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Full text of "Culture methods for invertebrate animals;"

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Marine Biological Laboratory Library 

Woods Hole, Massachusetts 




Gift of Bostwick H. Ketchum - 19?6 









CULTURE METHODS 

FOR 

INVERTEBRATE ANIMALS 



THE PREPARATION OF THIS COMPENDIUM 

WAS MADE POSSIBLE BY A GRANT FROM 

THE NATIONAL RESEARCH COUNCIL 






CULTURE METHODS 

FOR 

INVERTEBRATE 

ANIMALS 

A Compendium prepared cooperatively by 

American zoologists under the direction 

of a committee from Section F of 

the American Association for 

the Advancement of Science 




PAUL S. GALTSOFF 

FRANK E. LUTZ • PAUL S. WELCH 

JAMES G. NEEDHAM, Chairman 

i by many specialists whose names appear 
connection with their respective 
contributions to this volume. 




Ithaca, New York 
Comstock Publishing Company, Inc. 

l 937 



COPYRIGHT 1937, BY 
COMSTOCK PUBLISHING COMPANY, INC. 



PRINTED IN THE UNITED STATES OF AMERICA 

BY R. R. DONNELLEY & SONS COMPANY, 

CRAWFORDSVILLE, INDIANA, AND CHICAGO, ILLINOIS 



PREFACE 



This book has been prepared as an aid to studies that 
require living animals in continuous supply. They 
are needed both jor teaching and for research, espe- 
cially for research in genetics, in parasitology, in experi- 
mental zoology, in economic entomology, and in nearly every 
field of applied biology. They are needed in aviaries, in 
aquaria, in fish culture experiment stations, and in zoological 
gardens where they are used as food for carnivorous animals. 
Such needs have seemed to justify an effort to gather to- 
gether the experience of many scattered workers and make 
their useful results more widely available. 

The task of the committee in charge has been a simple 
one — that of serving as a medium for the exchange of ex- 
perience by actual workers. So we have announced the under- 
taking to the zoological public and we have personally 
invited contributions from those we have known to be keeping 
cultures of small animals. We have put together with a 
minimum of editing what they have offered. Our responsi- 
bility ends here. The articles bear the names of their con- 
tributors. We do not guarantee the methods outlined. Indeed, 
there are great difficulties in keeping cultures going, and 
failures will occur betimes even in the hands of careful 
workers. Constant alertness is required to see that fit con- 
ditions are maintained, and a large measure of ingenuity is 
often necessary to adapt places and circumstances to keeping 
conditions fit. 



vi Preface 

Thus this is a compendium in the older sense of that word 
fcon and pendere, weighing together), rather than a digest. 
Perhaps condensation may be in order later when more 
animals have been managed successfully and more expe- 
rience has accumulated. This is only a beginning. The gaps 
in our knowledge are numerous and very obvious. Com- 
paratively few invertebrate animals have been tried as yet, 
and very few have been managed with complete success. 

These pages will record the experience of those among us 
who have had some measure of success in rearing various 
invertebrate animals, and will give the methods they have 
found most useful. It is to be expected that these methods 
may be improved by further trial. That economic values and 
gains to health and comfort will grow out of further work 
along these lines is altogether probable, for the scientific use 
of our animal resources has only just begun. 

Some very simple procedures are included. We have not 
forgotten the needs of the high school teacher who is wise 
enough and diligent enough to teach zoology with the saving 
grace and with the quickening thrill that comes from the use 
of living materials. 

The Committee wishes to acknowledge the efficient aid 

of its secretary, Miss Mary E. Davis, who has carried a 

very large share of the burden of this compilation. The task 

has been lightened by receipt of many valuable voluntary 

contributions, and the response from those to whom we have 

appealed for culture methods has been both prompt and 

generous. 

Paul S. Galtsoff 

Frank E. Lutz 

Paul S. Welch 

James G. Needham, Chairman 



TABLE OF CONTENTS 



Introduction 

By James G. Needham, Cornell University 

General Methods of Collecting, Maintaining, and Rearing 
Marine Invertebrates in the Laboratory 
By Paul S. Galtsoff, U. S. Bureau of Fisheries 

i. Collecting 

2. Marine Aquaria 

3. Artificial Seawater 

4. Methods of Securing Food for Marine Invertebrates 

5. Methods of Obtaining a Culture of a Single Food Species 

Collecting and Rearing Terrestrial and Freshwater Invertebrates 
By F. E. Lutz, J. G. Needham, and P. S. Welch 

1. Collecting and Handling Living Specimens 

Aquatic Animals 
Aerial Insects . 

2. Cages and Shelter 

3. Cage Management 

Feeding and Watering . 
Temperature and Humidity 
Sanitation 



PAGE 

3 



5 

13 
27 

3 1 
36 



40 
41 
44 
46 
48 
48 
48 

49 



Phylum I. Protozoa 

Class Mastigophora 

Growth of Free-Living Protozoa in Pure Cultures . . . 51 

By R. P. Hall, University College, New York University 

Cultivation of Protozoa 59 

By John P. Turner, University of Minnesota 

Polytoma Cultures 61 

By Josephine C. Ferris, University of Nebraska 

Culturing Euglena proximo, 61 

By J. A. Cederstrom, University of Minnesota 

A Culture Medium for Free-Living Flagellates (Reprinted) . .62 
By James B. Lackey, U. S. Public Health Service 

vii 



viii Contents 

PAGE 

Culture of Some Flagellates and Ciliates 63 

By Paul Brandwein, Washington Square College, New York University 

A Simple Method for Culturing Trypanosoma lewisi ... 64 
By Reed O. Christenson, University of Minnesota 

Notes on Culturing Certain Protozoa and a Spirochaete Found 

in Man 65 

By M. J. Hogue, University of Pennsylvania Medical School 

Frog and Toad Tadpoles as Sources of Intestinal Protozoa for 

Teaching Purposes (Reprinted) 68 

By R. W. Hegner, School of Hygiene and Public Health 

Class Sarcodina 

Protozoan Cultures 69 

By George R. La Rue, University of Michigan 

Culture of Some Freshwater Rhizopoda 72 

By Paul Brandwein, Washington Square College, New York University 

Amoeba 74 

By William LeRay and Norma Ford, University of Toronto 

Stock Cultures of Amoeba proteus (Reprinted) . . . -75 
By H. W. Chalkley, U. S. Public Health Service 

The Culture of Amoeba proteus Leidy partim Schaeffer . . .76 

By D. L. Hopkins, Duke University, and D. M. Pace, Johns Hopkins 
University 

Culturing Amoeba proteus and A. dubia 80 

By H. R. Halsey, Columbia University 

Cultivation of Mayorella {Amoeba) bigemma on Euglena gracilis . 81 
By John C. Lotze, Ohio State University 

The Culture of Flabellula mira 83 

By D. L. Hopkins, Duke University, N. E. Rice, Brenau College, and 
H. E. Butts, Wellesley College 

Valkampfia calkinsi and V. patuxent (Abstracted) .... 86 

By C. M. Breder, Jr., New York Aquarium, and R. F. Nigrelli, New 
York University 

Notes on Various Media Used in the Culture of Intestinal Protozoa . 86 
By Charles A. Kofoid and Ethel McNeil, University of California 



Contents ix 

PAGE 

In Vivo Cultivation of Intestinal Protozoa in Parasite-Free 

Chicks (Reprinted) 89 

By R. W. Hegner, School of Hygiene and Public Health 

A Method of Culturing Arcellae 9 1 

By E. D. Miller, University of Virginia 

Culturing Arcellae 9 2 

By Bruce D. Reynolds, University of Virginia 

Methods of Culturing Testacea 9 2 

By A. B. Stump, University of Virginia 

Culture Methods for Marine Foraminifera of the Littoral Zone . 93 
By Earl H. Myers, Scripps Institution of Oceanography 

Class Sporozoa 

Maintenance of Laboratory Strains of Avian Plasmodium and 

Haemoproteus 9° 

By Clay G. Huff, University of Chicago 

Class Ciliata 

Culture Media for Opalinidae (Abstracted) 98 

By Maynard M. Metcalf, Johns Hopkins University 

Culture Medium for the Ciliate Lacrymaria (Abstracted) . . 99 
By Y. Ibara, Johns Hopkins University 

The Culture of Didinium nasutum 100 

By C. Dale Beers, University of North Carolina 

Controlled Cultures of Freshwater Ciliates 103 

By Alford Hetherington, Stanford University 

Cultivation of Colpidium campylum 108 

By T. M. Sonneborn, Johns Hopkins University 

The Culture of Colpidium campylum 109 

By C. V. Taylor, J. O. Thomas, and M. G. Brown, Stanford University 

A Culture Method for Colpoda 109 

By M. S. Briscoe, Storer College 

Stock Cultures of Colpoda (Reprinted) no 

By Joseph H. Bodlne, Iowa State University 



x Contents 



PAGE 



The Culture of Colpoda cucullus II0 

By H. Albert Barker and C. V. Taylor, Stanford University 

Some Methods for the Pure Culture of Protozoa . . . .112 

By William Trager, Rockefeller Institute for Medical Research 

Thyroid Cultures of Paramecia (Reprinted) 119 

By William L. Straus, Jr. 

