight of the Acarina allergies
were attributed to mites and one to ticks. The dry ingre¬
dients in artificial rearing media caused most of the

Table 1. — Sources of allergies reported on questionnaires of the
Insect Allergy Committee

Response1

Source Number Percent

Lepidoptera . 69 61.1

Coleoptera . 13 11.5

Hymenoptera . 12 10.6

Orthoptera . 11 9.7

Acarina . 9 8.0

Diptera . 9 8.0

Diets 6 5.3

Host animals . 6 5.3

Other2 . 13 _ 11.5

Total . 146 .

‘Multiple responses from 113 people.

2One or more responses to: Aranea, Hemiptera, Isoptera, Neurop-
tera, silk, tissues, or hemolymph.

263

allergies related to diet. All allergic reactions to host
animals were caused by rodents or rabbits. More than
68% of the people suffering allergies were exposed to the
allergen daily; 12.4% were exposed weekly. Although
eight people reported observing symptoms on first con¬
tact, and one person reported having had 20 years of ex¬
posure before the allergy developed, most estimates
ranged from 1 week to 7 years with a mode of 12 months
(20.8%). Most (77.0%) allergens came from airborne
material (table 2). Sneezing and a runny nose were the
commonest (68.1%) allergic reactions (table 3).

Of those persons reporting arthropod allergies, 28%
stated that they had had to stop work or transfer; 49%
had consulted a physician; and 47% said that the allergy
had required some type of medication or medical treat¬
ment; 24% reported having used self-treatment (over-the-
counter antihistamines); 25% were using prescription
drugs; and 4% were receiving desensitization
inoculations.

Table 2. — Suspected origins of allergic responses reported on
questionnaires of the Insect Allergy Committee

Suspected origin

Response'

Number

Percent

Airborne material .

87

77.0

Contact .

72

63.7

Sting .

8

7.1

Bite .

4

3.5

No response .

9

8.0

Total .

180

'Multiple responses from 113 people.

Table 3. — Type of allergic reaction reported on questionnaires of
the Insect Allergy Committee

Response'

Symptoms

Number

Percent

Sneezing and runny nose .

77

68.1

Skin irritation .

70

61.9

Eye irritation .

70

61.9

Breathing difficulty .

38

33.6

Anaphylactic shock .

3

2.7

Other2 .

4

3.5

Total .

262

‘Multiple responses from 113 people.
2Headache, nausea, faintness, fever.

Major Health Problems
Allergies and dermatitis

The insect-allergy survey showed that occupational
health hazards associated with arthropod rearing usually
affect the respiratory system, the skin, or the eyes.
Rhinitis and bronchial asthma have resulted from ex¬
posure to silkworms (Inasawa et al. 1973, Tadani et al.
1974, Saakadze 1976, Fuchs 1979) and other moths
(Stevenson and Mathews 1967; Smith 1973, pp. 405-407);
and other occupational respiratory allergies have been
caused by bees (Yunginger et al. 1978); flies (Ordman
1946, Ebeling 1975); cockroaches (Bernton and Brown
1964); beetles (Smith 1973, pp. 413-416; Ebeling 1975);
locusts (Frankland 1953, Fuchs 1979); screwworms,
Cochliomyia hominivorax (Coquerel) (Gibbons et al. 1965,
Herrman 1966); mites (Wharton 1976, Fuchs 1979); mos¬
quitoes, owlflies, termites, and the introduced pine saw-
fly, Diprion similis (Hartig) (Wirtz 1980). Airborne
cuticular fragments, hairs, scales, and dried tissues and
feces are abundant in most rearing areas. Some people
can be extremely allergic to insect fragments (Perlman
1958, Marchand 1966, Fuchs 1979), and these allergens
can sensitize through inhalation or contact (Bernton and
Brown 1969, 1970). Two laboratories rearing Lepidoptera
reported that 53% of their personnel developed allergies
to moth scales even though exhaust hoods, protective
masks, and laboratory coats were used routinely (Wirtz
1980). Sensitivity to the German cockroach, Blattella ger-
manica (Linnaeus), has been correlated with the intensity
and duration of exposure (Bernton and Brown 1967a). In¬
sectary personnel, museum curators, food inspectors or
handlers, and entomologists who have repeated close con¬
tact with arthropods are especially susceptible to
allergies of arthropod origin. An absence of allergic symp¬
toms does not mean that arthropods or their products are
not causing sensitizations. Bernton and Brown (1967b)
reported that when “nonallergic” people had skin tests
with extracts from seven commonly reared insect pests of
human food, 25% of the tests were positive.

Dermatitis, a well-documented allergic response to arthro¬
pods (Barnard 1966, James and Harwood 1969, Ebeling
1975), was reported by 63.7% of the people responding to
the insect-allergy survey. Implicated sources include
beetles (Scott 1962), moths and butterflies (Smith 1973,
pp. 405-407; Kwangtung Occupational Diseases Preven¬
tion and Treatment Hospital and Chung Shan Medical
College 1974; Perlman et al. 1976; Press et al. 1977;
Guseinov 1977), cockroaches (Bernton and Brown 1964),
locusts (Wirtz 1980), theraphosid spiders (Ratcliffe 1977),
and mites (Alexander 1972, TerBush 1972, Krantz 1978).
The twospotted spider mite, Tetranychus urticae (Koch),
was the source of four of the nine allergies caused by
Acarina. One respondent stated that everyone who

264

reared this mite in his laboratory had developed a contact
allergy (Wirtz 1980). More than 20 dermestid species that
damage insect collections have been recorded (Ebeling
1975), and a single larva can release over 3,000 urticating
hastisetae (Okumura 1967, Mills and Partida 1976), which
are capable of penetrating the skin (Smith 1973, pp.
413-416). Unlike the hollow stinging hairs of many
Lepidoptera larvae, which may contain toxins, irritations
from hastisetae appear to be solely mechanical.

Hastisetae also cause eye irritation, either directly enter¬
ing the eye or indirectly moving from the arthropod to
the worker’s hands and then to the eyes (Ebeling 1975,
Mills and Partida 1976).

Envenomization

Since many arthropods can inject venom or salivary
secretions by means of a stinger, mouthparts, or ur¬
ticating hairs, they may be a serious health hazard to
people rearing or collecting them (American Academy of
Allergy, Insect Allergy Committee 1965; James and Har¬
wood 1969; Barnard 1973). The stinging arthropods in¬
clude Hymenoptera (bees, wasps, hornets, ants, velvet
ants) and Scorpiones (scorpions). In the United States,
Solenopsis spp. and Pogonomyrex spp. ants, and the Cen-
truroides spp. scorpions pose the most serious problem to
“nonallergic” people (Biery 1977, Frazier 1969). Allergic
reactions to honey bee stings affect many beekeepers and
their families (Yunginger et al. 1978, Lichtenstein et al.
1979). Honey bees and wasps are the most common
causes of anaphylaxis (Barnard 1973, Gottlieb 1979).

Allergic reactions to any of the stinging arthropods can
result in death. Every year, stings actually kill at least
50-100 persons in the United States (Lichtenstein 1977,
U.S. Food and Drug Administration 1979, Gottlieb 1979).
While some people become increasingly sensitive to
stings, many have no forewarning that they might have
anaphylactic reactions (Barnard 1973). Fortunately, only
11 of the 113 respondents to the insect-allergy survey
listed allergies to stinging Hymenoptera: 9 to honey bees
and 2 to the fire ants, Solenopis spp. (Wirtz 1980).

The biting or piercing arthropods include the Thysanop-
tera (thrips), Anoplura (sucking lice), Heteroptera (true
bugs and allies), Diptera (mosquitoes and biting flies),
Siphonaptera (fleas), Acarina (mites and ticks), Aranea
(spiders), and Chilopoda (centipedes). All of these, be¬
cause they can inject salivary secretions or venoms
through specialized feeding appendages, are a potential
health hazard to insectary personnel (Frazier 1969, James
and Harwood 1969, Ryckman 1979, Ryckman and Bentley
1979). As with stings, allergic responses cause most
of the severe reactions to bites (Derbes 1971). Reactions
vary greatly. Some people suffer no visible effects, not
even swelling in the area of multiple bites, while others

react violently to a single bite (Scott 1966, James and
Harwood 1969).

The larvae of more than 50 species of Lepidoptera can
envenomize through urticating hairs. These specialized
setae are usually hollow and connected to a gland that
releases a toxin. The intensity of irritation varies with
the species, sensitivity of the victim, and the exposed
tissue (Smith 1973, pp. 405-407; Scott 1966). If ur¬
ticating hairs get into the eye, they may cause blindness
(James and Harwood 1969).

Chemical secretions

Arthropods are noted for the diversity of defensive
chemicals they produce and their unique delivery systems
(Roth and Eisner 1962, Eisner and Meinwald 1966).
Defensive secretions are usually a mixture of these reac¬
tive, irritating, or toxic compounds: carboxylic acids,
alcohols, aldehydes, ketones, esters, lactones, phenols,

1.4- quinones, hydrocarbons, steroids, and other
miscellaneous compounds— for example hydrocyanic acid
(Blum 1978). Many of these defensive exudates have con¬
stituents that significantly increase their effectiveness by
facilitating spreading or penetration. Delivery systems
range from localized exudation and reflexive bleeding to
remarkably accurate defensive sprays (Eisner and Mein¬
wald 1966). People rearing, collecting, or accidentally
exposing themselves to these defensive chemicals may
encounter risks ranging from temporary skin irritation to
chronic exposure to suspected carcinogens. For example,
certain tenebrionid beetles release a mixture of

1.4- benzoquinones when stressed. Even when few of these
beetles are present, they can be detected by the quinone
odor they give to an infested commodity. Their quinones
are highly reactive compounds with suspected mutagenic
and carcinogenic activity (Ladisch 1965, U.S. En¬
vironmental Protection Agency 1979). At least six other
species of stored-products arthropods can also be
detected by their odor, but the composition of these ex¬
udates is unknown (Wirtz, in press).

Parasites and pathogens

Many commonly reared or collected arthropods can
parasitize humans (arthropodiasis) or be vectors of
pathogenic organisms. Arthropodiasis may appear to be
an unlikely health hazard under normal rearing condi¬
tions, but it can easily happen if sound laboratory prac¬
tices are not strictly enforced. Over 100 species of fly
larvae have been identified in cases of myiasis in man by
James (1947), James and Harwood (1969), and Palmer
(1970). Other cases have involved accidental enteric
myiasis, canthariasis (beetles), scoleciasis (moths) and
acariasis (mites) (James and Harwood 1969, Ebeling
1975). In one instance, a specimen collector got occupa-

265

tional “myiasis” by using an aspirator during insect col¬
lection; about 2 months after he had finished the summer
collecting during which he used the aspirator 4-6 hours
per day, 69 specimens from 4 insect orders were passed
from the antrum of his sinus; apparently, the insects
entered the sinus as eggs that had passed through the
brass screen on the aspirator (Hurd 1954).

Arthropods are biological and mechanical vectors and in¬
termediate hosts of human disease-producing organisms
(for example, see James and Harwood 1969 and Smith
1973). Also, many human pathogens grow on synthetic
rearing diets and other organic matter present in many
insectaries. P. P. Sikorowski (see “Microbial Contamina¬
tion in Insectaries. Occurrence, Prevention, and Control”),
analyzing the problem of employee safety and microbial
contamination in the insectary, concluded that humans
are a major source of these pathogens. Filth flies and
domestic cockroaches are often implicated as vectors of
pathogenic organisms (James and Harwood 1969, Palmer
1970). Evidence for fly involvement is present for more
than 60 human diseases (Greenberg 1973). At least 4
strains of poliomyelitis virus, 40 pathogenic bacteria, the
eggs of 7 species of pathogenic helminths (Roth and
Willis 1957, 1960; Frishman and Alcamo 1977), and
several species of fungi and protozoa (Beatson 1976) have
been isolated from wild cockroaches. Laboratory colonies
established from wild populations should be checked
carefully for human pathogens.

Generally, micro-organisms pathogenic to insects and
used as active ingredients in microbial-control products
will not harm man, other animals, or plants (Steinhaus
1957). Certain arthropod viruses, the baculoviruses, are
surprisingly host specific and appear to be restricted to
invertebrate hosts (Tinsley 1979). In 1973, the U.S. En¬
vironmental Protection Agency exempted arthropod virus
from tolerance-residue requirements stating that it
presented no known hazard to human health (Falcon
1976). But the desired viral agent must be carefully
isolated and purified and then positively identified
(Tinsley 1979). Bacteria pathogenic to insects are also be¬
ing mass-produced after extensive field testing with no
recognized hazards to human health or the environment
(David 1975, Arata et al. 1978). Even though micro¬
organisms already tested seem to be harmless to humans
and the environment, potential health hazards must be
considered when new biological control agents are being
developed and tested (Tinsley 1979).

Conclusions

Limiting contact with causative agents will effectively
reduce health hazards associated with exposure to ar¬
thropods. Decreasing exposure to allergens, hazardous
chemical and physical products, and pathogenic micro¬

organisms is best accomplished by design or modification
of laboratories, enforcing general sanitation requirements,
and using protective equipment. If prophylaxis is im¬
possible, people who continue to work with arthropods
that cause them serious allergies should consider using a
sting kit and getting desensitization inoculations
(Lichtenstein et al. 1979). Because any strong allergic
reaction is potentially serious, the sufferer should consult
an allergist.

Identification and documentation of specific occupational
hazards associated with exposure to arthropods should
continue. This is the goal of the Insect Allergy Commit¬
tee. An expanded survey has been distributed; the results
will be used to inform insectary personnel, managers, and
administrators what the hazards are. But the lack of
specific allergenic fractions prevents the confirmation of
work-related symptoms and the preparation of specific
desensitization inoculations. Information from the ex¬
panded survey and from reliable research data will enable
us to address the physical, ethical, and legal problems
associated with human exposure to potentially hazardous
arthropods and their products.

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268

Systems Analysis and Automated
Data Processing In Insect Rearing

A System for the Biting Gnat Culicoides
variipennis and Mosquitoes

By David H. Akey, Robert H. Jones, and Thomas E. Walton1

Introduction

Quality-control tests are being used increasingly to de¬
termine the fitness of insects produced for laboratory re¬
search or in mass-rearing programs (Chambers 1977).

Daily records must be kept of environmental variables,
their parameters, and changes in nutritional regimens;
other pertinent data concerning rearing and quality con¬
trol must be recorded for each generation or production
run. The recordkeeping and analyses needed to process
the data are an added burden to the task of rearing in¬
sects. To alleviate this problem, we enlisted the help of
personnel in the U.S. Department of Agriculture, Science
and Education Administration, Administrative Manage¬
ment, Communications Data Services Division (CDSD)
and explored how systems analysis can be used for
gathering, analyzing, and retrieving data associated with
the production of insects.

Systems analysis has been defined as “the approach to a
system that is the opposite of trial and error. All in¬
fluences and constraints are identified and evaluated in
terms of their impact on the various parts of the system’’
(FitzGerald and FitzGerald 1973). Put another way, sys¬
tems analysis is “the examination of an activity, pro¬
cedure, method, technique, or a business to determine
what must be accomplished and the best method of ac¬
complishing the necessary operations” (Sippl and Sippl
1972). The increasing sophistication of computers in the
last 20 years has contributed to the increasing sophistica¬
tion of systems analysis. In fact, it is frequently viewed
as part of a broader concept of a management informa¬
tion system that provides information used by manage¬
ment for decisionmaking (Lucas 1974, Youssef 1975).
Personnel responsible for rearing insects should view
systems analysis as a way to organize insect-rearing
operations in efficient and logical sequences. Such
organization facilitates timely analysis, makes data
retrieval possible, and aids decisionmaking. Computers
are tools used to achieve these goals and should be

'Research entomologist, supervisory research entomologist, and
research/location leader, U.S. Department of Agriculture,
Agricultural Research Service, Arthropod-Borne Animal
Diseases Research Laboratory, Denver, Colo. 80225.

viewed as “devices that can be harnessed to serve informa¬
tion needs and improve organization efficiency” (Alexander
1974). For more details about systems analysis and
related technology, see Bingham and Davies (1972) or
Gore and Stubbe (1975).

Insect rearing can be viewed as a business that can
benefit directly from the use of systems analysis and of
automated data processing (ADP) with computers. As an
example of how such an operation works, we present the
methods used to apply systems analysis to the produc¬
tion of colonies of the biting gnat Culicoides variipennis
(Coquillett)— the primary vector of bluetongue virus in
the United States— and of several colonies of mosquitoes.
Although the procedures detailed here were designed for
these Diptera, they can probably be used readily with
most insect colonies. To our knowledge, this is the first
time that systems analysis and ADP have been applied
to insect rearing.

Methods and Materials

We began the application of systems analysis to insect
rearing by having the rearing operation reviewed by
personnel from the CDSD to determine if the proposed
project was feasible. A systems analyst and a computer
programer then worked with us to determine what
portion of the rearing operation would be easiest to incor¬
porate into an information system and to select com¬
ponents of the rearing operation for inclusion in the
system. We decided to limit the first information system
(the one described in this paper) to data that could be
recorded on a single 80-column coding form. To simplify
this presentation, all examples refer only to C. variipennis
unless otherwise stated.

C. variipennis gnats were reared with the methods of
Jones et al. (1969), and their size was monitored by dry
weight and wing length (Akey et al. 1978). Techniques for
rearing mosquitoes generally followed those described by
Gerberg (1970). The larvae were reared in 38- by 23- by
6-cm white enamel photographic developing trays and fed
powdered rabbit food and/or a high-protein chow (Purina
Special Steer 32, Ralston Purina Co., St. Louis, Mo.) or
feed supplement (Jones et al. 1969). Adults were held in
30.5-cm3 cages.

269

All data were recorded manually and then keypunched on
80-column cards with IBM (International Business Ma¬
chines Corp., White Plains, N.Y.) machines. An IBM
370/168 computer, associated peripherals, and operating
systems at the U.S. Department of Agriculture, Wash¬
ington Computer Center, Office of Automated Data Sys¬
tems, were used for processing and storing data. A
Teletype (Teletype Corp., Skokie, Ill.) model 43 teleprinter
with a Com Data (Com Data Corp. Skokie, Ill.) model
302F2-33 modem (a device that changes computer-
compatible digital signals to telephone-compatible analog
signals and vice versa) was used as a terminal for time-
share operation (TSO). It was operated at half-duplex
(unidirectional communication over a line) over voice-
grade telephone line at 300 bauds. The computer was
operated with both standard software (collections of com¬
puter programs) and with programs written specifically
for this project. Before data on quality control were
keypunched, preliminary analyses were performed with
an HP9830 (Hewlett-Packard Calculator Products Divi¬
sion, Loveland, Colo.) calculator system programed with
either the 98930A Stat Pac 1 or the General Statistics
Pac. All other analyses were performed with the IBM
370/168 programed with the Statistical Analysis System
(SAS, SAS Institute, Raleigh, N.C.; At this time, SAS
can be run only on IBM mainframe computers).

Entry of Data

Flow charts

and source and record documents

The first step in devising a system for data entry was to
review the daily rearing operations that had been listed
and ordered in logical sequences on four check sheets;2
one of these is shown in fig. 1. In the second step, we
reviewed the data-collection forms used with the check
sheets (for example, fig. 2). Next, data recorded merely as
a convenience were separated from essential data. Also,
data that were not permanently recorded were
reevaluated to determine if they were needed. All essen¬
tial data were divided into groups of related fields (par¬
ticular categories of data treated as a set or group). The
fields were arranged in logical sequences for data collec¬
tion and recording.