A Combined Culture Method and Indicator for Paramecium 

(Reprinted) "9 

By Robert T. Hance, University of Pittsburgh 

Paramecium I2 ° 

By William LeRay and Norma Ford, University of Toronto 

A Culture Medium for Paramecium (Reprinted) . . . .120 
By Lauren E. Rosenberg, University of California 

Cultivation of Paramecium aurelia and P. multimicronucleatum . 121 
By T. M. Sonneborn, Johns Hopkins University 

Paramecium multimicronucleatum: Mass-Culturing, Maintain- 
ing and Rehabilitating Mass-Cultures, and Securing Con- 
centrations I22 

By Edgar P. Jones, University of Akron and University of Pittsburgh 

Culturing Paramecium caudatum in Oat Straw Infusion . . .127 
By George A. Smith, Eugenics Record Office 

A Culture Method for Paramecium multimicronucleatum and 

Oxytricha jallax l2 & 

By A. C. Gtese, Stanford University 

The Cultivation of Nyctotherus ovalis and Endamoeba blattae 

(Reprinted) 128 

By Harry E. Balch, University of California 

A Method for Culturing Bursaria truncatella (Reprinted) . .129 
By Amos B. K. Penn, Tsing Hua University, Peiping, China 

Uroleptus mobilis (Abstracted) 13° 

By Gary N. Calkins, Columbia University 

Methods for Culturing Pleurotricha 13 1 

By Amos B. K. Penn, Tsing Hua University, Peiping, China 



Contents xi 

PAGE 

Stylonethes sterkii (Abstracted) 13 2 

By Laura Garnjobst, Stanford University 

A Method for Inducing Conjugation Within Vorticella Cultures . 132 
By Harold E. Finley, West Virginia State College 

Miscellaneous Classes and Microbiology 

A Novel Method of Obtaining Protozoa 134 

By W. H. Davis, Massachusetts State College 

Permanent Cultures (Reprinted) 134 

By Frederick Bauer, Rhode Island State College 

Food Organisms for Marine and Halobiont Animals . . . . 134 
By R. M. Bond, Santa Barbara School, Carpinteria, California 

Wheat-Grain Infusion l 35 

By John W. Nuttycombe, University of Georgia 

Phylum II. Porifera 

Notes on the Cultivation and Growth of Sponges from Reduc- 
tion Bodies, Dissociated Cells, and Larvae 137 

By H. V. Wilson, University of North Carolina 

Phylum III. Coelenterata 

Class Hydrozoa 

Hydras Uo 

By Libbie H. Hyman, New York City 

The Culture of Some Miscellaneous Small Invertebrates . . .142 
By Paul Brandwein, Washington Square College, New York University 

Class Scyphozoa 

Rearing the Scyphistoma of Aurelia in the Laboratory . . .143 
By F. G. Gilchrist, Pomona College 

Class Anthozoa 

Sagartia luciae *44 

By Donald W. Davis, The, College of William and Mary 



xii Contents 

PAGE 

Rearing Coral Colonies from Coral Planulae 145 

By Thomas Wayland Vaughan, Scripps Institution of Oceanography 

Phylum V. Plathelminthes 
Class Turbellaria 

Culture of Stenostomum oesophagium 148 

By Margaret Hess, Judson College, Marion, Alabama 

Cultivation of Stenostomum incaudatum 148 

By T. M. Sonneborn, Johns Hopkins University 

The Culture of Microstomum 149 

By M. Amelia Stirewalt, University of Virginia 

Geocentrophora applanatus 151 

By William A. Kepner, University of Virginia 

Culture of Planaria [=Eu plan aria] agilis 151 

By Rosalind Wulzen, Oregon State College, and Alice M. Bahrs, St. 
Helen's Hall Junior College 

Planaria 153 

By William LeRay and Norma Ford, University of Toronto 

Collection and Culture of Planaria 153 

By George R. La Rue, University of Michigan 

Planarians 154 

By Libbie H. Hyman, New York City 

Class Trematoda 

The Parasitic Flatworms 156 

By H. W. Stunkard, University College, New York University 

Epibdella melleni 158 

By Theodore Louis Jahn, State University of Iowa 

Class Cestoidea 

Intermediate Stages of Cestodes 160 

By Reed O. Christenson, University of Minnesota 

Phylum VI. Nemertea 

Methods for the Laboratory Culture of Nemerteans . . . .162 
By Wesley R. Coe, Yale University 



Contents xiii 

PAGE 

Phylum VII. Nemathelminthes 
Class Nematoda 
Recovering Infective Nematode Larvae from Cultures . . .166 
By G. F. White, U. S. Bureau of Entomology and Plant Quarantine 

Rearing Trichinella spiralis ^8 

By Reed O. Christenson, University of Minnesota 

The Growth of Hookworm Larvae on Pure Cultures of Bac- 
teria (Reprinted) l 7° 

By Oliver R. McCoy, School of Hygiene and Public Health 

Culturing Eggs of the Fowl Nematode, Ascaridia lineata . . . 171 

By J. E. Ackert, Kansas State College 
The Life History of the Swine Kidney Worm (Abstracted) . .172 

By Benjamin Schwartz, U. S. Bureau of Animal Industry 

Cultivation of a Parasitic Nematode 17 2 

By William Trager, Rockefeller Institute for Medical Research 

Culturing Parasitic Nematode Larvae from Silphids and Re- 
lated Insects l 73 

By C. G. Dobrovolny, University of Michigan 

Anguilla aceti (Abstracted) 173 

By George Zebrowski, Buck Creek, Indiana 

Artificial Cultivation of Free-Living Nematodes . . . . 174 
By Asa C. Chandler, Rice Institute 

Phylum VIII. Trochelminthes 
Class Rotatoria 

A Culture Medium for Hydatina senta 176 

By Josephine C. Ferris, University of Nebraska 

Class Gastrotricha 

Method of Cultivation for Gastrotricha 176 

By Charles Earl Packard, University of Maine 

Phylum IX. Bryozoa 
Class Ectoprocta 

Bugula flabellata and B. turrita 178 

By Benjamin H. Grave, De Pauw University 



xiv Contents 

PAGE 

Culturing Freshwater Bryozoa x 79 

By Mary Rogick, College of New Rochelle 

Phylum XIII. Annelida 
Class Polychaeta 

Nereis limbata I ° 2 

By Benjamin H. Grave, De Pauw University 

A Method for Rearing Nereis agassizi and N. procera . . .184 
By John E. Guberlet, University 0} Washington 

Hydroides hexagonus I &5 

By Benjamin H. Grave, De Pauw University 

Sabellaria vulgaris I ^7 

By Alex B. Novikoff, Brooklyn College 

Class Oligochaeta 

Cultivation of Enchytraeus albidus 19 J 

By Raymond F. Blount, University of Minnesota Medical School 

Laboratory Culture of Enchytraeus 193 

By Victor Loosanoff, U. S. Bureau of Fisheries 

Enchytraeid Worms I 93 

By William LeRay and Norma Ford, University of Toronto 

Tubificidae x 94 

By George R. La Rue, University of Michigan 

Earthworms x 95 

By Walter N. Hess, Hamilton College 

Culture of Allolobophora I 9° 

By R. N. Danielson, University of Minnesota 

Class Gephyrea 

Culturing Larvae of Urechis caupo r 97 

By G. E. MacGinitte, California Institute of Technology 

Class Hirudinea 

Laboratory Care of Leeches 201 

By J. Percy Mooee, University of Pennsylvania 



Contents xv 

PAGE 

Phylum XIV. Arthropoda 

Class Crustacea 

A Method for Rearing Artemia salina 205 

By R. M. Bond, Santa Barbara School, Carpinteria, California 

Culture of Cladocera 20 7 

By A. M. Banta, Brown University 

A Note on Banta's Culture Medium 211 

By George G. Snider, University of Cincinnati 

Cladocera Culture 2I1 

By Harold Heath, Hopkins Marine Station 

A New Culture Medium for Cladocerans (Reprinted) . . .212 
By Walter A. Chipman, Jr., U. S. Bureau of Fisheries 