In the third step, we used a specially designed record
document to list the fields. Headings were typed on a
general purpose 80-column coding sheet, 8 Vi by 14 inches,
that was revised several times from its conception in
1974 until a final document was approved in 1977. After
2 years of use, a more convenient form was printed on

2In computer terminology, these are known as flow charts in the
form of check lists.

8V2- by 11-inch paper (fig. 3). The fields on this document
are described in the procedure on coding data (appendix
A). Quality-control data on insect size were recorded on
two other record documents, one for pupal production
and one for wing length and dry weight.

Data coding and editing document

The fourth step was the development of a data-coding
system that was simple but comprehensive enough to
cover the variations that occurred in our rearing opera¬
tions. Additionally, we needed an editing document that
gave the ranges, limits, and code types (alphabetic,
numeric, or alphanumeric) for each field that was coded
(appendix A). The combined data-coding and editing
document was used as a guide for proper coding and
recording of data about insect rearing. It defined the per¬
missible relationships between codes for different fields
and was operated as a program in the computer so that
data entered were tested against the editing criteria.

Data that did not meet the criteria were listed in the
printout of the editing program and corrected if wrong;
data that did not meet the editing criteria but were cor¬
rect were left unchanged. Examples of the codes and per¬
missible values were posted for use by the technicians
who took care of our insect colonies. Some coded data are
shown in fig. 3. Data collected on the record document
(fig. 3) were keypunched on 80-column cards, entered in
the computer, and stored in data files (collections of
related records) on magnetic tape. Quality-control data
were reduced to mean, standard error, etc., with a
calculator before entry into the computer. The computer
program prepared by the systems analyst to perform the
editing listed in appendix A is shown in appendix B.

Retrieval of Data

Retrieval-request document

To produce reports on insect rearing, we first prepared a
retrieval-request document (appendix C) for the systems
analyst. This document was used as a guide for prepara¬
tion of the required software. It listed the items of data
input necessary for the software to generate the re¬
quested report. Also, the document listed examples of
hypothetical output.

Retrieval request reports via TSO terminal

The retrieval programs were written to make use of SAS,
a versatile and easy-to-use software computer package.
Retrieval requests were made from a terminal. The
desired SAS retrieval program was operated in an edit
mode that allowed input or changes to be made in the
program from the terminal; fig. 4 is an example of a

( Continued on page 279.)

270

DAILY MASTER CHECK SHEET
C. varlipennis

SHEET NO. 3 8
DATE 7-53 2%

FORM 005

DATE

Y

X

ri 0

V

as

V

XV

r

V

of'

rp

V

ao

Y

V

$

as

V

a

INITIALS

for daily work

ap

JSJ.

th

E-P

E.P.

ER

ER.

J.H.

JH

sp

EP

ER

E.R

Time Start

S

/

<f*°°

/

0*°°

/

/

/

/

/

/

X

Record pan

temperatures

X

X

X

X

X

X

X

X

X

X

X

X

X

X

Light timers-

lights on

X

X

X

X

X

X

X

X

X

X

X

X

X

X

timers in use¬

setting correct

X

X

X

X

X

X

X

X

X

X

X

X

X

X

Pupae picked-

every other day

y

—

X

-

X

—

X

X

—

X

—

X

-

X

Larval Pans Setup*

Mon.Wed.Fri

3

-

3

-

3

-

-

3

-

3

-

3

-

—

Holding Pan Setup

Mon , Wed , Sat

/

-

/

-

-

/

-

/

-

/

-

-

/

-

Larval Pans Dumped

Mon ,Wed , Fr i

3

-

3

-

3

-

-

3

-

3

-

3

-

-

Check Pan Water
level - Daily

X

X

X

X

X

X

X

X

X

X

X

x

X

X

Correct filming

Daily

X

X

X

X

X

X

X

X

X

X

X

X

X

X

Determine food
requirements

X

X

X

X

X

X

X

X

X

X

X

X

X

X

Add food to larval pans;
bacterial starter

X

-

X

-

X

X

-

X

-

X

-

y

X

-

NB

X

X

X

X

X

X

X

X

X

X

X

X

X

X

dry;Alf, K, J*

K

J

K

J

X

JX

X

JX

J

K

X

K

K

J

Larval pan record
sheet

X

X

X

X

X

X

X

X

X

X

X

X

X

X

Adult setup record
sheet

X.

X

X

X

X

X

X

X

X

X

X

X

X

X

Adult holding record
sheet

X

X

X

X

X

X

X

X

X

X

X

X

X

X

Time out

vxP°

&

&

**

&

X”

/

/

Water circulators on

A and B

X

X

X

X

X

X

X

X

X

X

X

X

3

X

Check reservoirs
pump on

X

X

X

X

X

X

X

X

X

X

X

X

X

X

Pump running

X

X

X

X

X

X

X

X

X

X

X

X

X

X

Water level

X

X

X

X

X

X

X

X

X

X

X

X

X

X

c m j Mon, Wed,

Eggs Placed - ’ ’

Fri

X

X

-

X

-

-

X

-

X

-

X

-

-

Refrig. Temp.

(»o

(o-O

56

(,o

<X3

£.3

£ 3

<pZ

OS

6,0

J>S

(jP £

& O

*Letter or number

Figure 1.— Original check sheet to record insect rearing opera¬
tions used for the primary Culicoides variipennis colony. NB is
nutrient broth food; Alf, K, J are dry powdered foods— see
Jones et al. (1969).

SHEET NO. /£

ADULT HOLDING RECORD DATE 1+' 232

C_. varllpennls FORM _ 003

Colony OOO-Ah

DATE

2 W

227

2?o

27/

272

2^3

27V

27X

27£

Egg estimates*

Tu(C) ;F(A) ;Sun(B)

c

ooo

—

—

A

H 0,0 00

—

e

3otooo

—

C

so, ooo

C

soo

Dead flies removed
-Daily-

X

X

X

X

X

X

X

X

X

New cage set up*

M(A);W(B);F (C)

—

S

—

C

—

—

A

—

B

By use of flies from*

—

/029-

1093

—

/099-
109 i>

—

—

1099-
/ / o/

—

/ /oz-

iios

Blood fed M(A) ;W(B) ;F(C)
Rabbit no. / *

—

3

-

c

—

—

A

—

3

Change: 2 sugars and

1 water-Daily*

X

X

X

X

X

X

X

X

X

Place egg dish*
M(C);Th(A);Sat(B)

c

—

A

—

B

—

£

c

—

Notes**

DATE

2n

272

3 oo

30/

302

303

30H

30S-

Egg estimates*
Tu(C);F(A);Sun(B)

—

A

90, ooo

—

3

6>0,000

3

/poo

C

sopoo

C

/ 0,000

—

2\

30,000

Dead flies removed
-Daily-

X

X

X

X

X

X

X

X

X

New cage set up*

M(A) ;W(B) ;F(C)

—

C

—

—

A

—

s

-

c

By use of flies from*

—

//C#f-

///£•

-

-

/// 7-
II ZO

—

nu¬
ll 25

-

U2X,-

!'33

Blood fed M(A) ;W(B) ;F(C)
Rabbit no. *

-

c

-

—

A

—

s

-

c

Change: 2 sugars and

1 water-Daily*

X

X

X

X

X

X

X

X

X

Place egg dish*

M(C) ;Th(A);Sat(B)

A

-

A3

3

c

c

—

A

-

Notes**

Blood Feeding Host _ *Use letters A,B,or C, or "All"

Temperature & Relative Humidity_ _ _°C/ _ %RH **Indicate under "Notes"Mainten-

Pho toper iod : Light _ _( hr) ance of old cage.

Figure 2.— Original data-collection form for
recording holding and maintenance data for
adult flies for the primary Culicoides variipen-
nis colony. BF is blood feeding of flies from
cages (4-digit numbers in left column) that
were combined into cages A, B, or C used to
propagate the colony.

272

NOTE: * Day = Julian Day last 2 digits. COLONIES - ADULT PRODUCTION

Shaded numbers indicate card columns data begins in. (Entomology)

3dAl QUVD -

o

—

- 1

ADULT

DISPOSITION

| Colony

| Davl

2

r'-

&

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0050

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<£>

In

—

LOCATION

Physical Parameters

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In

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ADULT NUTRITION

3rd Feeding 1

adA_L poo-j csj

Time

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83

83

EMERGENCE TIME

END
Pupae
Removed
from Cage

T ime

Lj

in

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o

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START
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Placed
in Cage

Time

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RECORD IDENTIFIER

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uouejauau cm

c

o

3

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Small

Cage

No.

3

Large

Cage

No.

4

m

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on

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-

-

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-

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-

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-

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-

-

-

-

-

-

273

Figure 3.— Colonies-adult-production record document for recording data used in the retrieval
system. The 6 lines of coded data are shown as examples. The coding system is given in appendix A.

Figure 4.—SAS pupal production program. Ar¬
rows (not part of program) indicate data inputs
that were allowed to stay as presented or
changed to conform with needs of user.

list

COMMENT PROGRAM m ;

• CHOOSE THE TYPE OF TIME UNIT ON RECORD;

♦ FOLLOUING THE MACRO MAC1 STATEMENT;
♦CHOOSE ONE: UK MON QTR YR;

MACRO MAC 1

•UK

♦ CHOOSE TYPE OF CUMULATION FOLLOUING THE MACRO MAC 1 A STATEMENT;

» CHOOSE ONE: CUM. UK CUM.MON CUM QTR CUM YR;

MACRO MAC 1 A
» CUM UK
7.

♦CHOOSE TIME UNIT OF REPORT FOLLOUING THE MACRO MAC 1 B STATEMENT;

♦ CHOOSE ONE: UEEK MONTH QUARTER YEAR;

MACRO MAC 1 B UEEKZ

• CHOOSE COUNTING METHOD(S);

♦ FORMAT MAT RE MULTIPLE (METHOD , METH- D OR METH= E ETC.);

• PLACE FOLLOUING MACRO MAC I C STATEMENT;

MACRO MAC 1 C

- METH- [i OR ME TH= E OR METH= F

7.

• SELECT START OF JULIAN DATE RANGE;

♦ PLACE AFTER MACRO MAC 1 D STATEMENT;

MACRO MAC ID

■ 76205

*

♦ SELECT END OF JULIAN HATE RANGE;

♦ PLACE AFTER MACRO MAC I E STATEMENT;

MACRO MAC IE

- 76247
7

♦ SPECIFY COLONY OR COLONIES DESIRED;

♦ ENCLOSE SC VALUE IN SINGLE QUOTES:

♦ PLACE AFTER MACRO MAC 1 F STATEMENT;

MACRO MAC 1 F

•SC* 'AA

♦ SPECIFY IF PLOTS ARE DESIRED FOLLOUING THE MACRO MAC 1 G STATEMENT;

♦ 1 =PL0T 2rNO PLOT;

MACRO MAC 1 G

1

7.

♦ THIS MACRO CONTAINS THE PLOTS;

MACRO MAC 1 H

PROC PLOT DAT A = RE AD Y ; FORMAT SC ISCC. METH IMET . ; BY SC METH;

PLOT TOTAL *MAC 1 A= • ; TITLE PUPAE COUNTS BY MAC 1 B ;

PROC PLOT DATA=REAOY; FORMAT SC ISCC. METH IMET.; BY SC METH;

PLOT CUM TOT *MAC 1 A= * ; TITLE CUMULATIVE PUPAE COUNTS BY MAC 1 B ;

PROC PLOT DAT ALREADY ; FORMAT SC ISCC. METH IMET.; BY SC METH;

PLOT TOTAL *MAC 1 A = T CUMTOT ♦MAC1 A- C /OVERLAY;

TITLE UEEKLY (T) AND CUMULATIVE <C) PUPAE COUNTS BY MAC 1 B ;

OPTIONS CENTER;

DATA OLD; INFILE BUG; INPUT

r<T 2 SC I 3-4 JD 17-18 BJD 16-18 YR H-15 DATEA 14-18
METH I 69 TOTAL 70-73 1 ;

UK=C£ IL ( BJD 7);

SASDATE 'DATE JUL (DATEA ) ;

MON=MONTH( SASDATE ) ;

OTR-CEIL ( MON/4 ) ;

IF METH= 1 : METH- 5 ' I METH= A THEN METH= D ;

IF ME TH= 3 ! METH= ' 7 ’ ! METH= C THEN ME TH= E ;

IF ME TH= 2 I ME TH= 6 ! METH='B THEN METH= ' F ;

IF MAC 1C ;

IF MAC1 D< = DATEA<' = MAC1 E AND ( MAC 1 F ) ;

PROC SORT BATA-OLD; BY MAC 1 ;

PROC MEANS NOPRINT MIN DATA-OLD: BY MACK
VARIABLES DATEA; OUTPUT OUT=TRYIT2 MIN=STARTDTE ;

DATA 0LD2;

MERGE TRYIT2 < IN-OK I ) OLD (IN=0K2) ; BY M AC 1 ;

[F OKI $ 0K2;

PROC SORT DAT A = 0L D2 ; BY SC METH DATEA;

PROC MEANS NOPRINT MEAN ST DERR SUM;

BY SC METH S TART DTE MAC 1 NOTSORTED;

VARIABLES TOTAL ; OUTPUT 0UT = GO1 MEAN-'MEAN STDERR-S _E_ SUM =T OT AL ;

PROC SORT; BY SC METH START DTE;

DATA READY; SET G0 1 ; BY SC METH ;

RETAIN CUM TOT 0 MAC 1 A 0;

IF FIRST. METH OR FIRST. SC THEN CUM. TOT-O;

IF FIRST. METH OR FIRST. SC THEN MACIAS;

CUM TOT rCUM TQT + TOT AL ;

MAC 1 A-MAC 1 A* 1 ;

PROC PRINT DATA=READY; FORMAT SC ISCC. METH IMET. MEAN S_E_ TOTAL 6.2 CUM TOT \
2 ; BY SC METH; ID MACK

VARIABLES STARTDTE MAC 1 A MEAN S.E. TOTAL CUM.TOT;

TITLE PUPAL PRODUCTION BY MAC 1 B ;

DATA READY; SET READY;

SELECT =MAC1G;

IF SELECT* 1 ;

MAC1H;

END OF DATA

274

e:<ec UOIOl.stat 'pi'

FILE SORTLIB NOT FREED, IS NOT ALLOCATED
FILE SORTUKOI NOT FREED, IS NOT ALLOCATED

FILE SORT UK02 NOT FREED, IS NOT ALLOCATED

FILE S0RTUKO3 NOT FREED, IS NOT ALLOCATED

FILE SYSOUT NOT FREED, IS NOT ALLOCATED

NOTE: THE JOB SEANJOI HAS BEEN RUN UNDER RELEASE 79. 2B OF SAS AT UASHINGTON COHPUTER CENTER.
NOTE: THIS IS AN EXPERIMENTAL TEST RELEASE OF SAS. PLEASE

REPORT ALL PROBLEMS TO YOUR INSTALLATION REPRESENTATIVE.

NOTE:

NOTE:

NOTE:

NOTE:

NOTE:

NOTE:

NOTE:

NOTE:

NOTE:

NOTE:

INFILE BUG IS:

DSNAME=SEANJ01 . D0 1 0 1 .TAD 1 7. DAT A,

UNIT=SYSDA,VOL=SER=DATA18,BISP=SHR,

DCB= (BLKSIZE=4840,LRECL= 1 21 ,RECFN=FB)

INFILE BUG HAS 5665 LINES.

DATA SET UORK . OLD HAS 26 OBSERVATIONS AND 12 VARIABLES. 138 OBS/TRK .
DATA SET UORK. OLD HAS 26 OBSERVATIONS AND 12 VARIABLES. 138 OBS/TRK.
DATA SET UORK.TRYIT2 HAS 7 OBSERVATIONS AND 2 VARIABLES. 612 OBS/TRK.
DATA SET UORK.OLD2 HAS 26 OBSERVATIONS AND 13 VARIABLES. 126 OBS/TRK.
DATA SET U0RK.0LD2 HAS 26 OBSERVATIONS AND 13 VARIABLES. 126 OBS/TRK.
DATA SET UORK.GOI HAS 7 OBSERVATIONS AND 7 VARIABLES. 258 OBS/TRK.
DATA SET UORK.GOI HAS 7 OBSERVATIONS AND 7 VARIABLES. 258 OBS/TRK.
DATA SET UORK. READY HAS 7 OBSERVATIONS AND 9 VARIABLES. 192 OBS/TRK.

PUPAL PRODUCTION BY UEEK 14:00 UEDNESDAY , DECEMBER 26, 1979 1

SC = AA = CULICOIBES VARI IPENNIS SONORA METHYLS OF PUPAE - - -

UK

STARTDTE

CUN_UK

MEAN

S.E.

TOTAL

C U H _ T 0 T

30

76205

1

4.75

0.75

19.00

19.00

31

7621 2

2

5.60

0.98

28.00

47.00

32

7621 9

3

4.50

0.65

18.00

65.00

33

76226

4

5.38

0.94

21.50

36.50

34

76233

5

3.13

0.83

12.50

99.00

35

76240

6

6.75

0.83

27.00

126.00

36

76247

7

7.00

7.00

133.00

NOTE: DATA SET UORK. READY HAS 7 OBSERVATIONS AND 10 VARIABLES. 168 OBS/TRK.

Figure 5.— SAS output from the pupal production program shown in fig. 4. The general data-
retrieval request is shown in appendix C, 1.

A

LARVAL TEMPERATURE DATA

BY UEEK

jl-hh-luliluililj vhk i i r c. nn i o

UK

STARTDTE

M E A N _ H I SE.HI

HI.MAX

HI. MIN

MEAN.L0 SE.L0

L0.MAX

L0.HIN

30

74205

28.09

0.04

28.5

28.0

25.86 0.10

26.5

25.5

31

7421 1

28.47

0.08

29.0

28.0

25.20 0.10

26.0

25.0

32

74218

28.75

0.08

29.0

28.5

25.00 0.16

26.0

24.0

33

74225

28.08

0.10

28.5

27.5

25.00 0.20

26.0

24.0

34

74232

28.46

0.26

29.5

27.0

25.50 0.24

26.5

24.0

35

74239

28.25

0.14

29.0

27.5

26.08 0.22

27.0

25.0

34

74244

27.47

0.17

28.0

27.5

26.17 0.17

26.5

26.0

B

TEMPERATURE DATA

FOR LARVAL

DEVELOPMENT

FOR SELECTED CAGES

1 1 PENN I S S

OL-hH-LULHUil'113 Villi

LCN

STARTDTE

END.

DTE

MEAN. HI

SE.HI

HI.MAX

HI. MIN MEAN.L0

SE.L0

L 0 _ M A X

L 0 _ M I N

2400

74155

761

71

27.47

0.14

28.50

26.50 25.44

0.16

26.50

24.00

2401

74154

741

72

27.47

0.14

28.50

26.50 25.47

0.15

26.50

24.00

2402

74154

74172

27.47

0.14

28.50

26.50 25.47

0.15

26.50

24.00

2403

74157

761

73

27.47

0.14

28.50

26.50 25.56

0.14

26.50

24.00

Figure 6.— A, Output from retrieval request for larval rearing temperature by week (see appendix C,
3). B, Output from retrieval request for histories of larval rearing temperatures for specific cages of
adult flies (see appendix C, 3).

276

A

SUMMARY OF ADULT

DISPOSAL

BY UEEK

- . —

- YR=76 SC=AA=

CULICOIBES UARIIPENNIS SONORA -

UK

STARTDTE CAGES COLONY

RESEARCH BOTH

DISCARD

30

76205

4

0.00

0.00 4.00

0.00

31

7621 1

15

8.00

7.00 0.00

0.00

32

76218

2

0.00

0.00 2.00

0.00

33

76225

12

10.00

2.00 0.00

0.00

34

76232

12

12.00

0.00 0.00

0.00

35

76239

4

0.00

0.00 4.00

0.00

36

76246

3

2.00

1.00 0.00

0.00

TOTAL

•

52

32.00

10.00 10.00

0.00

5! TOTAL

•

8

61 .54

19.23 19.23

0.00

B

USE OF FLIES - LISTING BY UEEK

--- SC=AA=CULICOI DES VAR1 IPENNIS SONORA WK=30

STARTDTE=76205 END.

DTE=762 1 0 . .

CAGE

UISP1

DISP2

2461

ADULTS FOR

BOTH

ADULTS FOR BOTH

2462

ADULTS FOR

BOTH

ADULTS FOR BOTH

2463

.

ADULTS FOR COLONY

2464

ADULTS FOR

RESEARCH

.

2465

.

ADULTS FOR COLONY

2466

ADULTS FOR

RESEARCH

.

2467

ADULTS FOR COLONY

2468

ADULTS FOR

RESEARCH

.

2469

.

ADULTS FOR COLONY

2470

ADULTS FOR

RESEARCH

.

2471

.

ADULTS FOR COLONY

Figure l.—A, Output from retrieval request for summary of
adult fly use (disposal) by week (see appendix C, 8). B, Output
from retrieval request for summary of adult fly use (disposal)
for specific cages of adult flies (see appendix C, 8).

A

FEEDINGS=1 SC=AB=PSOROPHORA COLUNBIAE START=74205 END_DTE=74247

DATE

CAGE

SEQUENCE

TYPE

74 220

113

1

102 SUCROSE

+ UATER

74230

114

1

102 SUCROSE

♦ UATER

FEEDINGS=3 SC=AA=CULICOIDES VARIIPENNI9 SONORA START=74205 END_DTE=74247 -

DATE

CAGE

SEQUENCE

TYPE

74207

2444

1

UATER ONLY

74208

•

2

102 SUCROSE ♦ UATER

74208

3

UATER ONLY

74208

2444

1

UATER ONLY

74213

•

2

102 SUCROSE + UATER

B

IAIAN DATE

MEAN U1N0

MEAN DRY

OF O.C.

NO. OF PUPAE

LENGTH <NM)

UE1GHT (NICROGRAN!

SAMPLE

PER HL.

♦OR- 8.E.

♦OR- S.E.

74238

702

1.43 +0R- 0.01

83.43 +0R- 2.39

74244

487

1.48 +0R- 0.02

87.30 +0R- 2.90

74232

712

1.44 +0R- 0.01

84.07 +0R- 3.11

74237

472

1.31 +0R- 0.01

93.78 +0R- 2.80

74244

470

1.50 +0R- 0.01

93.30 +0R- 2.49

74271

484

1 .47 +OR- 0.01

89.40 +0R- 2.24

Figure 8.— A Outputs from retrieval requests for summary of
nutritional regimes for a mosquito and a gnat colony (see appen¬
dix C, 10). B, Output from retrieval request for quality -control
data on fly size for a specific colony and time span (see appen¬
dix C, 12).

278

pupal production program. Then data inputs were entered
(for example, Julian dates,3 colony codes, and cage
numbers— appendix C). Next, the SAS program was run
and the report listed (printed) as output. The data inputs
were made or changed after the MACRO MAC 1 X
statements that appeared after each CHOOSE, PLACE,
or SELECT statement. The output generated by running
the SAS pupal production program (fig. 4) is shown in
fig. 5 (plot excluded). Examples of the tables produced
from some of the retrieval programs listed in appendix C
are shown in figs. 6-8, with the execution statements and
plots omitted for brevity. The actual computer operation
of the retrieval programs and the system itself required
several computer programs (appendix D).

Discussion

Since every insectary can benefit from logical flow charts
and well-planned data-collection documents, systems
analysis is advantageous even without being linked to a
computer, as it encourages efficient performance of tasks.
In particular, coding and recording of quantitative infor¬
mation increases the accuracy of data collection. Techni¬
cians find the ADP system easy to use and, because of
the flow charts and clearly marked forms, are unlikely to
forget duties or fail to record important information. The
system is an effective management tool since consistent
and easily reviewed documents are readily provided by
the computer to evaluate variables associated with insect
production— environmental conditions; nutritional
regimens; and the quality, quantity, and use of insects.
Therefore, one of our goals is to enter data into the com¬
puter at least weekly so that we can immediately change
procedures in response to quality-control and production
reports (see “Putting the Control in Quality Control in
Insect Rearing,” by D. L. Chambers and T. R. Ashley
and “The Closed-Loop System of Quality Control in In¬
sect Rearing,” by J. C. Webb).

Our system has evolved slowly; by 1980, we had used the
data-collection part for more than 4 years, the editing
part for 2 years, and the retrieval request for 6 months.
But the current data base is restricted to the one
80-column entry document. This limitation has caused
some codes (for example, for pupal production) to become
complex and some information to be lost (for example,
disposition of flies coded for research without an indica¬
tion of the recipient). So we are expanding the system to
include almost every aspect of our insect-rearing pro¬
gram. The subjects eventually covered by this expansion

Mulian date: a 5-digit system of recording dates that begins
with the last 2 digits of the year followed by 3 digits to record
the day in a numbering system between 1 and 365/6 for each
year; Feb. 15, 1981, would be 81046.

will include adult production, physical variables and their
parameters, pupal and egg production, quality control,
larval production and use of immatures, production costs,
and shipments to other laboratories. We will also enter
the data through the TSO terminal rather than from com¬
puter cards.

For those who might wish to use systems analysis but
are unfamiliar with computer technology, we recommend
an introductory computer course that will give them a
working knowledge of the terminology. Then, a systems
analyst could be consulted before a new system for a
specific insect-rearing operation is planned or created.

The application of systems analysis should include an
assessment and examination of the organization and
operation of the insect-rearing program, evaluation of ex¬
isting data-collection forms and other documents, comple¬
tion of a feasibility study determining the usefulness of
ADP to the particular program, design, and implementa¬
tion. After the system is in place, there should be an
evaluation, and changes should be made to improve the
system if necessary. The techniques required to ac¬
complish these steps include flow charting, summarizing,
coding, and form designing. Also, computer software and
hardware needs would have to be considered. We en¬
courage those who incorporate these concepts into insect¬
rearing programs and hope that this report will foster use
of systems analysis and information-system theory in the
management of insect rearing.

Acknowledgments

We thank the personnel of the CDSD for their assistance,
cooperation, and support in this project; particularly J.
Belt, B. Davies, A. Gardiner, C. Gerhardt, J. Littrell, C.
Mitchell, J. Palmes, and L. Penney. We especially ap¬
preciate the help of A. Weiner (U.S. Department of
Agriculture) in planning and preparation of data forms.

References

Akey, D. H.; Potter, H. W.; and Jones, R. H.

1978. Effects of rearing temperatures and larval den¬
sity on longevity, size, and fecundity in the
biting gnat Culicoides variipennis. Ann. En-
tomol. Soc. Am. 71: 411-418.

Alexander, M. J.

1974. Information systems analysis. 424 pp. Science
Research Associates, Chicago.

Bingham, J. E., and Davies, G. W. P.

1972. A handbook of systems analysis. 191 pp.
Halsted Press, New York.

Chambers, D. L.

1977. Quality control in mass rearing. Annu. Rev. En-
tomol. 22: 289-308.

FitzGerald, J. M., and FitzGerald, A. F.

279

1973. Fundamentals of systems analysis. 531 pp.
John Wiley & Sons, New York.

Gerberg, E. J.

1970. Manual for mosquito rearing and experimental
techniques. Am. Mosq. Control Assoc. Bull. 5,
109 pp.

Gore, M., and Stubbe, J.

1975. Elements of systems analysis for business
data processing. 427 pp. Wm. C. Brown Co.,
Dubuque, Iowa.

Jones, R. H.; Potter, H. W.; and Baker, S. K.

1969. An improved larval medium for colonized

Culicoides variipennis. J. Econ. Entomol.

62: 1483-1486.

Lucas, H. C., Jr.

1974. Toward creative systems design. 147 pp.
Columbia University Press, New York.

Sippl, C. J., and Sippl, C. P.

1972. Computer dictionary and handbook. 2d ed., 778
pp. Howard W. Sams and Co. and Bobbs-
Merrill Co., New York.

Youssef, L. A.

1975. Systems analysis and design. 228 pp. Reston
Publishing Co., Reston, Va.

280

Appendix A.— Computer Codes and Editing Criteria for the Colonies-Adult-Production Record Document, Figure 3
Column

number Description

1-15 . I. Record identifier.— Alphanumerically coded data are stored in 15 columns comprising 7 fields that identify each

record (a horizontal line of data in figure 3) from all other records.

1 . . A. File code. — 1 -column numeric field to denote investigator or research project. Valid entry is 1 (entomology).

2 . B. Record type. — 1 -column numeric field to denote type of research record used. Valid entry is 1 (colonies-adult-

production record).

3-9 . C. Population.

1. Colony.— 2-column, alphabetic field to record genera, species, and particular population of insects that per¬
tain to data in the record. Valid entries for established colonies are AA-BZ. Code ZZ means no adult insects
are present, and only location data in 53-68 are being recorded for the year and day given in 14-18;
columns 5-13, 19-52, and 69-79 must be blank.

Species

AA—Culicoides variipennis (000) . Texas 1957.

AD —Aedes aegypti . Texas 1974.

AF —Culex tarsalis . Colorado 1972.

AG —Culiseta inomata . Colorado 1972.

AK —Culicoides variipennis (036) . Idaho 1973.

AP— Aedes taeniorhynchus . Florida 1975.

5-8 . 2. Large cage No.— 4-column numeric field that identifies large emergence cages. The number is assigned se¬

quentially for each large cage when it is established. For each colony, numbers must follow sequentially and
may not be duplicated unless each duplicate is accompanied by a different small-cage number in 9-11. Valid
entries are 0001-9999.

9-11 . 3. Small cage No.— 3-column numeric field that identifies small cages set up sequentially from 1 large cage.

Numbers may not be duplicated if they are accompanied by an identical large-cage number in 5-8. Small-cage
numbers are usually issued by the user of flies so that field 9-11 is usually blank. Valid entries are 001-999.

12-13 . D. Generation.— 2-column numeric field to denote generation of colony. Parent generation (P) is 01. The number is

sequential for successive generations. If the colony is a population without discrete generations, the code is 00.
The same generation number is valid for multiple records with the same colony code, but nonsequential genera¬
tion numbers are invalid (the generation number must be the same or in sequence). Valid entries are 00-99.

14-15 . E. Year.— 2-column numeric field that denotes the year that data were collected. The last 2 digits of the year are

entered. Valid entries are less than or equal to the current year and greater than or equal to 74.

16-28 . II. Emergence rime.— Numerically coded data are stored in 13 columns comprising 4 fields to record start and end of in¬

sect emergence.

16-22 . A. Start. (Pupae placed in cage).

1. Julian day.— 3-column numeric field that denotes Julian day. Valid entries are 001-365 for regular years and
001-366 for leap years.

19-22 . 2. Time.— 4-column numeric field to record time of day when pupae are placed in the cage. Military time is used.

Laboratory practice is to record to the closest hour; for some special tests, the entry may be to the nearest
minute. Valid entries are 0001-2400.

23-28 . B. End. (Pupae removed from cage).

23-24 . 1. Day.— 2-column numeric field to record last 2 digits of Julian day. Entries must be greater than or equal to

the Julian day in 16-18, and elapsed days from Julian day in 16-18 must be less than or equal to 14. Valid
entries are 00-99.

25-28 . 2. Time.— 4-column numeric field to record the time when pupae are removed from the cage. (See IIA2.) Valid

entries are 0001-2400.

281

Column

number Description

29-52 . III. Adult nutrition.— Numerically coded data are stored in 24 columns comprising 9 fields to record adult-nutrition data.

A new feeding regimen begins when different nutrition is given to a cage of flies, and the code for food type must
change.

29-36 . A. 1st Feeding.

29-30 . 1. Day.— 2-column numeric field to record the last 2 digits of the Julian day on which the 1st feeding regimen

was begun. Entries must equal the last 2 digits of the Julian day in 16-18. Valid entries are 00-99.

31-34 2. Time placed.— 4-column numeric field to record the time the 1st feeding was begun. (See IIA2.) Valid entries

are 0001-2400.

35-36 . 3. Food type.— 2-column numeric field to record the first type of nutrition given. Water is always present for

emerging adults in the pupal dish; so the lst-feeding regimen is coded 01 if no other nutrition was given, and
values in 16-22 are used for day and time. Valid entries are 01-05.

Codes

01— water only,

02— sucrose solution (5%-10%) with water given also,

03— sugar cubes and water,

04— raisins and water,

05— sugar solution for 1 day alternated with water only for 2 days.

37-44 . B. 2nd Feeding.

37-38 1. Day.— 2-column numeric field to record the last two digits of the Julian day on which the 2d feeding regimen

was begun. Columns 37-38 are blank if 29-36 are blank. The value in 37-38 must be greater than or equal
to the value for 29-30. If the value in 29-30 equals that for 37-38, then the value in 31-34 must be less than
the value in 39-42. Valid entries are 00-99.

39-42 . 2. Time placed.— 4-column numeric field to record the time the 2d feeding regimen was begun. (See IIA2.) If

the value in 37-38 equals that for 29-30, then the value in 39-42 must be greater than the value in 31-34.
Valid entries are 0001-2400.

43-44 . 3. Food.— 2-column numeric field to record the 2d type of nutrition given. (See IIIA3.) Columns 43-44 must be

blank or contain a value different from that in 35-36. Valid entries are 01-05.

45-52 ... C. 3rd Feeding.— \i 37-44 are blank, then 45-52 must be blank.

45-46 . 1. Day.— 2-column numeric field to record last 2 digits of the Julian day on which 3d feeding regimen was begun.

Columns 45-46 are blank if 37-44 are blank. The value in 45-46 must be greater than or equal to the value
in 37-38. If the value in 37-38 equals that for 45-46, then the value in 39-42 must be less than the value in
47-50. Valid entries are 00-99.

47-50 . 2. Time placed.— 4-column numeric field to record the time 3d feeding regimen was begun (See IIA2.) If the

value in 45-46 equals that for 37-38, then the value in 47-50 must be greater than the value in 39-42.
Valid entries are 0001-2400.

51-52 3. Food type.— 2-column numeric field to record the 3d type of nutrition given. (See IIIA3.) Columns 51-52

must be blank or have a value different from that in 43-44. Valid entries are 01-05.

53-68 IV. Location. — Numerically coded data are stored in 16 columns comprising 7 fields to record data on physical variables

(see “physical parameters” in fig. 3) for insect-rearing rooms for the Julian day entered in columns 16-18. The 7
location data fields, 53-68, may not be blank if the serial code in 3-4 is ZZ.

53-54 A. Room.— 2-column numeric field to code the room in the building where environmental data are taken. Valid en¬

tries are 01-09.

Codes

01— building 106-B, colony room 1,
02— building 106-B, colony room 2,
03— building 106, colony room,

04— building 106-A, colony room,
09— budding 106, laboratory.

282

Column

number Description

55-68 . B. Physical parameters. — Variables in a room or incubator for a given day.

55-60 . 1. Larval pan temp. °C.

55-57 . a. High.— 3-column numeric field to record highest temperature (taken by hand). Enter temperature to 1

decimal place with decimal point assumed between 56-57. The value must be greater than or equal to the
low temperature value in 58-60. If a value is in 55-57, then 58-60 may not be blank. Valid entries are
blank or 150-400.

58-60 b. Low.— 3-column numeric field to record lowest temperature (taken by hand). Enter temperature to 1

decimal place with a decimal point assumed between 59-60. The value must be less than or equal to the
high temperature value in 55-57. If a value is in 58-60, then 55-57 may not be blank. Valid entries are
blank or 150-400.

61-68 . 2. Adult holding.

61-64 . a. Temp. °C.

61-62 (1) H.— 2-column numeric field to record the highest temperature on a high-low thermometer. Enter

temperature to the nearest whole number. The value must be greater than or equal to the low in
63-64. If a value is in 61-62, then 63-64 may not be blank. Valid entries are blank or 15-40.

63-64 (2) L.— 2-column numeric field to record the lowest temperature on a high-low thermometer. Enter

temperature to the nearest whole number. The value must be less than or equal to the high in
61-62. If a value is in 63-64, then 61-62 may not be blank. Valid entries are blank or 15-40.

65-68 . b. RH. — Relative humidity.

65-66 (1) H.— 2-column numeric field to record the highest relative humidity on a hygrothermograph. Enter

relative humidity in percent to the nearest whole number. The values must be greater than or equal
to the RH-value in 67-68. If a value is in 65-66, then 67-68 may not be blank. Valid entries are blank
or 00-99.

67-68 . (2) L.— 2-column numeric field to record the lowest relative humidity on a hygrothermograph. Enter

relative humidity in percent to the nearest whole number. The values must be less than or equal to
the high RH-value in 65-66. If a value is in 67-68, then 65-66 may not be blank. Valid entries are
blank or 00-99.

69-73 . V. Pupal Prod.— Alphanumerically coded data are stored in 5 columns comprising 2 fields to record the counting method

and the number of pupae collected.

69 . A. CM.— Counting method; one-column alphanumeric field to record method used for pupal counts in 70-73. The

numeric code records pupae picked (harvested) on the same date (same batch). The alphabetic code records pupae
combined from different picking dates. Valid entries are 1-7 and A-C.

Codes

1— number of pupae per milliliter;

2— estimate;

3— actual count;

4— pupae picked previously and remaining pupae moved to new cage;

5— pupae picked, placed in more than 1 cage, and all cages labeled “5;” the value equals the number of pupae per
milliliter;

6— pupae picked, placed in more than 1 cage, and all cages labeled “6;” the value is estimated;

7— pupae picked, placed in more than 1 cage, and all cages labeled “7;” the value equals the actual count.

A— pupae, from different picking dates, combined; the value equals the number of pupae per milliliter;

B— pupae, from different picking dates, combined; the value is estimated;

C— pupae, from different picking dates, combined; the value equals the actual count.

70-73 . B. A^o.— 4-column numeric field to record the number or amount of pupae. If codes that record the number of pupae

per milliliter in column 69 are 1, 5, or A, a decimal point is assumed between 72-73. If the value for 70-73 is 0000,
then the code for the counting method in column 69 must be 4, A, B, or C. If column 69 has an alphabetic code,
then 0000 in 70-73 means pupae are recorded on a trailer card (a second card); the value in 70-73 (with an
alphabetic code in column 69) records only new pupae that were placed with the remaining older pupae. Valid en¬
tries are 0000-9999.

283

Column

number Description

74-79 VI. Adult disposition.— Numerically coded data are stored in 6 columns comprising 4 fields to record whether flies were

used for research and/or for a colony; 2 divisions of adult disposition— research and colony— contain 2 fields each to
record how and when flies were used.

74-76 . A. Research.

74 . 1. How. — 1-column numeric field to record each large cage of flies that was used for research. If the value in col¬

umn 74 is 3, then the value in column 77 must be 3. If column 74 is blank, then the value in column 77 must
be 1 or 5. The values 1 and 5 may only be used in column 77; 3 may be used in both columns 74 and 77. Valid
entries are blank, 2 or 3.

Codes

1— adults to colony,

2— adults to research,

3— adults to both,

4— undefined,

5— discarded.

75-76 . 2. Day.— 2-column numeric field to record the last 2 digits of the Julian day when flies were used for research.