A Culture Medium for Daphnia (Reprinted) 213 

By R. M. Bond, Santa Barbara School, Carpinteria, California 

Methods for Culturing Daphnia 214 

By Arthur D. Hasler, U. S. Bureau of Fisheries 

Daphnia Culture 2I 5 

By Alfred W. Schluchter, Dearborn, Michigan 

Propagating Daphnia and Other Forage Organisms Intensively 

in Small Ponds (Abstracted) 216 

By G. C. Embody, Cornell University, and W. O. Sadler, Mississippi College 

Daphnia Culture 2I 9 

By Libbie H. Hyman, New York City 

Chydoridae 22 ° 

By Charles H. Blake, Massachusetts Institute of Technology 

Culture Methods for Pelagic Marine Copepods 221 

By George L. Clarke, Woods Hole Oceanographic Institution and 
Harvard University 

Notes on the Cultivation of Tigr'wpus fidvus and Calanus 

finmarchicus 22 4 

By R. M. Bond, Santa Barbara School, Carpinteria, California 

Culture Methods for Cyclopoid Copepods and Their Forage 

Organisms 22 ° 

By R. E. Coker, University of North Carolina 



xvi Contents 

PAGE 

Suggestions for Culturing Ostracods 228 

By Esther M. Patch and Lowell E. Noland, University of Wisconsin 

Cypridae 22 9 

By Charles H. Blake, Massachusetts Institute of Technology 

A Suggested Imitation of a Woodland Pool 229 

By Charles H. Blake, Massachusetts Institute of Technology 

Maintenance of Land Isopods 230 

By John L. Fuller, Massachusetts Institute of Technology 

Note on Keeping Ligia oceanka in the Laboratory . . . .231 
By John Tait, McGill University 

Hatching and Rearing Pandalid Larvae 231 

By Alfreda Berkeley Needler, Biological Board of Canada 

A Method for Rearing Small Crangonidae 233 

By Hugh H. Darby, College of Physicians and Surgeons 

Hatching and Rearing Larvae of the American Lobster, Homa- 

rus americanus 233 

By Paul S. Galtsoff, U. S. Bureau of Fisheries 

A Crayfish Trap (Reprinted) 236 

By E. C. O'Roke, University of Michigan 

Culture Methods for Marine Brachyura and Anomura . . .237 
By Josephine F. L. Hart, Pacific Biological Station 

Notes on Rearing the Pacific Edible Crab, Cancer magister . .239 
By Donald C. G. MacKay, Pacific Biological Station 

Class Arachnoidea 

Feeding Notes for Certain Arthropods 242 

By Lucy W. Clausen, American Museum of Natural History 

Laboratory Care of Tarantulas 243 

By W. J. Baerg, University of Arkansas 

Keeping Avicularia avicularia in the Laboratory . . . .244 
By Mary L. Didlake, University of Kentucky 

Culture of Lathrodectus mactans, the Black Widow Spider . .244 
By Elizabeth Burger, The College of William and Mary 



Contents xvii 

PAGE 

Culture of Non-Predacious, Non-Parasitic Mites .... 245 
By Arthur Paul Jacot, Asheville, North Carolina 

Tick Rearing Methods with Special Reference to the Rocky 

Mountain Wood Tick, Dermacentor andersoni Stiles . . 246 
By Glen M. Kohls, U. S. Public Health Service 

Parasitic Water Mites 256 

By John H. Welsh, Harvard University 

Breeding of Neotetranychus buxi, a Mite on Boxwood . . .257 
By Donald T. Ries, Ithaca, New York 

Class Myriapoda 

Euryurns erythropygus (Abstracted) 258 

By Hugh H. Mlley, Ohio State University 

Rearing of Scutigerella immaculata (Abstracted) . . . .259 
By George A. Filinger, Ohio Agricultural Experiment Station 

Class Insecta 

Rearing of Thysanura 259 

By G. J. Spencer, University of British Columbia 

Methods of Rearing Lepismatids 261 

By J. Alfred Adams, Iowa State College 

Rearing of Collembola 263 

By G. J. Spencer, University of British Columbia 

Remarks on Collembola (Abstracted) 263 

By Charles Macnamara, Arnprior, Ontario 

A Method for Rearing Mushroom Insects and Mites (Reprinted) . 265 
By C. A. Thomas, Pennsylvania State College 

Rearing Mayflies from Egg to Adult 266 

By Helen E. Murphy, Phoenix, N. Y. 

Culture Methods for the Damselfly, Ischnura verticalis . . .268 
By Evelyn George Grieve, Cornell University 

Methods of Rearing Odonata 270 

By P. P. Calvert, University of Pennsylvania 



xviii Contents 

PAGE 

Rearing the Stonefly, Nemoura vallicularia (Abstracted) . . .273 
By Chen-fu Francis Wu, Yen Ching University, Peiping, China 

Rearing Fall and Winter Plecoptera (Abstracted) . . . .274 
By Theodore H. Frison, Illinois Natural History Survey 

Laboratory Colonies of Termites 275 

By Esther C. Hendee, Limestone College, Gafjney, South Carolina 

Troctes divinatoria (Abstracted) 279 

By O. W. Rosewall, Louisiana State University 

Lipeurus heterographus (Abstracted) 280 

By F. H. Wilson, Tulane University 

Oligotoma texana (Abstracted) 281 

By Harlow B. Mills, Montana State College 

Dermaptera 282 

By B. B. Fulton, North Carolina State College 

Grylloblatta 282 

By Norma Ford, University of Toronto 

Care and Rearing of Blattella germanica 283 

By C. M. McCay and R. M. Melampy, Cornell University 

Cockroaches (Reprinted) 283 

By John M. Kelley, General Biological Supply House 

Two Species of Praying Mantis (Abstracted) 284 

By Mary L. Didlake, University of Kentucky 

Ceuthophilus (Abstracted) . . . . . . . .285 

By T. H. Hubbell, University of Florida 

On Rearing Gryllidae 286 

By B. B. Fulton, North Carolina State College 

Culture Methods for Grasshoppers 287 

By E. Eleanor Carothers, State University of Iowa 

The Grouse Locusts 292 

By Robert K. Nabours, Kansas State College 

Rearing Thrips tabaci (Abstracted) 295 

By Burl Alva Slocum, University of Nanking 



Contents xix 

PAGE 

Rearing Hog Lice on Man 296 

By Laura Florence, N. Y. Homeopathic Medical College 

Notes on Rearing a Scutellerid 298 

By H. M. Harris, Iowa State College 

A Method of Rearing Four Species of Plant Bugs . . . .299 
By F. G. Mundinger, New York State Agricultural Experiment Station 

Perillus bioculatus (Abstracted) 3°° 

By Harry H. Knight, Iowa State College 

Corizus hyalinus and C. sidae (Abstracted) 300 

By Philip A. Readio, Cornell University 

Lygaeidae 3 01 

By F. M. Wadley, U. S. Bureau of Entomology and Plant Quarantine * 

Rearing Methods for Chinch Bugs, BUssus hirtus .... 303 
By Kenneth E. Maxwell, Cornell University 

Reduv'ms personatus (Abstracted) 304 

By Philip A. Readio, Cornell University 

How to Rear Mesovelia 3°5 

By C. H. Hoffmann, U. S. Bureau of Entomology and Plant Quarantine 

A Method of Rearing Two Species of Nabidae 306 

By F. G. Mundinger, New York State Agricultural Experiment Station 

Breeding and Rearing Cimex lectularius 307 

By R. M. Jones, Liquid Carbonic Corporation, Chicago 

Hydrometra (Abstracted) 3° 8 

By J. R. de la Torre Bueno, Tucson, Arizona 

Three Species of Gerridae (Abstracted) 308 

By William E. Hoffmann, Lingnan University 

Winter Food for Waterbugs in Aquaria (Reprinted) . . 309 

By William E. Hoffmann, Lingnan University 

Velia watsoni (Abstracted) 3™ 

By William E. Hoffmann, Lingnan University 

Saldula major and S. pallipes (Abstracted) 3 11 

By Grace Olive Wiley 



xx Contents 

PAGE 

Rearing Notonectidae (Abstracted) 3 12 

By Clarence O. Bare, Sanford, Florida 

Pelocoris carolinensis (Abstracted) 3 X 3 

By H. B. Hungerford, University of Kansas 

Curie t a drakei (Abstracted) 3 X 3 

By Grace Olive Wiley 

Gelastocoris oculatus (Abstracted) 3 X 4 

By H. B. Hungerford, University of Kansas 

Rearing Cercopidae 3 X 5 

By Kathleen C. Doering, University of Kansas 

Artificial Feeding of Leafhoppers (Abstracted) 3 l6 

By R. A. Fulton and J. C. Chamberlin, U. S. Bureau of Entomology 
and Plant Quarantine 

The Beet Leafhopper, Eutettix tenellus (Abstracted) . . .317 
By Henry H. P. Severin, University of California 

Culture Methods for the Potato Psyllid 3 X 7 

By George F. Knowlton, Utah Agricultural Experiment Station 

A Useful Cage for Rearing Small Insects on Growing Plants 

(Abstracted) 3 20 

By E. A. Hartley, New York State College of Forestry 

The Nasturtium Aphid, Aphis rumicis 3 21 

By H. H. Shepard, University of Minnesota 

Aphis maidi-radicis (Abstracted) 3 2 3 

By J. J. Davis, Purdue University 

Rearing Methods for Aphididae 3 2 3 

By F. M. Wadley, U. S. Bureau of Entomology and Plant Quarantine 

Culture of Aphids 3 2 4 

By A. Franklin Shull, University of Michigan 

Methods for Rearing Mealybugs, Pseudococcus sp 326 

By Stanley E. Flanders, University of California 



Contents xxi 

PAGE 

Methods Used in Rearing the Mealybug, Pseudococciis com- 

stocki 329 

By W. S. Hough, Virginia Experiment Station 

Methods of Collecting and Rearing Neuroptera . . . .331 
By Roger C. Smith, Kansas State College 

Bittacus 335 

By Laurel R. Setty, Park College 

On Rearing Triaenodes 336 

By Wynne E. Caird, Cornell University 

A Method of Collecting Living Moths at Sugar Bait (Re- 
printed) 337 

By R. P. Gorham, Dominion Entomological Laboratory, Fredericton, N. B. 