Values must be greater than or equal to the Julian day in 16-18 and less than or equal to the value in 78-79.
Valid entries are 00-99.

77- 79 . B. Colony.

77 . 1. Dow.— 1-column numeric field to record large cage of flies used for a colony. (See VIA1.) If the value in col¬

umn 77 is 3, then the value in column 74 must be 3. If column 77 is blank, then the value in column 74 must be
2. Valid entries are blank, 1, 3, and 5.

78- 79 2. Day.— 2-column numeric field to record last 2 digits of the Julian day when flies were used for the colony.

If the value in column 77 is 3, then the value in 78-79 must be greater than or equal to that in 75-76. Valid
entries are 00-99.

80 . VII. Card type.— Numerically coded data are stored in a 1-column field to distinguish the primary card from trailer cards.

The field may not be blank. Valid entry is 1 for the primary card.

284

Appendix B. — Computer Program Written From the Editing Document, Appendix A, to Verify Data4

L18T "SEAMJOI .84101 .COLVERIF .COBOL '
'8EANJ01 .80101 .C9LVER1F.C0I0L7
OHIO IDENTIFICATION B1VI810N.

00020 PROGRAM-ID. COLVERIF.

00030 MICHMITTCN. DECEMBER, 1 477.
00040 ENVIRONMENT 9IV191M.

00030 CONF10URATION SECTION.

00060 SPECIAL -EASE 8.

00070 COt 18 TOPLIBE

00080 C8P 18 WORE .

00090 80NRCE -COMPUTER. IDM-370.

00100 OBJECT -COMPUTER. IDM-370.

OOttO INPUT-OUTPUT SECTION.

00120 FILE-CONTROL.

SELECT TRANS
SELECT MA8TR
SELECT EL1BT
SELECT ERN8G
DATA IIV18I0N.
FILE SECTION.

A8II0M TO UT-8-IN1.
A88I8M TO UT -3-OUT 1 .
ASSIGN TO UT-8-0UT2.
ASSIGN TO 8T-8-IN2.

F0

00130
00140
00154
00160
00170

ooieo

00190 '

00200
00210
00220
00230
00240
00230
00200
00270 0
00280 •

00290 • BASTE IS THE OUTPUT FILE. ALL RECORDS THAT ARE ERROR FREE ARE
00300 • WRITTEN TO THIS FILE. THIS FILE CONTAINS ONLY BATA FROM THIS
00310 • PASS.

00320 •

00330 FB
00340
00330
00340
00370
00384 01
00390 •

00400 o LIST IS THE PRINTER TRANSACT I6N/ERR0R LIST.

00410 «

00420 FB
0043®

00444
00439
00460
00470 01

00460
00490
OOSOO
00510 •

00321 •

00334 •

00340 •

00330 •

00360 •

00370 «

00380 •

00390 •

00600 •

00619 •

00424
00430
00640
00630
00660
00470
00680
00640
00700
00710 •

00720 •

00730 •

00740 •

00730 77 A PIC 999

00760 77 LINEZ PIC 94

00770
00780
00790 •

00800 77 DAY-STORE PIC 49

00810
00820
00830
00840
00830 77

00860 77

00870 77

00680 77

00890 77

00900 01

00910 • *

00920 • DUPE31 AND DUPE82 ARE USED TO CHECI FOR 80 COLUHN DUPLICATES IN
00930 • THE TRANSACTION FILE.

00940 •

00930
00960
00970

TRANS IS THE INPUT TRAN8ACTIBN FILE.
TRANS

REC0RDIN8 NODE 18 F
LABEL RECORDS ARE STANDARD
BLOCK CONTAINS 0 RECORDS
DATA RECORD 13 INPT.

INPT PIC 11121).

MA8TS1

RECORDING NODE IS F
LABEL REC0ND8 ARE STANDARD
BLOCK CONTAINS 0 SECONDS
DATA RECORD 18 OUT.

OUT PIC 1(121).

ELI8T

RECORDING RODE IS F
LABEL SECONDS ARE STANDARD
BLOCS CONTAINS 0 RECORDS
DATA RECORD 18 PRT.

PRT.

03 FILLER PIC I.

03 BODY PIC 1(121).

03 FILLER PIC 1(11).

ERNB8 IS THE FILE OF ERROR HE88A8EB. EACH ERROR MESSAGE HAS A

SEQUENTIAL NUMERIC CODE IN OC 79-80. ERROR HE88A0ES CAN BE
ADDED AS NEEDED, INCREA8IN8 THE SUBSCRIPT AND INCREA8INB THE
VALUES IN 6TATENENT8 • 2 77 TO REFLECT THE INCREASED SIZE
( INCREMENTS OF 80 >, C TOT 0 278 OCCURS X TINES' < I ACRE BENTS
OF 1 ), 8TRT 6 289 ( INCREHENT8 OF 1 ) AND ST BT • 290 OCCURS
X TIKES' ( INCNEHCRTS OF 1 ). SINCE EACH HE88A8E CARRIES ITS OUN
SUBSCRIPT VALUE, THESE RECORDS DO NOT HAVE TO BE IN SEQUENCE
UHEN THET ARE READ IN.

FB

01

ERH8Q

RECORDINS BODE IS F
LABEL RECORDS ARE STANDARD
BLOCK CONTAINS 0 RECORDS
DATA RECORD 18 E-N.

ER.

03 FILLER PIC 1(78).

03 E-NO PIC 99.

U0RK1R8- STORAGE SECTION.

A IS INCREKEHTED IN 6THT N 440, AND IS USED TO CALCULATE THE DATS
UNTIL EMER8EHCE.

VALUE OOO.
VALUE 94.

DAY-STORE RECEIVES THE FIRST FEED1N6 DAT FOR NUMERIC COUP ARISONS.

VALUE 00.

B IS INCREMENTED IN THE ERROR CHECK LOOP (STMT • 707).

7 B PIC 49 VALUE 00.

181 PIC 9 VALUE 0.

MRED PIC 9(3) VALUE OOOOO.

MROTE PIC 9(3) VALUE OOOOO.

HERR PIC 9(3) VALUE OOOOO.

HD-8W PIC X VALUE ' '.

P1DJN PIC X ( 1 1 ) VALUE HIBH-VALUE8.

01

DUPE81 .

03 DUPEH0LD1 PIC X(121>.

03 FILLER REDEFINES DUPEH0L81.

00900
00990
OtOOO 01
01916
01020
01030
01040
01030 •
01040 •
01070 •
01080 •
04090 01

01109
01119
01129
01139
01149 01
01139
01169
01170 •
01169 •
01190 •
01200 •
01210 01
01220
01230
01240
01230
01260
01270 •
01289 •
01290 •
01301 01
01310
01129
01330
01340
01350
01369
01379
01380
01399
01400
01419
01420
01439
01440
01430
01469
01479
01486
01499
01300
01310
01320
01330
01349
01330
01369
01370
01390
01399
01609
01610
01620
01639
01640
01650
01660
01676
01600
01699
01790
01710
01720
01739
01740
01730
01760
01770
01786
01790
01600
01819
01620
01830
01849
01830
01869
01870
01880
01890
01900
01910
01420
01930
01940
01930
01960
01479
01980
01990

03 RECH0LD1 PIC X<114).

03 FILLER PIC X(7).

DHPC82.

03 DUPEH0LD2 PIC X(121).

03 FILLER REDEFINES DUPEH0LD2.

03 RECH0LD2 PIC X(I14>.

03 FILLER PIC 1(7).

J-DAI-CALC IB USED IN EMERGENCE CALCULATIONS A8 WELL A8 INTER-DATE
RELATIONSHIPS.

J-DAY-CALC.

03 J-DAY-DA8E PIC 994.

01 FILLER REDEFINES J-DAY-0A9E .

03 FILLER PIC 9.

03 IE8ULT PIC 44.

FILLER.

03 EOD PIC X VALUE ' '.

96 END-OF-DATA VALUE 'X7.

BASEDTE-DNK BREAKS DOWN THE IA8E JULIAN DAY 80 THE LA9T 2 DIBITS
ARE AVAILABLE FOR COMPARISONS.

BA8EDTE-DRK.

03 HOLD PIC 949.

03 BREAK REDEFINES HOLD.

03 FILLER PIC 9.

03 LA8T-2 PIC 99.

03 L9T-2 REDEFINES LAST-2 PIC XX. 8

U0RK-REC RECEIVES THE TRANSACTIONS ONE AT A TINE.

UORK-REC.

03 ID1 .

03 FILE-TYPE PIC X.

88 FILE-OK VALUES '1'.

03 RECORD-TYPE PIC X.

ee RECORD-OK VALUES 'I7.

03 SERIAL -CODE PIC XX.

08 SER-CD-OB VALUES AA THRU 'IZ' 7ZZ7.

88 AK-CGLBWY VALUE AK'.

88 ZZ-COLONY VALUE ZZ'.

03 Li-CABE PIC XXIX.

66 LAR8E-GABE-0K VALUES 0001' THRU '9999'.

SB Z2-CA6E VALUE '

03 SM-CA8E PIC XXI.

68 8HALL-CA0E-OK VALUES 000 THRU '499' 7
01 8ENERAT10N PIC XX.

SB GENERA! 10N-8K VALUES 01' THRU '99' 7 7.

88 AA-9ENERATIGN VALUE 00'.

02 JD.

03 BASE-DATE.

03 D-YEAR PIC XX.

08 YEAR-VALID VALUES '74' THRU '497.

88 LEAR-YEAR VALUES '76' '80 84' 88' '92' 96'.

03 D-DAY PIC XXX.

88 DAY-VALID VALUES 001 THRU '363'.

68 DAY-366 VALUE '3667.

03 ENER8ENCE-8-T1NE PIC XXXX.

SB STARI-TIHE-OK VALUES OOOI' THRU 2400'.

86 AB-8TRT-TINE VALUE '0001' THRU 2400'.

01 EMERGENCE -END- DAY PIC XX.

88 VALID-END-BATE VALUES 00' THRU '99'.

03 ENF.R-END REDEFINES EMERGENCE -END- DAT PIC 99.

02 FILLER.

03 EHER6CNCE-E-T1HE PIC XXXX.

88 END-TINE-OK VALUES '0001' THRU '2400'.

86 AK-END-TIME VALUE 0061 THRU 2400'.

03 FEED1 -DAT PIC XX.

88 VALID-FEED1-DAT VALUES 00' THRU '49' 7 7.

03 FI -CHI REDEFINES FEED1-DAY PIC 99.

03 FEED1-TIKE PIC XXXX.

88 VALID- 1ST -FEED- TIME VALUES 'OOOI7 THRU 2400'

86 AK-1ST-FEEB-TIHE VALUE OOOI' THRU '2400' '

03 TTPE-F80D1 PIC XX.

08 VALID-FO0B1 -TYPE VALUES '01' THRU '03' '

03 FEED2-DAT PIC XX.

86 VAL ID-FEED2-DAT VALUES 00' THRU 99' ' 7.

80 N0-2ND-FEED-DAY VALUES ' '00'.

03 FEED2-TIME PIC XXXX.

88 VALID-2ND-FEED-T IKE VALUES 0001' THRU 2400' '

88 AK-2ND-FEED-TIKE VALUE OOOI7 THRU '2400' '

88 H0-2ND-FEED-TIME VALUES 7 ' OOOO'.

03 TYPE-FOOD2 PIC XX.

08 VALID-FOOD2-TYPE VALUES 01 THRU '03' '

03 FEED3-DAY PIC XX.

86 VALID-FEED3-DAY VALUE '00' THRU '99' ' '.

80 WO- 3RD-FEED-DAY VALUES ' ' '00'.

03 FEED3-TIKE PIC XXXX.

88 VALII-3RD-FEED-TIME VALUES 0001' THRU '2400' '

88 AK-1RD-FEED-TIKE VALUES 0000' THRU '2400' '

66 N0-3RD-FEED-TIBE VALUES ' ' '0000'.

03 TYRE-FQ0D3 PIC XX.

88 VAL ID-FOOD3-TYPE VALUE6 ' '01' THRU 05'.

03 ROOK-NO PIC XX.

88 VALID-ROOM-NO VALUES 01' THRU 09'.

03 LARVAI-HI6H-TERP PIC XXX.

88 VAL ID-LARVAL -HI 6H VALUE8 '150' THRU 400' '

03 LARVAL-LOU-TERP PIC XXX.

68 VALID-LARVAL-LOU VALUES '130' THRU '400' '

03 ADULT-HIBH-TEHP PIC XX.

88 0OOO-HI8H-TEKP VALUES '13' THRU '40 ' '.

“Program based on earlier and slightly different versions of
record and editing documents.

02001

02010

02020

02030

02040

02050

02060

02070

02080

02090

02100

02110

02120

02130

02140

02130

02160

02170

02180

02190

02200

02210

02220

02230

02240

02250

02260

02270

02280

02290

02300

02310

02320 •

02330 *

02340 •

02330

02360

02370

02380

02390

02400

02410

02420

02430

02440

02430

02460

02470

02480

02490

02500

02310

02320

02530

02540

02350

02360

02370

02580

02390

02600

02610

02620

02630

02640

02650

02660

02670

02680

02690

02700

02710

02720

02730

02740

02730

02760 •

02770 «

02780 •

02790

02800

02110

02820

02830

02840 *

02830 •

02860 •

02870 •

02880 •

02890 •

02900 •

02910

02920

02930

02940 •

02930 •

02960 •

02970 •

02980 •

02990 •

03000 •

03010

03 AD4JLT-L0W-TEMP PIC XX.

03020

03

FT

PIC

X.

81 GOOD-LOW-TEMP VALUES '15' THRU '40' ' '.

03030

03

RT

PIC

X.

03 ADULT-HIQN-RH PIC XX.

03040

03

SC

PIC

XX.

88 VALID-MIGH-RH VALUES '00' THRU '99' ' '.

03050

03

LC

PIC

XXXX.

03 ADULT-LOW-RH PIC XX.

03060

03

SMC

PIC

XXX.

88 VALID-LW-RH VALUES '00' THRU '99' ' '.

03070

03

GN

PIC

XX.

03 METHOD- AND-NO PIC XXXXX.

03080

03

II

PIC

XX.

88 BOTH-ZEROS VALUE '00000'.

03090

03

BD

PIC

XXX.

03 FILLER REDEFINES METHOD-ANI-NO.

03100

03

ES

PIC

XXXX.

03 COUNTING-METHOD PIC X.

03110

03

EEB

PIC

XX.

8D VALID-C-HETNOD VALUES 'A' THRU 'C' 'V THRU '7'.

03120

03

EET

PIC

XXXX.

88 NETHOD-MO-FL1ES VALUES 'A' THRU 'C' '4'.

03130

03

FID

PIC

XX.

03 K0-0F-FLIE8 PIC XXXX.

03140

03

FIT

PIC

XXXX.

86 ACCEPTABLE-RANGE VALUES 'OOOO' THRU '9999'.

03150

03

TF 1

PIC

XX.

88 NO-FLIES VALUE '0000'.

03160

03

F2D

PIC

XX.

03 BISPQSITI0N1 PIC X.

1 2/78CLH

03170

03

F2T

PIC

XXXX.

88 DI8P1 -VALID VALUES ' ' '2' '3'.

1 2/70CLH

03180

03

TF2

PIC

XX.

86 ADULT8-T0-D0TH1 VALUE '3'.

1 2/78CLM

03190

03

F3D

PIC

XX.

88 600D-W-BLANK VALUED '2' ' '.

12/78CLM

03200

03

F3T

PIC

XXXX.

03 DISP1 -DAY PIC XX.

1 2/78CLH

03210

03

TF3

PIC

XX.

88 VAL I D-D I SP 1 VALUES '00' THRU '99' ' '.

12/7BCLM

03220

03

RN

PIC

XX.

03 DISP0SITI0N2 PIC X.

12/78CLM

03230

03

LHT

PIC

XXX.

88 DIBP2-VALID VALUE8 ' ' '1' '3' '5'.

1 2/78CLM

03240

03

LLT

PIC

XXX.

88 ADULT8-T0-DOTH2 VALUE '3' '3'.

1 2/78CLM

03250

03

AHT

PIC

XX.

68 BLANK-VALUE VALUE ' '.

1 2/78CLM

03260

03

ALT

PIC

XX.

03 DISP2-DAY PIC XX.

1 2/78CLM

03270

03

AHR

PIC

XX.

88 VAL I D-D 1 SP 2 VALUES '00' THRU '99' ' '.

1 2/78CLM

03280

03

ALR

PIC

XX.

03 CARB-TTPE PIC X.

03290

03

CM

PIC

X.

88 CARD-TYPE-OK VALUE '1'.

03300

03

NF

PIC

XXXX.

03 8UAL ITT- CONTROL -DAT A PIC X(34).

03310

03

D 1

PIC

X.

03 FORM-TYPE PIC X.

03320

03

DID

PIC

XX.

03 BATCH PIC X(6).

03330

03

D2

PIC

X.

03340

03

D2D

PIC

XX.

HEAD 1 THRU HEAD6 ARE HEADINGS FOR THE ERROR LIST.

03350

03

CT

PIC

X.

03360

01 GET-DATE

01 HEAD1 .

03370

02

FILLER

PIC

03 FILLER PIC X(44) VALUE

03380

02

THIS-YEAR

PIC

AGRICULTURAL RESEARC'.

03390 PROCEBURE BIVISION.

03 FILLER

PIC X< 36)

VALUE

03400

INITIALIZE.

'H SERVICE

03410

OPEN INPUT TRANS ERHSG OUTPUT MASTR ELIST.

03 FILLER

PIC X < 32 >

VALUE SPACES.

03420

MOVE ZEROS TO J-DAY-BASE ERROR-KEY.

HEAI2.

03430

MOVE SPACES TO UORK-REC ERROR-TABLE U-LINE.

03 FILLER

PIC X ( 44 )

VALUE

03440

PERFORM ERROR-LOAD UNTIL END-OF-DATA.

ARTHROPOD-

BORNE ANIMAL DISEASE'.

03450

HOVE CURRENT-DATE TO 6ET-DATE.

03 FILLER

PIC X( 36)

VALUE

03460

GO TO STARTT.

' RESEARCH

LABORATORY

03470

•

03 FILLER

PIC X ( 32 )

VALUE SPACES.

03480

• ERROR-LOAD READS THE DECK OF ERROR MESSAGES AND LOADS THEM INTO

HEAD3.

03490

• ERROR-TABLE (CC79-80).

03 FILLER

PIC X( 44)

VALUE

03500

•

DENVER FEDERAL CENTER - D'.

03510

ERROR-LOAD.

03 FILLER

PIC X(28)

VALUE

03320

READ ERMSG AT END MOVE 'X' TO EOD.

'ENVER, COLORADO

03530

MOVE EH TO E-TIL (E-NO).

03 MM

PIC XX.

03340

•

03 FILLER

PIC X

VALUE '/'.

03550

• STARTT IS THE DRIVER OF THE PROGRAM.

03 DD

PIC XX.

03560

•

03 FILLER

PIC X

VALUE '/'.

03570

STARTT.

03 YY

PIC XX.

03580

HOVE ' TO EOD.

03 FILLER

PIC X(S2)

VALUE SPACES.

03590

PERFORM MAIN THRU HAIN-XIT UNTIL END-OF-DATA.

HEAD4.

03600

DISPLAY MASTER RECORDS REAB HRED.

03 FILLER

PIC X( 44)

VALUE

03610

DISPLAY MASTER RECORDS WRITTEN HROTE.

TRANSACTION / ERROR'.

03620

DISPLAT MASTER RECORDS WITH ERRORS HERR.