A Method for Breeding Clothes Moths on Fish Meal . . .338 
By Grace H. Griswold, Cornell University 

The Goldenrod Gall-Maker, Gnorimo schema gallaesolidag'mis 

(Abstracted) 340 

By R. W. Leiby, North Carolina State Department of Agriculture 

Mass Production of Sitotroga cerealella 340 

By Stanley E. Flanders, University of California 

Sitotroga Egg Production 345 

By Stanley E. Flanders, University of California 

Notes on Breeding the Oriental Fruit Moth, Grapholitha molest a . 345 
By W. T. Brigham, Connecticut Agricultural Experiment Station 

Breeding Methods for Galleria mellonella 349 

By T. L. Smith, College of the Ozarks, Clarksville, Arkansas 

Laboratory Breeding of the European Corn Borer, Pyrausta 

nubilalis (Abstracted) 352 

By L. J. Briand, Parasite Laboratory, Belleville, Ontario 

Rearing Ephestta kuehniella Larvae in Quantity . . . -355 
By P. W. Whiting, University of Pennsylvania 

Rearing Polyphemus Moths . -357 

By R. W. Dawson, University of Minnesota 



xxii Contents 

PAGE 

Breeding Lymantriid and Saturniid Moths 359 

By William Tracer, Rockefeller Institute for Medical Research 

The Columbine and Iris Borers 361 

By Grace H. Griswold, Cornell University 

Methods for the Laboratory Culture of the Silkworm, Bombyx 

mori 362 

By William Trager, Rockefeller Institute for Medical Research 

Further Notes on Breeding Lepidoptera 364 

By W. T. M. Forbes, Cornell University 

Butterflies 365 

By John H. Gerould, Dartmouth College 

Craneflies 368 

By J. Speed Rogers, University of Florida 

Methods of Rearing, Manipulating, and Conserving Anopheline 

Imagines in Captivity 376 

By Mark F. Boyd, Thomas L. Cain, Jr., and J. A. Mulrennan, Station 
for Malaria Research, Tallahassee, Florida 

A Mosquito Rearing Cage 383 

By F. C. Baker, U. S. Bureau of Entomology and Plant Quarantine 

Laboratory Breeding of the Mosquitoes, Culex pipiens and 

C. jatigans 386 

By Clay G. Huff, University of Chicago 

The Culture of Mosquito Larvae Free from Living Micro- 
organisms 389 

By William Tracer, Rockefeller Institute for Medical Research 

Psychoda alternata and P. minuta (Abstracted) .... 390 
By C. L. Turner, Beloit College 

Methods of Collecting and Rearing Ceratopogonidae . . .391 
By Lillian Thomsen, Bethany College, Lindsborg, Kansas 

Methods for Propagation of the Midge, Chironomus tentans 

(Abstracted) 392 

By William O. Sadler, Mississippi College 

Chironomus crista tus (Abstracted) 395 

By Hazel E. Branch, University of Wichita 



Contents xxiii 

PAGE 

A Method for Studying the Hessian Fly and Other Insects 

(Abstracted) 396 

By James W. McColloch, Kansas State Agricultural Experiment 
Station 

Ceroplatinae and Macrocerinae (Abstracted) 398 

By G. H. Mansbridge, Imperial College of Science and Technology 

Culture Methods Used for Sciara 399 

By Helen B. Smith, Johns Hopkins University 

Culture of Sciara . 400 

By F. H. Butt, Cornell University 

A Laboratory Method for Rearing Sciara and Phorid Flies 

(Abstracted) 401 

By M. D. Austin and R. S. Pitcher 

Simulium omatum (Abstracted) 402 

By John Smart, University of Edinburgh 

Methods for Collecting and Rearing Horseflies 405 

By H. H. Schwardt, University of Arkansas 

The Culture of Aphidophagous Syrphid Flies 409 

By C. L. Fluke, JR-, University of Wisconsin 

Culture of the Drone Fly, Eristalis tenax 410 

By William L. Dolley, Jr., C. C. Hassett, W. B. Bowen, and George 
Phillies, University of Buffalo 

Methods Used in Rearing Leschenaultia exul . . . . . 411 
By Henry A. Bess, Ohio State University 

A New Technique for Rearing Dipterous Larvae (Translated) . .412 
By Julio Garcia-Diaz, University of Puerto Rico 

Sarcophaga bullata 412 

By Roy Melvin, U. S. Bureau of Entomology and Plant Quarantine 

Cochliomyia americana and C. macellaria 413 

By Roy Melvin, U. S. Bureau of Entomology and Plant Quarantine 

The Culture of Blowflies 414 

By Dwight Elmer Minnich, University of Minnesota 

Rearing Maggots for Surgical Use 418 

By G. F. White, U. S. Bureau of Entomology and Plant Quarantine 



xxiv Contents 

PAGE 

Rearing Larvae of the Cluster Fly, Pollenia rudis . . . .427 
By R. M. DeCoursey, Connecticut State College 

Stomoxys calcitrans 428 

By Roy Melvin, U. S. Bureau of Entomology and Plant Quarantine 

Rearing the Housefly, Muse a domestica, Throughout the Year . .429 

By Henry H. Richardson, U. S. Bureau of Entomology and Plant 
Quarantine 

The Culture of Muscina stabulans 432 

By Evelyn George Grieve, Cornell University 

Borboridae (Reprinted) 434 

By J. W. Wilson and Norman R. Stoll, Rockefeller Institute for 
Medical Research 

Notes on Breeding the Apple Maggot, Rhagoletis pomonella . . 436 
By Philip Garman, Connecticut Agricultural Experiment Station 

Piophila casei 437 

By Don C. Mote, Oregon State Agricultural College 

Culture Methods for Drosophila 437 

By A. H. Sturtevant, California Institute of Technology 

Pseudolynchia maura 446 

By Clay G. Huff, University of Chicago 

The Calosoma Beetle (Calosoma sycophanta) (Abstracted) . . 447 

By A. F. Burgess and C. W. Collins, U. S. Bureau of Entomology and 
Plant Quarantine 

Breeding and Rearing Haliplidae (Abstracted) .... 448 
By Jennings R. Hickman, Michigan State Normal School 

Rearing Gyrinidae (Abstracted) 45° 

By Melville H. Hatch, University of Washington 

Hydrophilidae (Abstracted) 45 x 

By E. Avery Richmond, Pennsylvania State College 

Silpha inaequalis (Abstracted) 45 2 

By Milton T. Goe, Portland, Oregon 

Staphylinidae (Abstracted) 453 

By Helen G. Mank, Lawrence, Massachusetts 



Contents xxv 

PAGE 

Batrisodes globosus (Abstracted) 454 

By Orlando Park, Northwestern. University 

Rearing Methods for Wireworms 455 

By W. A. Rawlins, Cornell University 

Scirtes tibialis (Abstracted) 457 

By Walter C. Kraatz, University of Wisconsin 

Breeding Dermestes vulpinus Throughout the Year (Ab- 
stracted) 45 8 

By A. G. Grady, Research Laboratories, Rohm & Haas Co., Inc. 