03 FILLER

PIC X(36)

VALUE

03630

CLOSE TRANS MASTR ELIST ERHSG.

' LIST INB

03640

STOP RUN.

03 FILLER

PIC X < 32 )

VALUE SPACES.

03650

•

HEADS.

03660

• MAIN IS THE BEGINNING OF THE MAJOR PERFORM STATEMENT. THE EDIT OF

03 FILLER

PIC X( 44 )

VALUE

03670

• EACH TRANSACTION RECORD STARTS HERE AND PROGRESSES THROUGH

' 1

2

3 4

03680

• MAIN-EXIT (STMT N 766). IT CONTAINS INDIVIDUAL FIELD VALIDITY

03 FILLER

PIC X ( 36)

VALUE

03690

• CHECKS, INTER-FIELD RELATIONAL EDITS AS WELL AS ERROR CHECK

' 3

6

7 8'.

03700

• AND PRINT ROUTINES.

03 FILLER

PIC X<52>

VALUE SPACES.

03710

•

HEAD6.

03720

MAIN.

03 FILLER

PIC X< 44)

VALUE

03730

READ TRANS INTO WORK-REC AT END HOVE 'X' TO EOD.

' 1 2345678901 2343678901 2343678901 2343678901 234' .

03740

IF END-OF-DATA GO TO MAIN-XIT.

03 FILLER

PIC X ( 36)

VALUE

03750

ABD 1 TO MRED.

'367D901 2345678901 2345678901234567890' .

03760

IF ZZ-COLONY GO TO FILE-CK.

03 FILLER

PIC X ( 32 )

VALUE SPACES.

03770

IF ID1 * PIDJH MOVE 'X' TO E-KEY (54). MOVE ID1 TO PIDJH. 7/7

ERROR-TABLE IS THE RECEIVING AREA FOR THE ERROR MESSAGES.

01 ERROR-TABLE.

03 ERR- T BL PIC 1(4320).

03 E-TIL REIEFINEB ERR- T BL OCCURS 54 TIMES.

03 E-M PIC X ( 78) .

03 E-N PIC XX.

ERROR-KEY HAS A 1 CHARACTER FLAG FOR EACH ERROR MESSAGE. AS AN
ERROR IS DETECTED, AN 'X' IS ROVED TO E-KEY (XX) WHERE XX IS
THE ERROR MESSAGE SUBSCRIPT. THEN, AS THE ERROR ROUTINE PROGRESSES
IT PRINT8 THE MESSAGE CORRESPONDING TO THE RELATIVE POSITION
WHERE AN 'X' IS FOUND.

01 ERROR-KET.

03 ERR-KET PIC X<34).

03 E-KEY REDEFINES ERR-KET PIC X OCCURS 54 TINES.

U-LINE HAS DATA ELEMENTS THAT CORRESPOND TO DATA ELEMENTS IN THE
TRANSACTION RECORD. WHEN A FIELD IS FOUND TO BE IN ERROR, ' '
(UNDERSCORES) ARE MOVED TO THE AREA CORRESPONDING TO THE FIELI
IN ERROR AND PRINTED AFTER THE RECORD 3UPPRES6ING LINE FEED TO
UNDERLINE THE DATA ELEMENT IN ERROR.

01 U-LINE.

03780

03790

03800

03810

03820

03830

03840

03850

03860

03870

03880

03890

03900

03910

03920

03930

03940

03950

03960

03970

03980

03990

04000

04010

04020

04030

MOVE UORK-REC TO DUPEH0LD1 .

IF RECH0LD1 NOT » RECH0LD2 MOVE DUPEH0LB1 TO DUPEH0LD2

ELSE MOVE X' TO E-KET (54) MOVE DUPEH01D1 TO

DUPEH0LD2.

FILE-CK VALIDATES CC 1.

X' TO E-KEY (01 ) MOVE

FILE-CK.

IF FILE-OK GO TO RECORD-CK ELSE HOVE

v T0

> RECORD-CK VALIDATES CC 2.

RECORD-CK.

IF RECORD-OK GO TO SER-CK ELSE MOVE 'X' TO E-KET (02) MOVE
v TO RT.

» SER-CK VALIDATES CC 3-4.

SER-CK.

IF SERIAL-CODE = ZZ' PERFORM ZZ-CHECK THRU ZZ-EXIT GO TO
RE C -PR T .

IF SER-CD-OK GO TO LRG-C6E-CK ELSE MOVE X TO E-KEY (03)
MOVE ALL V T0 sc*

i

► LRG-CGE-CK VALIDATES CC 5-8.

286

04040
04050
04060
04070
04080
04090
04100
04110
04120
04130
04 1 40
04130
04160
04170
04180
04190
04200
04210
04220
04230
04240
04230
04260
04270
04280
04290
04300
04310
04320
04330
04340
04350
04360
04370
04380
04390
04400
04410
04420
04430
04440
04430
04460
04470
04480
04490
04500
04510
04320
04530
04540
04550
04560
04570
04580
04390
04600
04610
04620
04630
04640
04650
04660
04670
04680
04690
04700
04710
04720
04730
04740
04750
04760
04770
04780
04790
04000
04810
04820
04830
04840
04850
04860
04870
04880
04890
04900
04910
04920
04930
04940
04950
04960
04970
04980
04990
05000
03010
03020
03030
03040
05030

LR6-C8E-CK .

IF SERIAL-CODE =* 'll' AND ZZ-CAGE GO TO SH-CGE-CK.

IF LARGE-CAQE-OK GO TO SH-CGE-CK ELSE HOVE 'X' TO E-KEY (04)

HOVE ALL V TO LC.

•

• SH-CGE-CK VALIDATES CC 9-11.

*

SH-COE-CK.

IF SHALL-CAGE-OK NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (03)

HOVE ALL V TO SHC.

• 8EN£RAT ION-CK VALIDATES CC 12-13.

•

GEMERAT ION-CK .

IF SERIAL-CODE “ AA' AND AA-6ENERATI0N 00 TO B-DATE-CK.

IF 6EHERAT ION-OK NEXT SENTENCE ELSE HOVE X' TO E-KET (06)

HOVE ALL TO 6N.

• B-DATE-CK VALIDATES BASE DATE CC 14-18.

•

B-DATE-CK.

IF DAY-366 AND LEAP-YEAR 08 TO EHER-S-CK.

IF YEAR-VALID NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (07) HOVE
ALL ™ BI.

IF B-TEAR > THI9-YEAR HOVE 'X' TO E-KEY (07) HOVE
ALL V TO DI.

IF DAY-VALID NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (08) HOVE
ALL V TO DD.

•

• EHER-S-CK VALIDATES CC 19-22.

EHER-S-CK.

IF AK-COLQHY AND AK-STRT-TIHE 60 TO EHER-E-CK.

IF START-T IHE-OK NEXT SENTENCE ELSE HOVE X' TO E-KEY (09)

HOVE ALL ' ' TO ES.

IF JB NOT NUHERIC HOVE 'X' TO E-KEY (09) HOVE ALL TO ES.

• EHER-E-CK VALIDATES CC 23-24 AND PERFORHS CALCULATIONS ON DAYS

• TILL EHER8ENCE.

•

ENER-E-CK.

H8VE B-DAY TO J-DAY-DA8E .

IF VALID-END-DATE NEXT SENTENCE ELSE HOVE ' X ' TO E-KEY (10)

HOVE ALL V TO EED 80 TO E-T-CK.

•THE FOLLOUIN8 5 LINES UERE ADDED ON 2/28/79 AFTER THE PR06RAN

• ABENDED AFTER F1NDIN6 A BLANK IN THE EHER-ENB FIELD

IF EHER-END NOT NUNERIC
DISPLAY UORK-REC
DISPLAY ' '

DISPLAY ' • ENER6ENCE END DAY (23-24) ILLE8AL AS RECORDED'

00 TO STARTT.

•

PERFORH VALIDATE VARYINO A FRQH 1 BY 1 UNTIL RE8ULT »

EHER-END.

IF A < 15 80 TO E-T-CK ELSE HOVE 'X' TO
E-KEY (11) HOVE ALL TO EED 80 TO E-T-CK.

o VALIDATE 18 THE PERFORHED COHPUTATION OF DATS TILL ENER8ENCE.

•

VALIDATE.

ADD 1 TO J-DAY-BASE.

IF LEAP-YEAR AND J-DAY-DA8E > 366
COHPUTE J-DAY-BASE ■ J-DAY-BASE - 366 ELSE
IF J-DAY-BASE > 365

COHPUTE J-DAY-BASE - J-DAY-BASE - 365.

•

• E-T-CK VALIDATES CC 25-28.

•

E-T-CK.

IF AK-COLONY AND AK-END-TIHE 60 TO FEED1-D-CK.

IF END-TIHE-OI NEXT SENTENCE ELSE NOVE ' X ' TO E-KEY (12) HOVE
ALL TO EET.

o

• FEEB1-D-CK VALIDATES CC 29-30, AND ITS RELATIONSHIP TO THE BASE DATE.

o

FEED1-D-CK.

IF VALID-FEED1 -DAY NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (46)

HOVE ALL V T0 F,» ™ FEED 1 -T-CK .

HOVE FEED1 -DAY TO DAY-STORE
HOVE B-DAY TO J-DAT-DASE.

IF DAY-STORE < RESULT HOVE 'X' TO E-KEY (13) HOVE ALL ' '

Tfl FID.

HOVE B-DAY TO HOLD.

IF FEED1 -DAY NOT » LST-2 HOVE ’X' TO E-KET (49) HOVE ALL ' '

TO FID.

i

• FEED1-T-CK VALIDATES CC 31-34.

o

FEED1-T-CK.

IF AK-COLONY AND AK- 1 ST-FEED-T INE 80 TO FOOD1 -T YPE-CK.

IF VALID- 1ST -FEED- TINE NEXT SENTENCE ELSE HOVE 'X' TO
E-KEY (14) HOVE ALL V ™ FIT.

>

• FOOD1 -TYPE-CK VALIDATES CC 35-36.

0

FOOD1 -TYPE-CK.

IF VALID-FOOB1 -TYPE NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (15)

HOVE ALL V 78 TF1.

i

• FEED2-D-CK VALIDATES CC 37-38.

05060

05070

05080

05090

05100

05110

05120

05130

05140

05130

05160

05170

05180

05190

03200

05210

03220

03230

03240

05230

05260

05270

03280

05290

03300

03310

03320

03330

05340

05350

05360

03370

03380

05390

05400

05410

05420

03430

05440

05450

03460

03470

03480

05690

05500

03510

05520

03530

03540

05550

03360

03370

03380

05390

03600

05610

05620

05630

05640

05650

03660

05670

03680

03690

03700

03710

05720

05736

05740

05750

05760

03770

05780

03790

05800

03810

03820

03830

03840

05850

05860

03876

05880

03890

03900

03910

03926

03930

05940

05950

05960

05970

05980

05990

06000

06010

06020

06036

06046

06030

06060

06070

FEED2-D-CK.

IF N0-2ND-FEED-DAY AND N0-2ND-FEED-TIHE GO TO FEEB3-D-CK.

IF VALID-FEED2-DAT NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (47)
HOVE ALL ' ' TO F2D 60 TO FEED2-T-CK.

• FEED2-T-CK VALIDATES CC 39-42 AS WELL AS ITS RELATIONSHIP TO CC 29-34.

FEED2-T-CK.

IF AK-COLONY AND AK-2ND-FEED-T I HE GO TO FOOD2-T YPE-CK .

IF VAL I D-2ND-FEED-TIHE NEXT SENTENCE ELSE HOVE X' TO E-KEY
(19) HOVE ALL V TO F2T .

IF FEEB1 -DAY - FEED2-DAY AND ( FEED2-TIHE NOT > FEED 1 - T I HE )
HOVE 'X' TO E-KEY (18) HOVE ALL V TO F2T .

• FOOD2-TYPE-CK VALIDATES CC 43-44 AS WELL AS ITS RELATIONSHIP TO

• CC 35-36.

FOOD2-TYPE-CK.

IF VAL ID-FOOD2-TTPE NEXT SENTENCE ELSE NOVE 'X' TO E-KEY (20)
HOVE ALL T0 7F2 80 T0 fE£D3-D-CK.

IF TYPE-F00D1 - TYPE-FOOD2 HOVE 'X' TO E-KEY (16) HOVE ALL
' ' TO TFl TF2.

• FEED3-D-CK VALIDATES CC 45-46.

•

FEED3-I-CK.

IF N0-3RD-FEED-DAY AND N0-3RD-FEED-TINE 00 TO ROON-CK.

IF VALIB-FEED3-DAY NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (48)
HOVE ALL V TO F3D 60 TO FEED3-T-CK.

• FEED3-T-CK VALIDATES CC 47-30 AS UELL AS ITS RELATIONSHIP TO

• CC 37-42.

•

FEED3- T-CK .

IF AK-COLONY AND AK-3RD-FEED- T INE 00 TO FOOD3-TYPE-CK.

IF VAL ID-3RD-FEEB-T INE NEXT SENTENCE EL8E HOVE 'X' TO
E-KEY (22) HOVE ALL TO F3T .

IF FEED2-DAY ■ FEED3-DAY AND (FEED3-TIHE NOT > FEED2-T IHE)
HOVE 'X' TO E-KEY (21) HOVE ALL TO F3T .

•

• FOOD3-TYPE-CK VALIDATES CC 51-32 AS UELL AS ITS RELATIONSHIP TO

• CC 43-44.

FOOD3-TYPE-CK.

IF VALID-FOOB3-TYPE NEXT SENTENCE EL8E HOVE 'X' TO E-KEY (23)
NOVE ALL V TO TF3 60 TO ROOH-CK.

IF TYPE-F00D2 - TYPE-FOOD3 HOVE 'X' TO E-KET (17) HOVE ALL

V T0 TF2 TF3-

• ROOH-CK VALIDATES CC 53-54.

ROOH-CK.

IF VAL ID-ROOH-NO NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (24)
HOVE ALL V TO RN.

• L-H-T-CK VALIDATES CC 55-37.

L-H-T-CK.

IF VALID-LARVAL-HI8H NEXT SENTENCE ELSE NOVE 'X' TO E-KEY
(25) HOVE ALL V TO LHT.

• L-L-T-CK VALIDATES CC 38-60 AS UELL AS ITS RELATIONSHIP TO CC 55-57.

IF VALID-LARVAL-LOU NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (26)

HOVE ALL ' ' TO LLT.

IF E-KEY (25) ■ 'X' OR E-KEY (26) - 'X' GO TO A-H-T-CK.

IF LARVAL-HI6H-TEHP < LARVAL-LOU-TEHP HOVE X' TO E-KEY (27)

HOVE ALL 70 W LLT.

*

• A-H-T-CK VALIDATES CC 61-62.

«

A-H-T-CK.

IF QOOB-H1GH-TEHP NEXT SENTENCE ELSE HOVE X' TO E-KET (28)

HOVE ALL TO AHT .

• A-L-T-CK VALIDATES CC 63-64 AS UELL AS ITS RELATIONSHIP TO CC 61-62.
A-L-T-CK.

IF GOOD-LOU-TEHP NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (29)

HOVE ALL 78 *LT.

IF E-KEY (20) - 'X OR E-KEY (29) • 'X' 60 TO A-H-H-CK.

IF ABULT-HI6H-TEHP < ADULT-LOU-TEHP HOVE 'X' TO E-KEY (30)

HOVE ALL V TO AHT ALT.

• A-H-H-CK VALIDATES CC 65-66.

•

A-H-H-CK.

IF VALID-HIGH-RH NEXT SEHTENCE ELSE HOVE 'X' TO E-KEY (31)

HOVE ALL 70 AHR-

«

• A-L-H-CK VALIDATES CC 67-68 AS UELL AS ITS RELATIONSHIP TO CC 65-66.

A-L-H-CK.

IF VAL ID-LOU-RH NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (32)
HOVE ALL TO ALR.

IF E-KEY (31) - 'X' OR E-KEY (32) * 'X' GO TO HETH-CK.

IF ADULT-HIOH-RH < ADULT-LOU-RH HOVE 'X' TO E-KEY (33) HOVE
ALL V TO AHR ALR.

•

• HETH-CK VALIDATES CC 69-73.

06000 HETH-CK.

06090 IF DOTH-ZEROS 60 TO DISPI-CK.

06100 IF VALID-C-HETMOD REIT SENTENCE ELSE HOVE 'X' TO E-KET <34 )

04110 HOVE ALL ' ' TO CR.

06120 IF ACCEPTARLE-RAHGE REIT SENTERCE ELSE HOVE 'X' TO E-KET (33)
06130 HOVE ALL V T0 *F •

06 140 •

06130 • DISPI-CK VALIDATES CC 74.

06160 •

06170 IISP1-CK.

06100 IF DISP1 -VALID NEXT SERTENCE ELSE HOVE 'X' TO E-KET (36) HOVE
06190 ALL TO D1.

06200 •

06210 • 11-IAT-CK VALIDATES CC 75-76.

06220 •

06230 D1-RAT-CK.

06240 IF VAL I D-D ISP 1 NEXT SERTENCE ELSE HOVE 'X' TO E-KET (37) HOVE

06250 ALL TO DID.

06260 •

06270 • DISP2-CK VALIDATES CC 77.

06280 •

06290 DISP2-CK.

06300 IF DISP2-VAL ID NEXT SERTENCE ELSE HOVE 'X' TO E-KET (30) HOVE
04310 ALL TO D2.

06320 •

06330 • D2-IAT-CK VALIDATES CC 70-79.

06340 •

06330 D2-DAT-CK.

06360 IF VALID-DISP2 NEXT SERTENCE ELSE HOVE 'X' TO E-KET (39) HOVE
06370 ALL *J TO D2D.

06300 •

06390 • C-TTPE-CK VALIDATES CC 00.

06400 •

06410 C-TTPE-CK.

06420 IF CARD-TYPE-OK REIT SENTERCE ELSE HOVE 'l' TO E-KET (41)

06430 HOVE ALL TO CT.

06440 •

06450 • CR088-CK IB THE IE8IRNIN6 OF THE HAJOR CROSS-FIELD EDITS VALIDATING
06460 • RELATIONSHIPS IETUEEN FIELDS IR THE TRANSACTION RECORDS. THIS
06470 * CHECK RUNS THRU CONTIHUEI AND TERHINATES THE STANDARD
06480 • TRANSACTION EDITS.

06490 •

06300 CR08S-CK.

06510 IF (RECORD-TTPE ■ ' 1 ' OR '2') AND (FILE-TTPE ■ 'V OR '2' OR

06520 '8') NEXT SERTENCE ELSE HOVE 'X' TO E-KET (43) HOVE ALL

06530 TO FT RT.

06540 IF AK-COLONT AND (ROOH-NO ■ '03' OR '09') HOVE 'X' TO
04330 E-KET (44) HOVE ALL TO SC RN.

06560 IF HETHOD-RO-FLIES AND NO-FLIES GO TO CONTINUE.

06370 IF RO-FLIES AND NOT HETH0D-N0-FLIE8 HOVE 'X' TO E-KET (45)

06380 HOVE ALL TO CR NF.

06390 CONTINUE.

06600 IF ADULTS-TO-BOTH1 AND NOT ADULTS-TO-BOTH2 HOVE 'X' TO E-KET

06610 (50) HOVE ALL '_' TO D1 D2.

06420 IF ADULTS-TO-BOTH2 AND NOT ABULTS-TO-BOTH1 HOVE 'X' TO

04630 E-KET (30) HOVE ALL TO Dl D2.

06640 IF BLANK-VALUE AND NOT GOOD-U-DLANK HOVE 'X' TO E-KET (31)

06450 HOVE ALL TO Dl D2.

06660 IF DISPOSITION! > ' ' NEXT SENTENCE ELSE 60 TO CONTINUE! .

06670 IF DISPOSITIOR2 - ' ' OR '1' OR '5' NEXT SENTENCE ELBE HOVE

06400 'X' TO E-KET (40) HOVE '_' TO Dl D2.

06690 CONTIHUEI.

06700

04710

IF AIULTS-TO-IOTH2 AND DISP2-IAY < DISPI-DAY HOVE '
E-KEY (52) HOVE ALL TO DID D2D.

X'

TO

04720

04730

IF SERIAL-CODE = 'AA
E-KEY (53) ROVE ALL '

AND BASE-BATE
.' TO SC BI BD.

'OOOOO' HOVE

'X'

' TO

06740

04730

IF SERIAL-CODE * 'AK'
E-KEY (53) HOVE ALL '

AND DASE-DATE
TO SC BI BD.

<

'73009' HOVE

'X'

’ TO

06760

06770

IF SERIAL-CODE - 'AH'
E-KET (33) HOVE ALL '

AND DASE-DATE
J TO SC Dl DD.

<

'74275' HOVE

'X'

' TO

06700

04790

IF SERIAL-CODE - 'AB
E-KEY (33) HOVE ALL '

AND DASE-DATE
TO SC BI BD.

<

'73157' HOVE

'X'

TO

06000

06810

IF SERIAL-CODE - 'AC'
E-KEY (53) HOVE ALL '

AND DASE-DATE
_' TO SC BI ID.

<

'73268' HOVE

'X'

TO

06020

06830

IF SERIAL-CODE • AD
E-KEY (33) HOVE ALL '

AND DASE-DATE
TO SC Dl BD.

<

'73280' HOVE

'X'

TO

06840

04850

IF SERIAL-CODE ■ 'AE'
E-KEY (53) HOVE ALL '

AND DASE-DATE
_' TO SC BI BD.

<

'73158' HOVE

'X'

TO

06660

06870

IF SERIAL-CODE - 'AF'
E-KEY (53) HOVE ALL '

AND BASE-DATE
J TO SC BI BB.

<

'72010' HOVE

'X-

' TO

06600

06890

IF SERIAL-CODE - 'AG'
E-KEY (53) HOVE ALL '

AND BASE-DATE
TO SC BI BD.

<

'72009' HOVE

'X

’ TO

06900

06910

IF SERIAL-CODE =» 'AT
E-KEY (53) HOVE ALL '

AND DASE-DATE
TO SC BI DD.

<

'74273' HOVE

'X

' TO

06920

06930

IF SERIAL-CODE - 'AJ'
E-KEY (33) HOVE ALL '

AND DASE-DATE
TO SC BI BD.

<

'74214' HOVE

'X

TO

06940

04930

IF SERIAL-CODE » 'AL'
E-KEY (33) HOVE ALL '

AND DASE-DATE
TO SC BI DD.

<

'75142' HOVE

'X

' TO

06960

06970

IF SERIAL-CODE - 'AH'
E-KEY (33) HOVE ALL

AND BASE-DATE
TO SC Dl BD.

<

'75142' HOVE

'X

' TO

06980

06990

IF SERIAL-CODE - 'AN'
E-KEY (33) HOVE ALL '

AND DASE-DATE
TO SC Dl BD.

<

'73142' HOVE

'X

' TO

07000

07010

IF SERIAL-CODE • 'AO'
E-KEY (33) HOVE ALL '

AND DASE-DATE
TO SC BI BD.

<

'75142' HOVE

'X

' TO

07020

07030

IF SERIAL-CODE - 'AP'
E-KEY (53) HOVE ALL '

AND DASE-DATE
' TO SC BI BD.

<

'73256' HOVE

'X

' TO

07040 •

07030 • REC-PRT . IN THIS PARAGRAPH, RECORDS UITH ERRORS ARE IDENTIFIED
07060 • AND PRINTED ALONG UITH U-LIHE UHICH UNDERLINES THE BAD FIELDS.
07070 •

07000 REC-PRT.

07090 IF LINEZ > 50 PERFORH HDR-PRT .

07100 IF ERR-KET ■ ALL '0' GO TO ALHOST-DONE.

07110 HOVE UORK-REC TO IODT URITE PRT AFTER 2 ADD 3 TO LINEZ.

07120 HOVE ' ' TO HD-SM.

07130 ADD 1 TO HROTE .

07140 HOVE U-LINE TO BODY URITE PRT AFTER NONE.

07150 HOVE SPACES TO BODY URITE PRT AFTER 1.

07160 ADD 1 TO HERR.

07170 IF HERR > 5689 DISPLAY 'TOO HANY ERRORS, ABORTING ' HOVE CLH07208
07180 'X' TO EOD.

07190 *

07200 • ERROR-LOOP CONTROLS THE PERFORHANCE OF THE ERROR SEARCH AND
07210 • PRINT PROCEDURE.

07220 •

07230 ERROR-LOOP.

07240 PERFORH KEY-CHECK VARY IN6 D FROH 1 BY 1 UNTIL B > 54.

07250 HOVE ALL ZEROS TO ERR-KET.

07260 HOVE ' ' TO HD-SU.

07270 •

07280 • KEY-CHECK CHECKS EACH OCCURARCE OF E-KET (XX) LOOKING FOR A VALUE
07290 • ' X ' AND PRINTING ERROR HESSA6E (XX) WHEN ENCOUNTERED.

07300 •

07310 KET-CHECK.

07320 IF LINEZ > 50 PERFORH HDR-PRT HOVE X' TO HD-SU.

07330 IF HD-SU • 'X' HOVE SPACES TOf BODY URITE PRT AFTER 1 ADD 1

07340 TO LINEZ HOVE ' ' TO HD-SU.

07350 IF E-KET (B) ■ 'X' HOVE E-H (B) TO BODY URITE PRT AFTER 1

07360 ADD 1 TO LINEZ.

07370 HOVE SPACES TO BODY.

07380 *

07390 • ALHOST-DORE. HERE RECORDS THAT ARE FOUND TO BE ERROR FREE ARE
07400 • WRITTEN TO THE HASTER FILE.

07410 •

07420 ALHOST-DONE.

07430 HOVE ZEROS TO J-DAY-BASE ERROR-KEY.

07440 HOVE SPACES TO UORK-REC U-LIHE.

07450 GO TO HAIN-XIT.

07460 •

07470 * ZZ-CHECK. IN THE CASE OF ZZ COLONY BATA, ONLY PART OF THE EDITS
07480 ♦ ARE PERFORHED. THOSE ZZ RECORDS ARE PROCESSED THRU ZZ-EXIT AND

07490 • THEN PASSED TO THE ERROR PRINT PORTION OF HAIN .

07500 •

07310 ZZ-CHECK.

07520 IF DAT-366 AND LEAP-TEAR GO TO ZZ-A.

07330 IF YEAR-VALID NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (07) HOVE

07540 ALL '_' TO BI.

07550 IF DAT-VALID NEXT SENTENCE ELSE HOVE 'X' TO E-KET (00) HOVE

07360 ALL TO BD.

07570 ZZ-A.

07500 IF VALID-ROOH-NO NEXT SENTENCE ELSE HOVE 'X' TO E-KET (24)

07390 HOVE ALL ' ' TO RN.

07600 IF VAL ID-LARVAL- HI 6H NEXT SENTENCE ELSE HOVE 'X' TO E-KEY

07610 (23) HOVE ALL TO LHT.

07620 IF VAL ID-LARVAL-LOU NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (26)

07630 HOVE ALL TO LLT .

07640 IF E-KEY (25) » 'X' OR E-KEY (26) - X' GO TO ZZ-B.

07650 IF LARVAL-HIGH-TEHP < LARVAL-LOU-TEHP HOVE 'X' TO E-KET (27)

07660 HOVE ALL TO LHT LLT.

07670 ZZ-D.

07600 PERFORH A-H-T-CK.

07690 IF OOOD-LOU-TEHP NEXT SENTENCE ELSE HOVE 'X' TO E-KEY (29)

07700 HOVE ALL TO ALT.

07710 IF E-KEY (20) « X' OR E-KEY (29) * 'X' 00 TO ZZ-C.

07720 IF ADULT-HIGH-TEHP < ADULT-LOU-TEHP HOVE 'X' TO E-KEY (30)

07730 HOVE ALL '_' TO AHT ALT.

07740 ZZ-C.

07750 PERFORH A-H-H-CK.

07760 IF VALID-LOU-RH NEXT SENTENCE ELSE HOVE 'X' TO E-KET (32)

07770 HOVE ALL TO ALR.

07700 IF E-KEY (31) ■ 'X' OR E-KEY (32) = 'X' GO TO ZZ-EXIT.

07790 IF ADULT-HI6H-RH < ADULT-LOU-RH HOVE 'X' TO E-KEY (33) HOVE

07800 ALL ' ' TO AHR ALR.

07810 ZZ-EXIT.

07820 EXIT.

07830 HAIN-XIT.

07840 EXIT.

07050 •

07860 • HDR-PRT IS THE PERFORHED HEADER PRINT ROUTINE.

07070 •

07800 HDR-PRT.

07090

HOVE

HEAD1

TO

DODY

URITE

PRT

AFTER

TOPLINE

07900

HOVE

HEAD2

TO

BODY

URITE

PRT

AFTER

1.

07910

HOVE

HEAD3

TO

BODY

URITE

PRT

AFTER

1 .

07920

HOVE

HEAD4

TO

DODY

URITE

PRT

AFTER

1 .

07930

HOVE

HEADS

TO

DODY

URITE

PRT

AFTER

2.

07940

HOVE

HEAD6

TO

BODY

URITE

PRT

AFTER

1 .

07950

HOVE

07 TO

LINEZ.

READY

288

Appendix C.— Summary of the Retrieval-Request Document Based on Data Entered in the Record Document (Fig. 3) and
Codes in Appendix A

Information requested

1. Pupal production (69-73).6

2. Adult holding temperatures and relative humidities for a speci¬
fic colony or room (61-68).

3. Larval rearing-temperatures for a specific colony or room
(55-60). 7

4. Larval rearing-temperature history for a specific cage of adult
insects (55-60). 8

5. Data listing by date for a specific colony (1-80).

6. Data listing for a specific time interval for all colonies (1-80).

7. Data listing for a group of large cages for a specific colony
(1-80).

8. Use of flies (74-79).®

9. Listing of cages of flies outside a specific temperature range
(5-8, 9-11).

10. Listing of cages for a specific colony that received a specific
number of feedings for a specific time interval (5-8, 9- 11). 10

11. Universal retrieval program (any of the data in 1-80 by re¬
quest).

Data input required

Julian dates (as coded in columns 14-18) for the time interval
(time span). Colony code (as coded in columns 3-4).

Time unit6— l=day, 2=week, 3=month, 4=quarter, and
5=year.

Plot— l=yes, 2=no.

Julian dates for the time interval.

Colony code and room-number code (as coded in columns 53-
54) number code only.

Time unit (as above).

Julian dates for the time interval.

Colony code and room-number code or room-number code only.

Colony code.

Large-cage number or large-cage number and subcage number,
(as coded in columns 5-8 and 9-11).

Number of days required for larval development.

Julian dates for the time interval.

Colony code.

Julian date for the time interval.

Ranges of large-cage numbers.

Colony code.

Julian dates for the time interval.

Colony code.

Time unit (as above).

Julian dates for the time interval.

Colony code.

Temperature range (as coded in columns 55-64)— l=larvae
(a=high, b=low), 2=adult (a=high, b=low).

Number of days required for larval development.

Julian dates for the time interval.

Colony code.

Number of feedings— 1 = 1, 2 = 2, 3=3.

Objective— provide retrieval of data with the fewest restric¬
tions possible imposed by the program, but allow users
to restrict the data listed as much as they wish by specify¬
ing codes, for particular fields, that must be present for data
to be listed.

Input— user provides codes or variables acceptable by the cur¬
rent editing program for any of the fields of the colony-
production record (fig. 3). The inputs of the computer pro¬
gram correspond to the fields shown in figure 3. As the user

“See figure 5 for sample output.

6Time unit must be less than or equal to time interval.
7See figure 6(A) for sample output.

8See figure 6(B) for sample output.

“See figure 7 (A and B) for sample output.
‘“See figure 8(A) for sample output.

289

Information requested

Data input required

enters codes in more fields, the listing becomes more re¬
stricted (for example, if AA were entered in columns 3 and
4— colony— then the data listing would be only for that
colony but would include all data associated with the AA
code; if AA were entered in columns 3 and 4, and if the num¬
bers 3398-4240 were entered in columns 5-8— large-cage
numbers— then the listing would include all data associated
with the AA colony for large cages numbered from 3,398 to
4,240). Also, the computer program accepts inputs of two or
more codes or variables or a range of codes or variables for
the same field. In such cases, the data will be listed for all
codes or variables and/or their ranges, that have been en¬
tered as inputs.

12. Quality control of size for a specific colony; to list the Julian
date of the quality-control sample, the number of pupae per
milliliter, mean wing length (mm) ± S.E. (standard error),
and mean dry weight (pg)±S.E.“

Julian dates for the time interval.

Colony code.

Time unit (as above).

Listing by Julian date— l=yes, 2=no.

Listing by large-cage number— l=yes, 2=no. If yes, enter
large-cage number and/or range of numbers.

Plot and printout of primary and secondary polynomial regres¬
sion equation of dry weight (X-axis) times wing length ( Y-
axis)— l=yes, 2=no.

Plot and printout of primary and secondary polynomial re¬
gression equation of dry weight (X-axis) times the number
of pupae per milliliter (Y-axis)— Culicoides only; l=yes,
2=no.

“See figure 8(B) for sample output.

290

Appendix D.— Computer Programs Used to Operate
the Information Retrieval System

Program

Contents or programs

CLIST

S FORMAT .

ST AT .

STATBIG .

CNTL

DISKTAPE .

TAPEDISK .

LOADSTAT

EXEC. COLVERIF

LOADEXEC. COLVERIF

LOAD MODULES
COLPROD
FORMAT

SOURCE PROGRAMS

COLVERIF .

S FORMAT .

STATPGMS. A .

STATPGMS. B
DATA FILES
ERRMSGS

NEW. COLONY

TAD 17 .

Creates SAS-format module used by statistical analysis programs.
Executes statistical-analysis programs.

Executes large statistical-analysis programs.

Copies colony-production data base from disk to tape.

Copies colony-production data base from tape to disk.

Copies statistical programs, format module, and CLIST programs
from tape to disk.

Executes colony-production editing program and error-message file
from disk.

Executes colony-production editing program and error-message file
from tape.

Contains editing program COLVERIF.

Contains SAS-format program S FORMAT.

Colony production editing program.

Format program for SAS statistical-analysis programs.
Statistical-analysis programs (see appendix C, 1-4 and 12).
Statistical-analysis programs (see appendix C, 6-12).

File of the texts of error messages printed out by colony-production
editing program.

Colony-production data for 1978.

Colony-production data for 1974-77.

Systems Management of Insect-
Population-Suppression Programs
Based on Mass-Production
of Biological-Control Organisms

By N. C. Leppla1

Introduction

Effective management defines and organizes the
variables that comprise a functional system and uses the
system to maintain itself. Having discovered that a finely
tuned organization will almost run itself, successful
managers have a nearly intuitive sense of order. So,
rather than managing hectically and anxiously from crisis
to crisis, they have become professionals who anticipate
and almost relish most administrative challenges.

One such challenge is to develop new technologies that
can be integrated with existing ones for reducing the im¬
pact of agricultural pests. The result is IPM (integrated
pest management), which includes chemical control, plant
and animal resistance, cultural practices, and biological
control (parasites, predators, pathogens, sterile-insect
technique, and genetic manipulation). Most of these
techniques depend on having large numbers of
laboratory-reared insects for their development and im¬
plementation. It would be wonderful if we could simply
collect some insects, rear and sterilize them, spray them
back into the field, and watch the pest population disap¬
pear. Unfortunately, there are no insecticidelike “magic
bullets.” No single set of rules and procedures for manag¬
ing population-suppression programs based on mass-
produced organisms exists; nor should it, since the
contributing elements can be fitted together in various
ways to solve different problems. This paper describes
elements of a generalized system for rearing and utilizing
insects that can be used as a framework for various pro¬
grams with various needs and purposes.

System Components

This conference has documented recent advances and ex¬
isting capabilities for handling the primary elements of
insect-rearing and utilization programs. It has also em¬
phasized the importance of organizing these elements in¬
to functional systems that can be managed effectively.

‘Research entomologist. Insect Attractants, Behavior, and
Basic Biology Research Laboratory, Agricultural Research Serv¬
ice, U.S. Department of Agriculture, P.O. Box 14565,

Gainesville, Fla. 32604

This approach of generating and operating unified
systems rather than dividing them into smaller, osten¬
sibly more manageable parts is a major technological
revolution (Kavanau et al. 1971). Such a system is more
than an aggregation of interrelated elements; it has a life
of its own so that it reacts as a unit to adjustments in its
parts. The system’s hierarchal structure can provide
several operational levels. Also, because the system re¬
quires cooperative effort, it may become the means for
accommodating divergent attitudes, conflicting motives,
and dissimilar ideologies.

The first step in developing an integral system is to ar¬
range the existing subsystems into a functional pattern.
For an insect-population-suppression program, the
primary functions are operation, quality control, and
perpetuation. “Operation” includes colonization, produc¬
tion, and utilization, the subsystems that directly con¬
tribute to organizational output. “Colonization” is
primarily strain development; production and utilization
emphasize process engineering. “Quality control” is the
monitoring and evaluation of insectary products and their
use, and it affects all phases of the program. “Perpetua¬
tion” includes all the elements of effective management
that facilitate interactions among the subsystems. These
subsystems and their elements are characterized by
relative permanence; but the operations, procedures, and
associated facilities and equipment vary as the system
evolves.

Operation

Colonization.— The first element of the colonization sub¬
system is characterization of the target population. The
pest’s geographical and temporal distributions, host
preferences, life history, etc., are determined by conduct¬
ing ecological surveys. The basis for a strategy for
behavioral or genetic control is derived from these data.

If, for example, the object is to disrupt reproduction, all
pertinent prereproductive and postreproductive processes
must be understood. Once the pest has been character¬
ized, the laboratory colony can be established with insect
collections of the appropriate composition. Desired
genetic traits are preserved by relaxing environmental
stress within the insectary and by adopting a suitable
maintenance strategy such as perpetual isolation,
hybridization, periodic infusion, or replacement. The

292

characterized insect becomes the biological standard
for measuring the effects of production and utilization
operations.

Production. — The elements of production are rearing,
engineering, and management. “Rearing” includes ac¬
quisition and storage of materials; diet formulation and
preparation; egg collection, treatment, and placement in
containers; larval development; removal and distribution
of pupae; adult maintenance; and microbial control.
“Engineering” encompasses facilities, equipment, in¬
strumentation (monitoring and control of processes and
environments), and maintenance. “Management” op¬
timizes, organizes, and allocates all available resources to
sustain the colony.

Utilization.— “Utilization” is the treatment, transport,
and deployment of population-suppression organisms.
Treatment may involve sterilization by irradiation,
chemicals, genetic alteration, or physical stress; also, it
usually includes some kind of marking (dye, isotopes,
etc.) and preconditioning (chilling, photoperiodic entrain¬
ment, acclimation, etc.). Obviously, the insects must be
transported quickly and under the best physiological con¬
ditions, an operation that is technically simple but often
difficult to accomplish in practice. “Deployment” is the
development and application of procedures and equip¬
ment for insect release. Like production, utilization
requires an integration of scientific and engineering
expertise (for example, see Smith 1977).