Carpet Beetles 459 

By Grace H. Griswold, Cornell University 

Mycotretus pulchra (Abstracted) 460 

By Harry B. Weiss, New Jersey State Experiment Station 

Hyper as pis lateralis (Abstracted) 460 

By H. L. McKenzie 

Hippodamia 13-Punctata (Abstracted) 461 

By Clifford Cutright, Ohio Agricultural Experiment Station 

Breeding and Rearing the Mexican Bean Beetle, Epilachna 

corrupta 4 DI 

By S. MarcovitcH) University of Tennessee 

Lindorus lophanthae (Abstracted) 462 

By Stanley E. Flanders, University of California 

Mealworms, Blapstinus moestus and Tenebrio molitor . . . 463 
By William LeRay and Norma Ford, University of Toronto 

The Culture of Tribolium conjusum 4°3 

By Thomas Park, School of Hygiene and Public Health 

A Method of Observing the Development of Tribolium con- 
jusum 466 

By Herbert S. Hurlbut, Cornell University 

Tenebrio Culture 4°7 

By W. M. Mann, National Zoological Park 

Tenebrio obscurus (Abstracted) 4^7 

By Richard T. Cotton, U. S. Bureau of Entomology and Plant Quarantine 



xxvi Contents 

PAGE 

Cisidae (Abstracted) 467 

By Harry B. Weiss, New Jersey State Experiment Station 

Methods of Breeding and Rearing Scarabaeidae .... 468 

By Henry Fox, V. S. Bureau of Entomology and Plant Quarantine, and 
Daniel Ludwig, University College, New York University 

A Successful Method of Rearing Trichiotinus 473 

By C. H. Hoffmann, U. S. Bureau of Entomology and Plant Quarantine 

Passalus cornutus (Abstracted) 474 

By Warren C. Miller, Bedford High School, Bedford, Ohio 

The Painted Hickory Borer, Cyllene caryae (Abstracted) . . 475 
By E. H. Dusham, Pennsylvania State College 

Calligrapha pnirsa (Abstracted) 476 

By C. N. Ainslie, U. S. Bureau of Entomology and Plant Quarantine 

Method of Rearing Diabrotica duodecimpunctata, the Southern 

Corn Root Worm 477 

By D. L. Wray, North Carolina State College 

Rearing Stored Food Insects for Experimental Use . . . . 47 8 
By L. Pyenson and H. Menusan, Jr., Cornell University 

The Duckweed Weevil, Tanysphyrus lemnae ..... 480 
By Minnie B. Scotland, New York State College for Teachers 

Boll Weevil Culture 481 

By F. A. Fenton, Oklahoma A. and M. College 

Grain Weevils 481 

By D. L. Lindgren, University of Minnesota 

Method of Rearing Calandra callosa, the Southern Corn Bill Bug . 483 
By D. L. Wray, North Carolina State College 

Method for Rearing Scolytus multistriatus 484 

By Philip A. Readio, Cornell University 

Pityogenes hopkinsi (Abstracted) 484 

By James A. Beal 

A Method for Breeding Cephidae 485 

By Donald T. Ries, Ithaca, N. Y. 



Contents xxvii 

PAGE 

A Method of Breeding Some Defoliators ... . 486 

By S. A. Graham, University of Michigan 

Technique of Culturing Habrobracon juglandis 489 

By P. W. Whiting, University of Pennsylvania 

Rearing Chelonus annulipes 49 1 

By Arlo M. Vance, U. S. Bureau of Entomology and Plant Quarantine 

Rearing Apanteles thompsoni 49 2 

By Arlo M. Vance, U. S. Bureau of Entomology and Plant Quarantine 

Methods of Producing Macrocentrus ancylivorus in Large 

Numbers for Colonization in Peach Orchards .... 493 
By Phhip Garman, Connecticut Agricultural Experiment Station 

Rearing of Aphidiinae -495 

By Esther W. Wheeler, University of North Dakota 

Hymenopterous Parasites of Gyrinidae (Abstracted) . . . 496 
By F. Gray Butcher, North Dakota College of Agriculture 

Rearing Paniscus . . . . , 497 

By Arlo M. Vance, U. S. Bureau of Entomology and Plant Quarantine 

Culture of Habrocytus cerealellae, a Parasite of the Angoumois 

Grain Moth 497 

By B. B. Fulton, North Carolina State College 

Breeding a Primary Parasite and Two Hyperparasites of the 

Geranium Aphid 499 

By Grace H. Griswold, Cornell University 

Production of Trichogramma S 00 

By Stanley E. Flanders, University of California 

Methods for Rearing Tiphiids and Scoliids 502 

By J. L. King, U. S. Bureau of Entomology and Plant Quarantine 

Laboratory Maintenance and Care of the Mound-Building Ant, 

Formica ulkei 508 

By A. M. Holmquist, St. Olaf College, Northfield, Minnesota 

Some Aids to the Study of Mound-Building Ants . . . .510 
By E. A. Andrews, Johns Hopkins University 



xxviii Contents 

PAGE 

Methods of Breeding Perisierola angulata, a Cocoon Parasite 

of the Oriental Fruit Moth 512 

By J. C. Schread, Connecticut Agricultural Experiment Station 

Rearing Laelius anthrenivorus 513 

By Arlo M. Vance, U. S. Bureau of Entomology and Plant Quarantine 

Culture of Aphelopus theliae 514 

By S. I. Kornhauser, University of Louisville Medical School 

Anteonine Parasites of Leaf hoppers (Abstracted) . . . .515 
By F. A. Fenton, Oklahoma A. and M. College 

Methods of Rearing Wild Bees, Wasps, and Their Parasites . .516 
By Charles H. Hicks, University of Colorado 

Phylum XV. Mollusca 

Class Amphineura 

Chaeto pleura apiculata 519 

By Benjamin H. Grave, De Pauw University 

Class Gastropoda 

Lymnaea [=Pseudosuccinea] columella (Abstracted) . . . 520 

By Harold S. Colton and Miriam Pennypacker, University of Penn- 
sylvania 

Rearing Aquatic Pulmonate Snails 522 

By Elmer Philip Cheatum, Southern Methodist University 

Rearing Aquatic Snails 523 

By Wendell Krull, U. S. Bureau of Animal Industry 

Rearing Terrestrial Snails 526 

By Wendell Krull, U. S. Bureau of Animal Industry 

Vivarium Methods for the Land Mollusca of North America . . 527 
By A. F. Archer, University of Michigan 

Culture Methods for Umax flavus 5 2 9 

By Emmett B. Carmichael, University of Alabama School of Medicine 

The Genus Crepidula 53 * 

By E. G. Conklin, Princeton University 

Culture Methods for Urosalpinx cinerea 532 

By H. Federighi, Antioch College 



Contents xxix 

PAGE 

Class Pelecypoda 
Spawning and Fertilization of the Oyster, Ostrea virginica . -537 
By Paul S. Galtsoff, U. S. Bureau of Fisheries 

The Cultivation of Lamellibranch Larvae 539 

By Herbert F. Prytherch, U. S. Bureau of Fisheries 

Cumingia tellinoides 543 

By Benjamin H. Grave, De Pauw University 

Rearing Teredo navalis 545 

By Benjamin H. Grave, De Pauw University 

Phylum XVI. Echinodermata 
Class Aster oidea 

Asterias forbesi 547 

By Henry J. Fry, Cornell University Medical College 

The Laboratory Culture of the Larvae of Asterias jorbesi . . . 550 
By Evert J. Larsen, U. S. Bureau of Fisheries 

Class Ophiuroidea 

Ophioderma brevispina 553 

By Caswell Grave, Washington University 

Class Echinoidea 

Arbacia punctulata 554 

By Henry J. Fry, Cornell University Medical College 

Notes on the Culture of Strongylocentrotus jranciscanus 

and Echinarachnius excentricus 55& 

By Martin W. Johnson, Scripps Institution of Oceanography 

Class Holothurioidea 

Notes on the Culture of Cucumaria 559 

By Martin W. Johnson, Scripps Institution of Oceanography 

Phylum XVII. Chordata 
Class Ascidiacea 
Notes on the Culture of Eight Species of Ascidians . . . . 560 
By Caswell Grave, Washington University 

Culture Methods for Ascidians 5°4 

By N. J. Berrhl, McGill University 



CONTRIBUTORS 



This list includes the names of those only who prepared material 

especially for publication in this volume. The names of 

other authors will be found in the index. 



Ackert, J.E. 
Adams, J. Alfred 
Andrews, E. A. 
Archer, A. F. 
Baerg,W.J. 
Bahrs, Alice M. 
Baker, F. C. 
Banta, A. M. 
Barker, H. Albert 
Beers, C. Dale 
Berrill,N.J. 
Bess, Henry A. 
Blake, Charles H. 
Blount, Raymond F. 
Bond, R.M. 
Bowen.W.B. 
Boyd, Mark F. 
Brandwein, Paul 
Brigham.W.T. 
Briscoe, M. S. 
Brown, M. G. 
Burger, Elizabeth 
Butt, F. H. 
Butts, H. E. 
Cain,T.L., Jr. 
Calvert, P. P. 
Carmichael, E. B. 
Carothers, E. Eleanor 
Cederstrom, J. A. 
Chandler, Asa C. 
Cheatum, Elmer P. 
Christenson, Reed O. 
Clarke, George L. 
Clausen, Lucy W. 
Coe, Wesley R. 
Coker, R. E. 
Conklin, E. G. 
Danielson, R. N. 
Darby, Hugh H. 
Davis, Donald W. 
Davis, Mary E. 
Davis, W. H. 
Dawson, R. W. 



DeCoursey, R. M. 
Didlake, Mary L. 
Dobrovolny, C. G. 
Doering, Kathleen C. 
Dolley.W.L.Jr. 
Federighi, H. 
Fenton, F. A. 
Ferris, Josephine C. 
Finley, Harold E. 
Flanders, Stanley E. 
Florence, Laura 
Fluke, C.L., Jr. 
Forbes, W. T. M. 
Ford, Norma 
Fox, Henry 
Fry, Henry J. 
Fuller, John L. 
Fulton, B.B. 
Galtsoff, Paul S. 
Garman, Philip 
Gerould, J.H. 
Giese, A. C. 
Gilchrist, F. G. 
Graham, S. A. 
Grave, Benjamin H. 
Grave, Caswell 
Grieve, Evelyn G. 
Griswold, Grace H. 
Guberlet, John E. 
Hall, R. P. 
Halsey, H. R. 
Harris, H. M. 
Hart, Josephine F. L. 
Hasler, Arthur D. 
Hassett, C. C. 
Heath, Harold 
Hegner, R.W. 
Hendee, Esther C. 
Hess, Margaret 
Hess, Walter N. 
Hetherington, Alford 
Hicks, C. H. 
Hoffmann, C. H. 



Hogue, M. J. 
Holmquist, A. M. 
Hopkins, D. L. 
Hough, W.S. 
Huff, Clay G. 
Hurlbut.H.S. 
Hyman, Libbie H. 
Jacot, Arthur Paul 
Jahn, Theodore L. 
Johnson, M. W. 
Jones, Edgar P. 
Jones, R.M. 
Kepner, William A. 
King, J. L. 
Knowlton, G. F. 
Kofoid, Charles A. 
Kohls, Glen M. 
Kornhauser, S. I. 
Krull, Wendell 
Larsen, E. J. 
La Rue, George R. 
LeRay, William 
Lindgren, D. L. 
Lotze, John C. 
Ludwig, Daniel 
Lutz, F. E. 
MacGinitie, G. E. 
MacKay, Donald C.G. 
Mann, William M. 
Marcovitch, S. 
Maxwell, Kenneth E. 
McCay, C. M. 
McNeil, Ethel 
Melampy, R. M. 
Melvin, Roy 
Menusan, H., Jr. 
Miller, E. D. 
Minnich, D. E. 
Moore, J. Percy 
Mote, Don C. 
Mulrennan, J. A. 
Mundinger, F. G. 
Murphy, Helen E. 



xxxi 



XXX11 



Contributors 



Myers, Earl H. 
Nabours, Robert K. 
Needham, J. G. 
Needier, Alfreda B. 
Novikoff, Alex B. 
Nuttycombe, John W. 
Pace, D. M. 
Packard, Charles Earl 
Park, Thomas 
Patch, Esther M. 
Penn, Amos B. K. 
Phillies, George 
Prytherch, H. S. 
Pyenson, L. 
Rawlins, W. A. 
Readio, Philip A. 
Reynolds, Bruce D. 
Rice, N. E. 
Richardson, H. H. 



Ries, Donald T. 
Rogers, J. Speed 
Rogick, Mary 
Schluchter, Alfred W. 
Schread, J. C. 
Schwardt, H. H. 
Scotland, Minnie B. 
Setty, Laurel R. 
Shepard, H. H. 
Shull, A.F. 
Smith, George A. 
Smith, Helen B. 
Smith, Roger C. 
Smith, T. L. 
Snider, George G. 
Sonneborn, T. M. 
Spencer, G. J. 
Stirewalt, M. Amelia 
Stump, A. B. 



Stunkard, H. W. 
Sturtevant, A. H. 
Tait, John 
Taylor, C. V. 
Thomas, J. O. 
Thomsen, Lillian 
Trager, William 
Turner, John P. 
Vance, A. M. 
Vaughan, Thomas W. 
Wadley, F. M. 
Welch, P. S. 
Welsh, John H. 
Wheeler, Esther W. 
White, G. F. 
Whiting, P. W. 
Wilson, H. V. 
Wray, D.L. 
Wulzen, Rosalind 



CULTURE METHODS 

FOR 

INVERTEBRATE ANIMALS 



INTRODUCTION 

By James G. Needham 

THE needs of animals determine all successful rearing practices. 
Their basic requirements are four: (i) Food; (2) Protection 
from enemies; (3) A suitable physical environment: these 
for individual livelihood; and also for the maintenance of 
successive generations; (4) Fit conditions for reproduction. 

Culturing animals doubtless began with collecting and caring for 
living specimens, and the first suggestions for supplying their needs in 
captivity were gained (as they are still to be gained) by carefully ob- 
serving them in their natural habitat. There they are seen eating their 
food, constructing their homes, eluding their enemies, accepting their 
mates, and rearing their offspring. There is no better way to proceed in 
the beginning than by imitating natural conditions. Our methods 
must be adapted to the .ways of the animal, for only in a very small 
measure will it change its ways for ours. Especially in the reproductive 
habits will it show readiness to go its hereditary way, and stubborn 
refusal to go any other. We may learn by experiment how best to 
meet its needs under indoor conditions. It is not so much close imita- 
tion of the natural environment, as careful feeding and attention to 
hygienic needs that make for permanent maintenance. Artificial devices 
may replace and may even better those found in nature (witness the 
movable-frame beehive as compared with the hollow tree), but the basic 
requirements of the animal remain ever the same. 

This book is concerned with methods of management of animal 
cultures under control. In the following pages will be found, first some 
general suggestions covering the principles of culture management by 
members of the Committee, followed by more specific and detailed 
methods for rearing particular groups or species, written by many 
individual contributors and collaborators. As stated in our call for 
such materials (Science 77:427, 1933), we have sought to obtain for at 
least one species of each considerable group of invertebrates "a fairly 
complete account of maintenance requirements, covering collecting 
methods and devices, cages and breeding quarters, plans for feeding 
and watering, cleaning and aerating quarters, breeding management, 
and all else that enters into the maintenance of the species through 
successive generations." When such an account was not available we 
have welcomed scraps of information that seemed likely to be helpful 
toward culture-keeping. The contributed articles vary therefore from 



4 Introduction 

mere suggestions as to how to get living specimens, to measured pro- 
cedures quantitatively determined. We have filled some of the more 
obvious gaps with abstracts and reprintings (initialed by the editor 
responsible) from available literature but we have by no means ex- 
hausted this source of material. 

The contributed articles are arranged as far as possible in systematic 
order following Pratt's Manual oj the Common Invertebrate Animals 
(revision of 1935), with the orders of insects following that of Com- 
stock's Manual jor the Study oj Insects (1926 edition) . Material within 
the insect orders is arranged according to the N. Y. State List oj Insects. 

Since from the nature of the book the different types of procedure 
are widely scattered through its pages, we have tried to make an index 
that would serve as a finding list for them, and we recommend that the 
user consult the index freely. 

We have assumed on the part of the reader some acquaintance with 
general zoology and some knowledge of elementary laboratory tech- 
nique. Limitations of space have necessitated that we restrict the text 
rather closely to collecting and culture methods. In doing so we have 
had to omit some interesting and valuable material, principally intro- 
ductory remarks, systematic discussion, life history details, and sugges- 
tions for the use of the materials in special fields. We have also done 
some condensing to avoid undesirable duplications, and we hope that in 
so doing we have not been unfair to any of our helpfully minded col- 
laborators. Papers by members of the scientific staffs of federal depart- 
ments and bureaus have come to us each bearing a statement that it is 
offered for publication with the permission of the head of the Depart- 
ment concerned. Receipt of these statements is hereby acknowledged; 
but the statements themselves are omitted from the text to make room 
for more useful material. 

It is hoped that this compilation on culture methods may stimulate 
interest in maintaining living animals in biological laboratories and may 
lead to further development of the proper technique. We will be glad 
to hear comments from the users of this book upon the usefulness of the 
methods here offered and to learn of new developments in the use of 
them; for it seems highly probable that both eliminations and improve- 
ments resulting from further trials will make necessary an early revision 
of this compendium. 



General Methods of Collecting, 
Maintaining, and Rearing 

Marine Invertebrates in the Laboratory 

j 

Paul S. Galtsoff, U. S. Bureau of Fisheries 

COLLECTING 

THE selection of the equipment for collecting marine animals is gov- 
erned by various considerations, of which the character of the bot- 
tom, depth of water, number of specimens the investigator desires to 
obtain, the purpose of collecting, and the animal which he seeks, are of 



Gin 




Fig. i. — The dip net. 

paramount importance. The following account describes the instru- 
ments which may be needed by an individual collector who desires to 
bring live material to his laboratory. The description of the method of 
collecting large numbers of specimens for museums and supply houses, as 
well as the account of the various oceanographic instruments used in a 
quantitative study of ocean life are beyond the scope of this book. The 
reader interested in this matter is referred to such books as: Murray 
and Hjort (1912); Johnstone (1908); Bulletin No. 85, Oceanography, 
of the National Research Council (1932) ; numerous publications of the 
Conseil Permanent pour I' Exploration de la Mer ; Wissenschajtliche 
Meeresuntersuchungen (abt. Helgoland and abt. Kiel), and to the de- 
scriptions of equipment given in the reports of various oceanographic 
expeditions. 