Quality control

“Quality control” is the monitoring of colonization, pro¬
duction, and utilization operations and their effects on
the products. Monitoring techniques are developed and
implemented, the data are periodically evaluated, and
recommendations are made for improving the system.
But, once tests and standards have been established,
routine measurements are made as procedures directly in¬
volved with performing an operation. So quality control
works like a servomechanism. A solid-state temperature
controller, for example, monitors the ambient air with
sensors, evaluates changes in electrical current within its
circuits, and actuates devices to heat or cool the air until
conditions stabilize. Quality control operates similarly to
maintain desired conditions; but, since it provides
tolerances and standards, it is more flexible than a single
regulatory device. It actually coordinates the operations
and insures the maintenance of acceptable standards of
colonization, production, and utilization.

Perpetuation

The ultimate goal of program management is to establish
a dynamic system that eliminates any long-term

dependence on individual managers and builds the
organization around program goals. An insect-population-
suppression program, like any business corporation,
should be created to preserve the organization’s conti¬
nuity and extend its capabilities beyond human limits.
Management must therefore think in terms of what
operations are needed and what procedures must be
followed rather than available skills and training. Certain¬
ly, key roles will emerge and be assumed by responsible
people, but performance and productivity should never be
limited by the limitations of the personnel.

If we assume that those associated most closely with an
operation know most about it, then, clearly, subsystems
of an insect-population-suppression program should be
managed internally. But, participatory management is
needed for the refinement and perpetuation of the sub¬
systems (King 1975). People who concentrate on im¬
plementing one set of operations may not be aware of
technologies that could improve their effectiveness and
are usually unable to see their relative importance. So it
is advantageous to cultivate an interdependent partner¬
ship and integrate available expertise to solve mutual
problems. Management provides this capability to
achieve this integration through organization, support,
communication, and regulation.

Invariably, successful managers are supportive. They not
only allocate resources where needed, but they also at¬
tempt to eliminate obstacles to employee success such as
poor work environments (ones that are hazardous, inflexi¬
ble, or constantly interrupted), deficiences in training or
experience, or inadequate coordination. Most employees
will achieve their potential if such obstacles are moder¬
ated or removed and if they are given a little motiva¬
tion in the form of material and emotional satisfaction.

Management is sometimes characterized as a sinister ac¬
tivity, wholly preoccupied with the formulation and en¬
forcement of regulations. Indeed, some antiquated
managers still consider this to be their primary function.
But enlightened managers know that the object of an
organized effort is productivity not regulation. They con¬
centrate on defining and measuring productivity and
regulating only those activities needed to acnieve a
suitable quantity and quality of work. In the case of
pupal irradiation, for example, management is concerned
with the dosage and number of sterilized pupae and not
with maintenance schedules, time cards, dress codes, etc.,
of the employees. Guidelines should specify output and
only those personal habits that directly affect it. This ap¬
proach maximizes the flexibility necessary for handling
contingencies and preserving employee autonomy, digni¬
ty, and professionalism.

Effective communication is the key to accomplishing

293

organizational, supportive, and regulatory goals because
it links subsystem elements, provides a way to incor¬
porate new technology, and facilitates the enforcement of
policies. It is the means by which deficiencies are cor¬
rected and achievements recognized. For communication
to be effective, people who work together should have fre¬
quent informal meetings; staff meetings should be held
less often but according to an appropriate fixed schedule.
Employees should always understand their roles in the
organization and never doubt the importance of their
contributions.

Factors Complicating
Systems Management

Population-suppression programs that depend on the effi¬
cient production and utilization of biological-control
organisms often appear unmanageable because they are
inherently complex, relying on technology that is derived
from many interrelated disciplines. Since coordinating
these disciplines is difficult, the field has remained un¬
necessarily compartmentalized. For example, it is impos¬
sible to deal with diets and containerization without
considering microbial contamination, and biological and
engineering problems are inseparable when rearing, trans¬
port, and release are automated. So management and
quality control are necessary to every aspect of a
program.

Another problem that hinders progress and demands
management coordination is the divergence of specialized
fields. Scientists, engineers, administrators, educators,
merchants, and other specialists may agree that the pur¬
pose of insect rearing is to provide a dependable supply
of organisms that meet acceptable biological standards,
but they have somewhat different attitudes and priorities
concerning the allocation of resources. The entomologist,
for example, is inclined to accept the dynamic and em¬
pirical processes that dictate relatively frequent changes
in materials and equipment, but the administrator may
question the expense, and the engineer might actually
propose that the insects be changed to save the hard¬
ware! These specialists are not inflexible, but they func¬
tion in different contexts whose compatibility they do not
always recognize. So they must be linked by effective
management. (For a worthwhile discussion of the role of
engineering in developing new technology, see Bailey
1978.)

Other problems management will have to deal with in¬
clude agricultural practices incompatible with biological-
control strategies, national and international politics,
long-term financial support, and imbalances in the pro¬
portions of basic research and dependent technology.
These considerations and others have somehow dictated
that massive programs be controlled by a few managers
and thus become limited in scope by their visions and
talents. But the most appropriate way to plan, operate,
analyze, and perpetuate these complex programs is to
adapt the concepts and principles of systems manage¬
ment. Systems management concentrates critical eval¬
uation on the program rather than on its individual
elements. The details can be organized and controlled by
means of modern computer technology, and the system
can serve as its own model for testing creative ideas
without disrupting existing operations and making costly
mistakes. Also, cooperation and interdependence are pro¬
moted through effective coordination. Management must
unify the complex contributing elements and provide
insect-population-suppression programs that will meet
agriculture’s future challenges. (For a humorous but
pointed discussion of management, see Ettinger 1970.)

References

Bailey, R. L.

1978. Disciplined creativity for engineers. 614 pp.

Ann Arbor Science Publishers, Ann Arbor,
Mich.

Ettinger, M. B.

1970. Standard methods for the mediocre science
manager. 86 pp. Ann Arbor Science Publishers,
Ann Arbor, Mich.

Kavanau, L. L.; Ancker, C. J., Jr.; and Schneidewind, N.

F.

1971. Systems analysis, design, and operation pro¬
cedures. In H. B. Maynard (ed.), Industrial En¬
gineering Handbook, pp. 8-3—8-33. McGraw-
Hill, New York.

King, J. R.

1975. Production planning and control. An introduc¬
tion to quantitative methods. 403 pp. Per-
gamon Press, New York.

Smith, D. B.

1977. Entomology-engineering cooperation— where do
we go from here? Bull. Entomol. Soc. Am.

23: 120-123.

294

The Insectary Manager

By W. R. Fisher1

Introduction

Rearing animals in the laboratory presents challenges
and problems unlike those associated with the production
of inanimate objects. This is especially true with the pro¬
duction of insects. First, each species goes through
distinct developmental stages that have different life
styles and different requirements for temperature, rela¬
tive humidity, lighting, diet, population density, contain¬
ment, and sanitation. So the environment in the rearing
facility must cater to the needs of each stage, particularly
immatures, whose needs are often radically different from
those of the adult. Second, insects in culture must be pro¬
tected against pathogens, parasites, and predators, which
can affect quality, reduce yields, and disrupt scheduling.
And a cannibalistic species must be protected from itself.
Third, during the course of production, insects must be
handled when, for example, they are placed on diet,
moved to a different container, or surface-sterilized. This
handling must be done carefully because eggs and pupae
are easily smashed, and soft-bodied larvae can be easily
punctured. Fourth, insect populations are characterized
by an element of inherent variability that can manifest
itself, regardless of environmental constancy, as subtle
differences in morphology, developmental rates, and
behavior. The type and degree of variation depend on fac¬
tors such as the stage of colonization and the time of
year. So identification of significant inconsistencies in
insect quality becomes difficult. Also, variability is in¬
creased by many elements in the rearing program itself,
including handling of insects and the types of diet. Cor¬
relating anomalies with specific causes and correcting for
them is therefore a complex task. So, the challenges and
problems associated with insect-production programs pro¬
vide a unique management opportunity that requires
trained, experienced individuals to establish and maintain
a quality production program.

Training and Experience

Two factors necessitate a higher level of training for
future insectary managers: the need for more efficient
production of quality insects and the increasing recogni¬
tion that manipulation of variables in the rearing pro-

‘Graduate research assistant, Department of Entomology and
Nematology, University of Florida; employed through a coop¬
erative agreement with the Insect Attractants, Behavior, and
Basic Biology Research Laboratory, Agricultural Research Serv¬
ice, U.S. Department of Agriculture, Gainesville, Fla. 32604.

gram will be needed to enable adjustment of that quality
to satisfy the specific needs of each release program. The
insectary manager should have academic training in
disciplines that are basic to the major areas of insect
production, such as the insects, their environment, and
insectary personnel. Courses in general entomology, in¬
sect physiology, pathology, and behavior provide an
understanding of insect life cycles, development, mor¬
phology, and diagnosis and treatment of disease, and
they form the basis for evaluating production efficiency
and insect quality. Courses dealing with insect ecology,
general microbiology, and general nutrition will illustrate
the effects of variables such as temperature, relative
humidity, and population density on insect growth and
development. They will also demonstrate microbiological
concepts, sanitary techniques, and nutritional require¬
ments of living organisms. A course covering main¬
tenance of environmental conditions will provide an
understanding of heating, cooling, and humidification
systems and how they are controlled and monitored.
Lastly, a basic course in personnel management will help
the manager in scheduling activities and organizing avail¬
able labor.

Formal coursework in understanding basic concepts is
valuable, but practical experience and training remain the
primary means of developing expertise in insect rearing.
Firsthand observations allow the manager to get a feel
for the program. By handling insects, performing proce¬
dures, and identifying problem areas, the manager is bet¬
ter able to evaluate the performance of the facility, the
equipment, and the personnel. For example, one can read
the recipe of ingredients and amounts used to prepare an
artificial diet, but one must observe the actual blending
action, texture, and gelling characteristics of the mixture
to be able to decide how to insure consistent batches.
Likewise, the manager must participate in the program
directly for a considerable length of time to gain the
knowledge needed for making decisions about subjective
matters such as the cause of abnormal fluctuations in
yield or insect quality or the course of action to take in
response to an outbreak of disease or dietary contamina¬
tion. With this experience, the manager should develop
an intuitive sense about the operations necessary for the
most appropriate response at the most advantageous
time. An inexperienced manager might overreact to an
apparent problem, making unnecessary or premature ad¬
justments in the program.

Responsibilities

The insectary manager must be an active participant in
the rearing program. He must know production require-

295

ments, as well as the responsibilities and activities of all
employees. He must know how to perform procedures
and how to use equipment. The manager must also insure
that all items needed for successful and safe completion
of all tasks are available. These include lab coats, gloves,
data books, stationary supplies, etc. Ordering any sup¬
plies, especially dietary ingredients or containers, should
be done by the manager well in advance of their use to
avoid disruption of the program because of unexpected
delays in availability or shipping.

Often, the items needed for successful rearing are un¬
available from suppliers and must be custom-built to
specifications based on the biology of the insect and
other considerations. For instance, one manager recog¬
nized that a cannibalistic species needed to be reared in
individual containers. At the same time, she realized that
this procedure depended on expensive, nonreusable plastic
cups and a labor-intensive effort to pour diet, plant eggs,
cap cups, and harvest pupae. She investigated a more effi¬
cient, reusable, multicellular system that would be a
significant savings in labor and money. Reports in¬
dicated, however, that these units were made of brittle
polystyrene plastic with square-cornered cells that pro¬
hibited adequate cleaning before reuse. The cell units
were heat labile and could not be sterilized in a steam
autoclave. Disinfection in bleach solution did not ade¬
quately destroy contaminants. This inadequate disinfec¬
tion was of major concern to the manager because she
wanted to reduce the incidence of disease organisms and
dietary contamination. She concluded that a new material
was needed that would be inexpensive and able to with¬
stand sterilization in an autoclave. Individual cells should
be round and seamless to aid cleaning and should be
large enough to contain enough diet for development and
pupation but small enough for efficient production.
Material covering the top of the cell unit should allow
adequate gas exchange for maintaining the necessary mi¬
croclimate in cells, and, at the same time, it must be of
small pore size to keep neonates from escaping. It must
also be rugged enough to withstand the damage from
mandibles of late-instar larvae. The final product con¬
sisted of liquid resin poured into a reusable, flexible mold
that was peeled away after the resin had hardened. The
covering material was porous polypropylene, commonly
used for containing insects. Both these materials are
autoclavable. This example illustrates the many variables
that must often be considered by the manager when mak¬
ing decisions about the rearing program.

reduce the occurrence of problems such as the spread of
disease throughout the facility. For example, unnecessary
personnel must be restricted from sanitary areas, and the
restriction must be enforced without exception. Even the
best rearing program cannot compensate for valid rules
that are not enforced. This example is an illustration of
the kind of procedures the manager should follow to solve
problems by treating their causes and not merely by
treating their symptoms. It is, in this illustration, more
advantageous to prevent disease by restricting the move¬
ment of personnel than it is to treat disease symptoms
by adding to the diet antimicrobial chemicals that may
affect insect quality.

Problems may develop as a result of equipment failure.
Knowing what to do in such an emergency is often dif¬
ficult unless prior thought has been given to potential
causes and alternative courses of action. Figure 1 is an
example of such a plan for dealing with potential prob¬
lems. In this dichotomously-branching flow chart, a prob¬
lem with the air-conditioning system causes an alarm to
be sounded. The manager follows the chart step-by-step

i

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Beyond the relatively obvious duties, one of the
manager’s primary responsibilities is to be aware of ex¬
isting or potential problems such as contamination or
equipment failure. He may then solve such problems or
reduce their impact should they threaten the program.
First, though, he must enforce regulations designed to

Figure 1.— “Trouble Shooting Chart #2: Data
Logger Alarm System for Abnormal
Temperatures in the Insectary’' (from Wiegand
1978). A person using this chart should be able
to identify the cause of problems associated
with the air-conditioning system.

296

to determine what action to take to resolve the problem.
Familiarity with the system, knowledge of the location of
the chiller, air handlers, boiler, etc., and insuring that ex¬
tra supplies such as fan belts and prefilters are on hand
are the responsibility of the manager. If the problem can¬
not be resolved easily, the chart indicates what profes¬
sionals to call. Anticipating potential problems in this
manner functions in several important ways: chart de¬
velopment requires that the manager understand the
system; the chart itself points very quickly to possible
solutions when time is crucial; and it is a written set of
guidelines that can be used by other individuals in the
absence of the manager.

The insectary manager should anticipate problems and
project a set of possible solutions before the problems oc¬
cur. This is further exemplified in figure 2, which sum¬
marizes the contamination potential in a multispecies
rearing facility I designed (Fisher 1978; see also “Produc¬
tion of Insects for Industry. The Dow Chemical Rearing
Program,’’ by W. R. Fisher). After completing the initial
layout and before construction began, I identified poten¬
tial sources of contamination, mode of entry into the
facility and its rooms, and measures to control them. If
any major, uncontrolled sources had been detected, the
design and procedures could have been changed before
completion of the building to insure a more sanitary
operation. This type of table is just as important for pro¬
grams already in operation. Obviously, such anticipation
is impossible for all problems, but even the experience in
thinking about the more probable ones will be helpful
in developing appropriate responses to those that are
unexpected.

The most basic requirement for a manager’s success is
proper training of employees. The initial training should
be in a classroom where discussions emphasize rearing
philosophy and concepts, including relevance and objec¬
tives of the rearing program; the significance of the
employee’s participation; insect biology, ecology, and
nutrition; the presence and transmission of microbial con¬
tamination; program efficiency; and insect quality. Begin¬
ning training by directly involving an employee in rearing
processes may be overwhelming and result in one who
does not thoroughly understand the concepts underlying
the program.

Training should then shift from the classroom to produc¬
tion areas, where the manager explains the logic of why
an activity is performed as well as how it is conducted.
This training will make the employee more conscientious
in performance of his duties while enabling him to more
readily identify developing problems. For example, ex¬
plaining that surface sterilization of eggs is required
because “that’s the way it’s always been done” is not
adequate. This explanation says nothing of how impor¬

tant this procedure is to disease prevention or of how it
will affect the eggs if not performed properly. The reason
for surface sterilization should be clearly stated: “One
way disease can be spread from one insect to another is
by contamination of the outside of the egg with virus,
bacteria, or other micro-organisms. When the larva chews
a hole in the egg during hatch, it may eat some of these
microbes and become diseased. Soaking the eggs in a
bleach solution for 5 minutes destroys these harmful
organisms. Part of the eggshell is dissolved in this pro¬
cedure, but this doesn’t hurt the eggs unless they remain
in the sterilant too long. Doing so can kill them. So it is
very important that this procedure be followed
precisely.”

The daily routines that the manager establishes for
employees must be flexible enough to cover holidays, sick
leaves, vacations, etc. When the routine is temporarily
altered, the manager then sets priorities and adjusts
schedules accordingly. Organization of responsibilities in¬
cludes consideration of equipment usage so that one
employee is not unnecessarily waiting for another to
finish. Employee activities should likewise progress
logically during the day. For instance, jobs that require
sanitary conditions, like preparation of artificial diets,
should be completed before operations are begun that
create greater contamination potential, such as
harvesting pupae or washing dirty containers.

The insectary manager is responsible for regular collec¬
tion of data so that abnormal trends in production levels
and insect quality can be observed. This requires han¬
dling data in a way, such as in process-control charting
(see “Putting the Control in Quality Control in Insect
Rearing,” by D. L. Chambers and T. R. Ashley), that
quickly and easily illustrates program status. Then, if
an anomaly or harmful trend is observed, the manager
takes appropriate action. For example, I once observed
that the incidence of wing deformities in adults of a lepid-
opteran species was increasing over time as were reduced
fecundity and premature death. Variables suspected of
causing this effect were analyzed. I concluded that a lack
of polyunsaturated fatty acids in the artificial diet caused
the deformities and other symptoms. These were then
eliminated by adding raw linseed oil to the diet. As in
this example, subtle changes can occur slowly, often im¬
perceptibly when observed on a daily or weekly basis; but
they may become manifest as a major problem when data
are analyzed over longer periods.

Finally, the insectary manager is the spokesman for the
program and the interface between it and the researchers
who actually use the insects (see “Management of Insect
Production,” by Charles P. Schwalbe and O. T.

Forrester). He coordinates the requirements of the rearing
program with those of the researchers towards a common

297

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goal. So changes in yields, adjustments in insect quality,
etc., desired by researchers should be requested of the
manager, who can then modify the program accordingly.

Conclusions

The insectary manager links the unique challenges and
problems of insect production with the practical applica¬
tion of rearing concepts. He links the control of con¬
tamination and the improvement of insect quality with
environmental monitoring and manipulation. He links
personnel with performance guidelines and effective use
of the facility. Finally, he links old, tested procedures and
techniques with new ones that improve program efficien¬
cy, safety, and capability to produce insects of better
quality. Philosophically, this requires that he view the
production program as a dynamic system that evolves as
new technology and techniques are developed. The
manager, as the interface between these ideas and his
program, has a thorough understanding of them both and
the best perspective for planning and making changes.

Acknowledgments

I thank Jeanne Wiegand, Department of Veterinary
Medicine, University of Florida, for her review and
discussions about insectary management.

References
Fisher, W. R.

1978. Contamination control in the Dow insect rear¬
ing facility. In W. R. Fisher and J. Wiegand,
(eds.), The Dow-Walnut Creek Insectary: An
Operational Manual, pp. 52-64. Dow Chemical
U.S.A., Walnut Creek, Calif.

Wiegand, J.