The dip net. The dip net (Fig. 1) is the handiest and most indis- 
pensable piece of equipment that can be used for many purposes and 
under a great variety of conditions. It consists of a conical net bag at- 
tached to a stout ring made of galvanized iron or preferably of brass and 



6 Marine Invertebrates 

fixed to a wooden handle. The bag usually measures about i foot in 
diameter at the opening and 18 inches in depth. The netting may vary 
from i inch mesh at the top with % inch at the bottom for larger speci- 
mens, to % of an inch at the top with % inch mesh at the bottom for 
smaller forms. For collecting minute organisms a bag made of bolting 
silk Nos. 12, 16, or 20 may be substituted. The handle is usually from 
6 to 7 feet long, but of course may be increased to any desired length. 




Fig. 2. — The pile scrape net. 

For picking up small specimens floating on the water a small dip net 
about 6 inches in diameter is sometimes preferable. 

The pile scrape net. This modification of the dip net is used for col- 
lecting organisms growing on piling and other underwater structures. 
The metal ring of the net is bent in such a way as to make it fit the curva- 
ture of the piling (Fig. 2) ; it is about % 6 °f an mcn m thickness and 12 



K-,— Ut -- It^t 




Fig. 3. — The square scrape net. 



inches in diameter with the curved scraper 1 inch in width. Attached 
to this frame is a net 18 inches in depth with the mesh % of an inch at 
the top and % of an inch or less at the bottom. The metal blade which 
is welded to the lower part of the iron frame works as a cutting knife 
when the net is pressed against the piling and is pulled up. The length 
of the handle varies according to local conditions from 6 to 20 feet. 

The square scrape net. This type is a modification of the pile scrape 
net from which it differs only in the shape of the frame, which is not bent 
(Fig. 3) as it is in the former type, and is provided with a straight 
cutting blade attached to the base. This net is very useful in collecting 
organisms growing on walls of various underwater structures such as 



Collecting 7 

stone breakwaters and docks. It may be used also for obtaining speci- 
mens living on hard or sandy bottoms. 

The oyster tongs. Oyster tongs, generally used by fishermen for tak- 
ing oysters from shallow water, may be used for collecting other bottom 
animals. This much used implement (Fig. 4) is made of a pair of rakes 
attached to the lower ends of two 
long handles fastened together like 
the blades of shears. The rakes are 
so fitted together that they rest 
upon the bottom parallel to each 
other when the handles are spread. 
By bringing the handles together 
the instrument is closed and the 
teeth of each rake interlock. The 
rakes are about 14 inches wide with 
the teeth 3-4 inches long and placed 
about 1% inches apart. Oyster 
tongs are usually available in four 
sizes, 8, ic, 12, and 16 feet long, 
although 20 foot handles are some- 
times used by the fishermen. Tong- 
ing may be efficiently carried out 
even by an inexperienced collector 
in water less than 12 feet deep. 

The digger. The most universal 
way of obtaining mollusks, worms, 
and other forms inhabiting muddy 
bottoms, is with rakes. The sim- 
plest type is the ordinary potato 
digger (Fig. 5) with four to six long 
thin prongs, and fitted with a handle about 5 feet long. The digger may 
be conveniently used on the exposed tidal flats. For working under 




Fig. 4. — The oyster tongs. 




Fig. 5. — The digger. 



8 



Marine Invertebrates 



water the back of the digger is covered with wire netting which holds the 
animals caught by the prongs. This instrument is commonly used for 
digging hard clams (Venus) and other mollusks. 

The basket rake. By fastening a basket of wire netting to the ordinary 
garden rake (Fig. 6) a very useful instrument may be made which is 




Fig. 6. — The basket rake. 

operated either by wading or from a boat. The shape and dimensions 
of the basket and the mesh of the wire netting vary greatly depending 
upon the locality and depth of the water in which the implement is to 
be used. The type of basket rake used for collecting Venus in the deep 
water of Cape Cod consists of an iron framework forming a curved bowl, 
the under edge of which is set with 20 steel teeth about 2% inches in 
length (Fig. 7). The bowl of the rake, strengthened by side and cross 




Fig. 7. — The basket rake, Cape Cod type. 



pieces of iron, is covered with a twine net dragging behind it. Sometimes 
teeth about 4 inches long are used. The rake weighs from 15 to 20 
pounds. There is great variety in the styles and sizes of basket rakes 
used by fishermen. The handles of the rakes vary from 23 to 65 feet 
according to the prevailing depth of the water. The rakes are often pro- 
vided with several detachable handles of various lengths. Although the 
handles are made of strong wood they break very easily when operated 
by inexperienced collectors, since they are flexible and very thin, not 
exceeding 1% inches in diameter. Operation of long rakes (65 feet) 
requires great skill. 



Collecting 9 

The clam hoe and the hooker. These instruments are used for digging 
in shallow water or on the exposed sand or mud flats. The clam hoe 
(Fig. 8) has four prongs about i 1 ^ inches wide and from 12 to 14 inches 
long, and a strong wooden handle about 4 feet in length. The instru- 
ment is suitable for digging in coarse sand or gravel. The hooker 





Fig. 8. — The clam hoe. 



Fig. 9. — The hooker. 



(Fig. 9) used in digging in the hard mud, has four thin, sharp prongs and 
a short handle. 

The shovel. An ordinary steel shovel is a valuable tool for collecting 
animals living in sand or mud on the beaches or in shallow water. 

The dredge. The dredge is the most efficient instrument for collecting 
bottom dwelling forms regardless of the depth of the water. Dredging 
at great depths is a difficult operation requiring costly equipment, but 
dredging in shallow water not exceeding 100 feet, may be carried out 
from a small boat and does 
not require special machinery. 
There are many types and sizes 
of dredges, ranging from small 
instruments about 1 foot wide 
to large commercial oyster 
dredges several feet wide and 
having a capacity of over 25 
bushels. The description given 
here refers only to small in- 
struments that may easily be used by the collector of scientific material. 

The dredge is always made of a rigid iron frame to which a bag made 
of heavy netting or interwoven chain rings is attached. The most com- 
monly used type is the so-called scraper or scallop dredge (Fig. 10) 
which consists of a triangular framework with an iron blade (B), 2 inches 
wide, set at an angle so as to dig into the bottom. On the upper side a 
raised cross bar connects the two arms. The net with a wooden hori- 
zontal bar at the end is fastened to the cross bar and to the top of the 
blade. Additional weight (A) may be put on the cross bar if it is desired 
that the instrument cut deeper into the bottom. The dredge shown in 
figure 10 has a metal sheet (A) which serves the double purpose of pro- 




Fig. 10. — The scallop dredge. 



10 



Marine Invertebrates 




Fig. ii. — The box dredge. 



viding additional weight and pushing the material scraped by the blade 
into the bag. The usual dimensions are as follows: arms, 2% feet; cross 
bar, 2 feet; blade, 2% feet long and 2 inches wide. For operating on 
rough bottoms the blade is set level or even with a slight upward incline, 
so the dredge will slide over the bottom. A dredge of this type may be 
made of any size and is very useful for general collecting. For small 

organisms the bags must be 
made of a fine twine netting 
or of some other strong, 
coarse material of the de- 
sired mesh. 

Instead of a cutting blade 
the dredge may be provided 
with a set of teeth. This 
type is commonly used for dredging oysters. 

The box dredge. The box dredge (Fig. n) consists of a rectangular 
iron framework 27 x 12 inches, with two folding arms and two cutting 
blades, one on each side of the dredge. A bag of coarse netting is at- 
tached to the blades. When in operation the two arms are tied together 
by a piece of string and the drag line is fastened only to one arm. If the 
dredge is caught under rocks the string breaks and the instrument may 
be saved by dragging it sidewise by one arm. This small dredge is very 
useful for general collecting. 

The triangular dredge. The triangular dredge (Fig. 12) has some ad- 
vantages over the other types because 
no matter which side rests on the 
bottom one of the blades will cut into 
the ground when the instrument is 
dragged. 

To facilitate the finding of a 
dredge in case the drag line snaps, a 
tail buoy is attached with a length of 
rope slightly greater than the depth 
of the water. When the instrument is being dragged slight shocks caused 
by the impact of the frame with rocks or other objects are conveyed 
along the rope and are easily noticed by the operator holding it in his 
hands or only touching it. He can easily feel the change in vibration 
when the dredge slides over the bottom without cutting into it. In this 
case more rope should be given out or the speed of the boat reduced. 