1978. Alarms. In W. R. Fisher and J. Wiegand, (eds.),
The Dow- Walnut Creek Insectary: An Opera¬
tional Manual, pp. 77-86. Dow Chemical
U.S.A., Walnut Creek, Calif.

299

Management of Insect Production

By Charles P. Schwalbe and 0. T. Forrester1

Introduction

“Management” is a generally used and deceptively sim¬
ple word that implies the accomplishment of an end by
judicious use of means. But close examination of situa¬
tions relying heavily on management reveals that the
means critical to accomplishment of the end are usually
complex and unwieldly. Many of today’s managers are ill-
equipped to deal with this complexity, even though sev¬
eral training programs, university curriculums, books,
etc., are available that deal with the theory and practical
applications of management concepts. Necessity often
places people with varying degrees of experience and
training in management roles. These newborn managers
usually do the best they can; but, for several reasons,
some are successful and others are not. Failures are
tolerable in systems equipped with fail-safes and backups
designed to minimize the effects of poor management,
but insect-rearing facilities and laboratories are usually
austere operations where fail-safes are uncommon. Here
we will examine what production managers in insect¬
rearing operations can do to prevent system failures and
show how the various components of insect production
depend on each other.

The Production Manager

Anyone responsible for an insect-rearing program is a
manager. The manager’s degree of responsibility depends
mainly on how big and how complex the production sys¬
tem is; it could be small-scale rearing for laboratory and
field research programs, pilot-scale production supporting
large experiments or pilot projects, or mass rearing in
direct support of operational line programs. But basic
managerial principles of planning, organizing, monitoring,
and upgrading apply to all cases. If these principles are
ignored, the rearing operation will probably fail. One per¬
son should be clearly responsible for the functioning of
the operation and should have the authority to carry out
the duties effectively. This person is a production
manager.

The production manager is the link among three major
units: technology and production groups within the pro¬
duction program and user groups for whom the insects

‘Center director and entomologist, Otis Methods Development
Center, Animal and Plant Health Inspection Service, U.S.
Department of Agriculture, Otis Air National Guard Base,
Mass. 02542

are produced. The manager insures that the technology
group designs a rearing system that will reliably and effi¬
ciently produce what the user needs. The manager must
also insure that pertinent technical information is
available to the program. Designing a rearing operation
without technical input from experts (engineers, physio¬
logists, geneticists, pathologists) would be foolish. Still,
by using commonsense, a manager can design and im¬
prove many operational details; an effective manager will
find the most expedient options. The technology group is
also instrumental in troubleshooting operations, resolving
technical problems, and conducting research for modify¬
ing and improving production sequences. A well-directed
technology group not only operates in the present, keep¬
ing things working, but also develops basic information
on innovative, more efficient production methods.

The production manager will also participate in the plan¬
ning of user programs, especially when large field tests or
operational insect-population-suppression programs rely
on receiving large numbers of insects regularly. The
reared insects will be a resource that line managers will
count on for their programs; they must know what rear¬
ing capabilities and limitations will be so that they can
plan the program's size and timing. In providing such in¬
formation, the production manager is able to show how
the rearing program might affect the user program.

Directing the technology group and helping plan the user
program are important duties for production managers,
but their most challenging role is to manage the various
resources in a production group in a way that is consis¬
tent with available technology and user needs. These
guide the methods, level, and duration of the rearing pro¬
gram. The production manager must have an effective
relationship with the production group so that the
technology is used most effectively and the user receives
quality insects on time and at the least possible cost.

Elements

of Production Management

Production management requires development of several
components (objectives, resources, operating plans, re¬
view, and technology development and implementation)
that, when compiled and coordinated, constitute a unified
production plan. This work plan should be carefully
prepared because it enables the rearing program to be
conducted in a uniform, systematic, and orderly fashion.
Failure to carefully consider any one of these components
and its place in the total system leaves the rearing pro-

300

gram vulnerable to breakdown. And failure to reassess all
elements regularly and to redesign them when necessary
may result in an outmoded plan.

Objectives

A production program must have established objectives.
A statement of objectives, telling how many insects will
be ready in a given time, is a necessary first step toward
developing plans for other production-management
elements. These statements can also tell program par¬
ticipants what is going on and, so, solve the common
problem of employees and cooperators not identifying
with the purposes of their activities.

Establishing how many insects will be available, and
when, is usually straightforward. The manager organizes
an annual production schedule (fig. 1). Such a plan iden¬
tifies production-delivery responsibilities and is the foun¬
dation for all other plans. A schedule of this nature is
especially useful when the program provides insects of
different stages or species to many users.

Objectives also define other important aspects of the pro¬
duction program, especially ways to improve the efficien¬
cy of operations. Improved quality standards, increased
yields, development of new techniques, implementation of
demonstrated techniques, training, improvement of the
physical plant, and staffing are all factors to be con¬
sidered in plotting a course of action. As with the produc¬
tion schedule, the objectives should be specific, quan¬
tified, and organized to show how long they will take to
fulfill.

Resources

The management of resources used in the production of
insects can be complicated. A well-planned, efficient rear¬
ing program requires adequate resources, but gross
surpluses are wasteful. Any activity will slow when the
supply of an essential resource dwindles, and insect rear¬
ing is no different. The production manager must devise
specific procedures to facilitate management of time,
space, personnel, materials, and funds.

Time. — Time-and-motion information is useful in deter¬
mining how much time is needed for accomplishing cer¬
tain activities (diet preparation, implanting, harvesting,
cleaning, etc.). The production manager should also know
how much time will be needed for future events such as
colony scale-up, diapause, insect developmental times,
purchasing and delivery of supplies, inventory utilization
rates, and hiring procedures. Failure to account for possi¬
ble lags in procurement of dietary ingredients, for exam¬
ple, could easily allow the inventory to run out.

Figure 1.— Stylized annual production
schedule.

Space.— Available space must be allocated and managed
so that all essential activities (supplies storage, diet
preparation, environmentally controlled holding,
harvesting, etc.) are accommodated. In arranging space,
the production manager must be especially aware of the
need for pathogen isolation or containment. Work areas
may require special utilities or environmental conditions
that will affect space allocation. The manager must also
consider what space will be needed for future modifica¬
tions of existing areas of the facility committed to a cer¬
tain function. It is not unusual, for instance, for space to
be a limiting resource when increased production is need¬
ed; all other resources may be available; but, if there is no
room to use them in, the amount of insects produced will
not increase.

Personnel.— In developing staffing patterns for the pro¬
gram, production managers must plan for the seasonal
nature of insect-production operations; so they often rely
on temporary employees. Proper training and motivation
of these temporary employees, in particular, help to in¬
sure that the rearing system continues to work and be ef¬
ficient. Resourceful managers will make certain that
employees know the program’s purpose and how they
contribute to its accomplishment no matter how insignifi¬
cant their jobs may seem to be.

Materials.— A wide variety of materials and equipment is
used in insect production, and these must be available
when they are needed. Rearing programs often have prob¬
lems when rearing methods require unique, little-used
materials since the commercial availability of these items
is likely to be sporadic. Alternatives should be identified
for use in the event of unforeseen shortages. Inventories

301

of materials such as containers, lids, and dietary ingre¬
dients are best expressed in units per 1,000 (or 1 million)
insects so that potential production can be easily deter¬
mined from inventory reports. When inventories reach a
certain level, resupply should be automatic and in quan¬
tities required to rear a given number of insects. This
way, cycles of production may begin with new inventories
of supplies. Procurement and contracting processes,
though time consuming, must be accounted for in a per¬
fectly tuned inventory-control system. Otherwise, sup¬
plies will run out while resupply is being processed. A
well-run inventory-control system will insure a steady
supply of materials.

Automated or semiautomated insect-rearing operations
that rely on costly equipment are likely to collapse when
mechanical breakdowns occur. But having replacement
machines is usually not possible, so parts should be
stocked to facilitate onsite repairs. If a breakdown is re¬
current and repair becomes routine, the machine should
be replaced or modified, or a preventive-maintenance plan
should be devised. Preventive maintenance is an excellent
way to lengthen equipment life and avert breakdowns.

Funds.— Fiscal management means getting the greatest
number of quality insects for the least money. The pro¬
duction manager must find low-cost supplies that con¬
form to established specifications. These should fulfill
quality requirements of the reared insects. For example,
bacteriological-grade agar need not be used if a less re¬
fined grade is adequate for normal growth, development,
and reproduction of the colony; or less costly gelling
agents such as carrageenins may be acceptable. If not
enough is known about the relationship between cost and
quality, favor quality.

Careful examination of a production system often reveals
many opportunities for economizing, but savings that
cause the user program to fail are meaningless. In many
cases, mechanization will be cost effective; time-and-
motion analyses, for instance, may help identify costly
operations requiring additional research to increase effi¬
ciency. At other times, simple reorganization of operating
sequences or rearrangement of work areas may increase
productivity.

Time, space, personnel, materials, and funds are the main
resources a manager has to work with in the production
program. When they are available in appropriate propor¬
tions, the rearing system operates effectively; imbalances
lead to inefficiency. Therefore, each rearing program must
reach its own balance and be prepared for necessary
readjustments.

Operating plans

Operating plans are written prescriptions with enough
detail to guide inexperienced workers through their jobs.
Such a plan is commonly called a standard operating pro¬
cedure (SOP); its purpose is to define and describe all
jobs conducted in a systematic production program. Con¬
structing an operating plan is difficult and requires the
production manager to decide how and when certain jobs
are performed. The manager should keep the SOP up to
date and must insure that its guidelines are followed. The
operating plan should be under constant review and
should be modified as refinements are made in the rear¬
ing operations. New methods should not be introduced in¬
to the system until they have been thoroughly tested and
made workable. A useful SOP has all rearing activities ar¬
ranged in chronological sequence in a looseleaf folder, so
modifications, additions, or deletions can be made easily.

The manager should have contingency plans prepared for
use in the event the operating plan is disabled. Normally,
the contingency plan will be less than ideal for several
reasons, but it will help maintain production while the
system is being repaired. For example, a contingency
plan would allow the use of a locally available gelling
agent after the stock of specified agar is depleted and
before a new supply is delivered. Shortages such as these
may be difficult to avoid for several reasons, and proven,
available alternatives should be identified. A contingency
plan should be used only when absolutely necessary.

An SOP also gives managers a written standard of
employee performance, and workers have a clear
understanding of what they are supposed to do. So the
SOP provides managers and workers with a common
ground that helps eliminate daily variations in the way
certain jobs are done and helps identify areas where
training can be provided to improve skills. If the
manager does not use a detailed operating plan, workers
will develop their own ways of accomplishing their tasks,
and those methods will vary from worker to worker.
Operating plans should embrace all aspects of the produc¬
tion program from the seemingly trivial, routine opera¬
tions to those of great complexity or sensitivity.

Review

If careful attention has been given to organizing objec¬
tives, resources, and operating plans, the rearing program
will probably be successful. The objectives, resource-
management scheme, and operating and contingency
plans are collated into a standard production work plan.
Of course, the rearing operation has not been set into
perpetual motion. Continual managerial guidance is re¬
quired to insure that everything is going according to
plan, since unforeseen and potentially catastrophic situa-

302

i

/ \

Implementation

Figure 2.— Technology development and im¬
plementation.

tions can arise quickly. Most problems will be easily iden¬
tifiable; solving them improves the system and prevents
similar difficulties in the future. So the manager should
make sure that quality is monitored and that the system
is under continual review.

Intensive monitoring of insect quality is a production
manager’s primary means for detecting system break¬
down. Other means are available; employee suggestions,
safety records, and cost analyses can all be valuable in¬
dicators of various deficiencies. The manager must
believe that an attempt should be made to prevent the
recurrence of any complaint or technical problem no mat¬
ter how small. Problems may be dealt with directly
through changes in scheduling or production sequence; or
they should be referred to the technology group if they
are technical. Later development in research or methods
may be needed; but, in many cases, data may already be
available to base alternatives on. If new technology is re¬
quired, it should be developed and implemented accord¬
ing to well-defined guidelines.

Technology development
and implementation

Reports of declining yield or quality may indicate a need
for improvements in operations. Operational adjustments
may also be made in response to increasing costs, declin¬
ing budgets, decreasing personnel ceilings, enlarged user
needs, or unavailability of materials. Production man¬
agers faced with any or all of these problems use the
review process to identify critical problem areas and
establish their order of importance so that efforts to im¬
prove technology can be properly focused.

Technology development and its implementation in the
rearing system is an interdependent process (fig. 2) with
research, methods development, and pilot testing inter¬
connected by a two-way flow of technology and informa¬
tion. If the process works properly, research leads to
methods development, which may discover problems that
require more research; if not, the new technology is pilot-
tested; any problems discovered there are returned to
research or methods. Finally, when all the problems are
solved, the technology is implemented. Theoretically, the
flow between pilot testing and implementation is one¬
way.

Research and development should be directed to areas
where needs are greatest. The technology group must be
sensitive to practical problems of production because
newly produced technology is seldom incorporated into
the SOP without large-scale testing and further adjust¬
ment. The manager must consider insect yield and the
quality and efficiency of associated rearing operations
when determining whether or not the new technology will
be applicable. Only in unusual cases of need should the
new technology be implemented without intermediate
pilot testing.

Conclusions

If the individual overseeing the operations of insect pro¬
duction does not appreciate the broad scope of the assign¬
ment, day-to-day problem-solving activities will replace
the more profitable long-term planning and management
of the facility. Appreciating the interactions of the com¬
ponents as discussed in this paper will facilitate smooth
operations and effective communication among all
participants.

303

Politics in Insect Rearing and Control

The Medfly Program in Guatemala and Southern Mexico

By Patrick Patton1

Politics play in important part in any program for insect
rearing and control. Even a localized program will have
internal and local politics to contend with. The larger the
territory covered by a program, the more important
politics become. In an international program, the scope
and depth of political problems can be enormous, and
they must be dealt with in many ways. The international
program to control the Mediterranean fruit fly, Ceratitis
capitata (Wiedemann), in Guatemala and southern Mexico
is an excellent example. It is the largest program of its
kind ever attempted, and political problems and their
solutions have been important to the program since its
inception.

The current program for controlling the Mediterranean
fruit fly (medfly) is an outgrowth of political events that
took place after the medfly became established in Costa
Rica (fig. 1) about 1955 and soon thereafter spread to
Nicaragua and Panama. Fearing that it would spread far¬
ther northward, the United States and Mexico joined
with several international organizations in the mid-1960’s
to begin a program to eradicate the medfly from
Nicaragua and maintain a barrier zone south of its border
with Costa Rica. The program was successful. But, as so
often happens with such delicately balanced programs, it
collapsed when much of the funding was discontinued, in
this case by the United States and Mexico. The medfly
was still far from their borders and therefore not con¬
sidered a critical problem.

Political and sociological changes in the region exacer¬
bated the medfly problem during the next 10 years. An
apparently effective quarantine between Nicaragua and
Honduras began to fail as transportation of agricultural
products between the countries increased. The chaos
caused by the earthquake that devastated Managua,
Nicaragua, in 1973 eliminated the remaining semblance of
a quarantine, and the medfly invaded El Salvador and
Guatemala in less than a year. The United States and
Mexico again became concerned, wanting to stop the
medfly before it crossed the Mexican border.

Cooperation was needed from the Governments of El
Salvador and Guatemala to construct a production facili-

'Subdirector de Servicios Fitosanitarios, Sanidad Vegetal, Sub-
direccion de Servicios Fitosanitarios, Mexico 21, D.F.

ty, produce 100 million sterile flies per week, and create
another barrier. But, in 1975, the Government of El
Salvador was unstable, the country on the brink of
revolution. Guatemala was an alternate possibility, but
its main crops were not heavily affected by the medfly
and were by other pests, so it was not very interested in
the problem. Guatemala did, however, want Mexican oil,
and this became a bargaining chip. So, in 1977,

Guatemala and Mexico formed the Comision Moscamed
to combat medflies in Guatemala.

The political negotiations that established this commis¬
sion were concluded as the first medfly was discovered
along the northern side of the Rio Suchiate between Mex¬
ico and Guatemala. By the end of 1977, 122 flies had
been detected. In the face of this invasion, Mexico sought
financial and technical support for a program to drive the
medfly out. Several national and international organiza¬
tions were approached. These included the Organization
of American States, the International Atomic Energy
Agency, the Food and Agriculture Organization (FAO),
and the agriculture and state departments of concerned
countries. After much negotiation, the program was
financed by means of a cooperative agreement that pro¬
vided for Mexico and the United States each to cover half
the cost. Several of the international organizations were
also to participate in the program. It was decided to con¬
struct a production facility at Metapa, Mexico; establish
a barrier zone between Mexico and Guatemala; and
eradicate the medfly from north of this border. The facili¬
ty began production in March 1979 and was producing
500 million sterile flies per week by 1980.

One of the political problems in an international program
is that opinions about how things should be done must
be handled with diplomatic sensitivity. When conflicting
advice comes from many nations and regions whose par¬
ticipation is necessary, someone must resolve the con¬
flicts tactfully and without harming the program. In the
medfly program, this problem is intensified because the
facility is managed by a group of people whose average
age is under 30, a perceived handicap that means they
have to defend their actions constantly and spend extra
time maintaining their credibility. The challenge for this
or any group is to evaluate available information and
adopt or modify the most appropriate technology without
making any enemies by rejecting someone’s advice. So, in
the medfly program, the Mexican general coordinator
reviews all suggestions and plans coming from various

304

INFESTED

COSTA RICA

PANAMA

PROTECTED

BELIZE

HONDURAS

NICARAGUA

Figure 1.— Areas of Central America and Mexico infested by and protected from the medfly.

people and agencies and then makes the decisions in con¬
junction with his counterparts. Dealing with the political
problems in this way, the Metapa group adapted the rival
Hawaiian and Viennese mass-rearing systems to the
much larger production quotas of Metapa and emerged
with the Metapan system.

These problems stem from the international aspects of
the medfly program. But, as with all insect-rearing pro¬
grams, local politics are also important. A good way to
avoid local political problems is to keep everyone affected
by a program informed about its purposes and methods.
But doing so is often difficult for the medfly program
because it encompasses diverse geographical areas,
climates, human cultures, agricultural practices, and fly
habitats. For example, consider a massive bait-spray and
sterile-insect release operation conducted along the border
between Mexico and Guatemala. The spray was 800 ml of

protein hydrolyzate plus 200 ml of malathion2 per hec¬
tare. The people living in the area did not understand
that this spray was not dangerous to them, their crops,
or their animals; nor did they understand the importance
of bait spraying to medfly control. So, whenever an
airplane was spraying, they would hide. And they blamed
the spray for everything from dead chickens to the com¬
mon cold. They were also afraid that this was some
drastic governmental form of birth control. Politicians
played on this fear to win votes; and newspapers, for
their own reasons, printed many negative articles about
the whole program. At one point, concerned about the
failure of their drought-stricken crops, 4,000 farmers
stormed my office at the production facility, blaming

20, 0-Dimethyl phosphorodithioate of diethyl mercaptosuccinate.

305

their problems on the bait spraying. So, even though ef¬
forts had been made from the beginning to educate the
people, a more intensive information campaign was devel¬
oped to explain the benefits of controlling the medfly.

The political problems inherent in such a program will
always be there. Program managers must, therefore, be
politicians. They have to convince potential funding

sources that the program is needed and can be successful.
They have to coordinate the contributions of program
participants with the skill of seasoned diplomats. They
have to insure that the public is well-informed about
what the program is doing and why. In short, there is
more to managing an insect-rearing program than just
rearing insects.

*U.S. GOVERNMENT PRINTING OFFICE: 1983-769-037:10

306

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