The material collected in the dredge must be washed free of mud and 
sorted. For this purpose it is convenient to have a set of sieves with 
various meshes into which the contents of a dredge is dumped and washed 
by dipping into the sea. (See p. 534 for drill trap dredge.) 




Fig. 12. — The triangular dredge. 
After Hagmeier. 



Collecting 1 1 

The tangle or mop. This implement (Fig. 13) may easily be made 
by attaching long loose cotton strands to an iron bar. When trailed along 
the bottom it captures echinoderms and spiny crustaceans with which it 
may come in contact. The tangle is widely used in Long Island Sound 
by the oystermen for removing starfish from their oyster bottoms and is 
available in various sizes and styles. 

The grapple. The grapple (Fig. 14) consists of a number of steel 
wires passed through a galvanized pipe about one foot long and 1% 





Fig. 13. — The tangle. 



Fig. 14. — The grapple. 



inches in diameter, the inside of which is filled with lead. The lower 
ends of the wires are bent back to form hooks, the upper being twisted to 
make a loop for the attachment of a line. The grapple is very useful 
for collecting submerged vegetation which may contain rich fauna of 
crustaceans, worms, Bryozoa, etc.* 

The plankton net. Small organisms suspended in the water are col- 
lected by means of a plankton net (Fig. 15) which is towed behind the 
boat or is allowed to sink and is then slowly hauled up. To avoid back- 
washing of the material caught in the net the rate of towing or hauling 
should not exceed 1 meter per second. Plankton nets may be made of 
various grades of bolting silk depending upon the size of the organisms 
one is planning to catch. For small planktonic forms No. 20 or 25 

♦Editor's Xote: Lacking a grapple, a substitute for it may be made by coiling a long 
piece of barbed wire in a circle, fastening it at the overlaps, and attaching a weight 
at one side and a throwline at the other. This will gather submerged vegetation effectively. 
J. G. N. 



12 



Marine Invertebrates 



should be used. It is advisable to have the silk part of the net sewn on a 
canvas collar folded over the metal ring and fastened to it by means 
of buttons. The lower end of the net is also made of 
a canvas collar which is slipped over the metal bucket 
and fastened to it by a clamp ring. The bucket has 
windows covered with bolting silk and is provided with 
a stop cock for draining. A small glass bottle or jar 
may be used instead of a bucket. Nets one foot in 
diameter are the most convenient ones to handle from 
a small boat. 

The plankton trawl. Planktonic forms living just 
above the bottom may be collected by means of a 
plankton net mounted in a horizontal position on a 
frame attached at right angles to a sheet of galvanized 
iron. When the plankton trawl (Fig. 16) is dragged it 
slides over the bottom and catches the organisms which 
otherwise escape capture. The metal sheet protects 
the net which does not come in contact with the bot- 
tom and therefore may be made of fine bolting silk. 
The glass bottomed box. In shallow waters the ob- 
servation and collection of bottom animals are greatly 
facilitated by using a water tight box with a pane of 
plate glass fitted in the bottom. The dimensions and 
shape may vary to suit individual purposes. The box 
used by the author is 1 1 inches high with square bottom and top, 9x9 
and n x 11 inches respectively (Fig. 17). It is fastened to a boat by 
a short line and is placed in the water with the glass side down. It 
smooths the ruffled surface of the water. 

Goggles. Goggles (Fig. 18) are widely used in the Orient by pearl- 




Fig. 15. — The 
plankton net. 




Fig. 16. — The plankton trawl. 

oyster fishermen and may be used to aid in the collection and observation 
of shallow water animals inhabiting the ocean floor. Those with wooden 
frames are the most convenient for they may be carved to fit the eye- 
sockets. Condensation of vapor on the inside may be avoided by follow- 
ing the method of the Philippine divers who rub the glass with tobacco 



Marine Aquaria 13 

soaked in water. Goggles of the type shown may be obtained in Hono- 
lulu in almost every store handling fishing tackle and other sporting 

goods. 

The diving helmet. The diving helmet, very useful in warm and clear 
waters, has become a part of the regular equipment of tropical marine 
stations. It may not be used by an individual collector, for the opera- 
tion requires a crew of at least two, and preferably three men to operate 
the pump and watch the diver. For an untrained person it is inadvis- 
able to descend beyond the 30 foot limit. 

Miscellaneous equipment. An experienced collector never forgets a 





Fig. 17.— The glass bottomed box. Fig. i 8.— Goggles. 

pocket lens of about 8 or 12 power magnification, mounted in a metal 
frame and suspended from the neck by means of a cord. 

For collecting among coral reefs and rocks a good crow bar and cold 
chisel are indispensable. 

Living forms should be placed immediately in a suitable container 
about % full of seawater. An ordinary milk can, preferably of white 
porcelain, is very convenient for this purpose. Small organisms may be 
placed in fruit jars, bottles, or vials. Overcrowding and exposure to 
direct sunlight should be avoided. All the containers should be kept 
open as long as possible. In a hot climate keeping the container packed 
in ice is sometimes necessary if the organisms are in transport for several 
hours. 

MARINE AQUARIA 

Aquarium tanks. Various types of aquarium tanks found on the 
market may be used successfully for keeping live material. A simple 
water-tight box with an inlet and outlet for seawater may be used for 
this purpose. A coat of black asphalt paint provides sufficient protec- 



14 Marine Invertebrates 

tion for the wood and is perfectly harmless even to the most delicate 
forms.* It may be constructed according to the desired size and shape. 
So-called "white wood" (trade name) untreated in any way but ab- 
solutely dry should be used. All wooden parts must be painted sep- 
arately before they are put together and a second coat of paint (No. 24 
Scotch heater, manufactured by Billing and Chapin Co., N. Y.) is ap- 
plied after the aquarium is assembled. To prevent cracking of the 
wood, the aquaria must be kept moist all the time. 'For mounting the 
glass a type of putty containing no toxic substances and remaining plastic 
for a long period should be used. Putties that harden quickly should 
be avoided for they may exert uneven pressure on the glass wall and 
cause it to crack.** 

For more critical experimental work in which a complete elimination 
of foreign substances is important, only glass containers should be used. 
All kinds of glassware found on the market may be used for the cul- 
tivation of marine forms. Fruit jars, battery jars, precipitating cylinders 
and more expensive pyrex containers of various shapes and dimensions 
may be suitable depending upon the requirements of the investigator. If 
running seawater is not needed small and medium-sized organisms may 
be kept successfully in ordinary finger bowls, 4 inches in diameter, or 
in so-called specimen dishes, 7 inches in diameter. These dishes made 
of heavy glass may be placed one on top of another and are very easy to 
handle. 

For delicate physiological work, as for instance the experiments on 
fertilization, only the best grade of glass or even quartz should be used. 
A cheap type of glassware is usually more or less discolored and of 
uneven thickness. These defects make it unsuitable for photography or 
for the observation of organisms through a low powered microscope. 
For these purposes a better type of glassware with parallel walls should 
be selected. For the cultivation of diatoms, other marine unicellular 
algae, and flagellates small round flasks are usually used. 

Success in cultivating marine organisms depends not so much on the 
shape and size of the container used as on the cleanliness of the glass. 
This fact cannot be overemphasized, for many observations have been 
ruined because of the contamination of the glass by toxic substances the 
presence of which was not suspected. It is a safe rule in experimenting 
with living forms never to use glassware which has been previously em- 

* If it is intended to keep the organisms under observation, a glass walled aquarium 
should be used. 

** The following recipe developed by Prof. Petrunkevitch was found to be excellent: 
place a portion of spar varnish in a can, stir in screened Portland cement until quite 
thick. Do not add more cement until the combined mass has stood one or two hours. 
As the mixture thickens in standing, the consistency should be nearly that of putty. Keep 
in tightly covered cans. 



Marine Aquaria 15 

ployed in the laboratory for some other purpose and may have been in 
contact with such toxic substances as corrosive sublimate, picric acid, 
salts of chromic acid, or formalin. There is a considerable difference 
in the degree of absorption of poisons by glass, corrosive sublimate for 
instance being the most difficult to remove. When chromium cleaning 
fluid has been used glassware must be meticulously washed to remove 
traces of chromium salts. Glassware available in marine laboratories 
should always be regarded with a certain degree of suspicion because 
of the impossibility of ascertaining the purposes for which it was pre- 
viously employed. 

New glassware just delivered from the factory need not be washed 
with cleaning fluid. It should be rinsed in water and, if necessary, 
washed with "Bon Ami" which is preferable to soaps and soap powders 
because it is more readily removed. After being used for keeping eggs 
and larvae the dishes should never be washed directly in freshwater 
because of the danger of the cytolyzed cells sticking to the glass (Just, 
1928). They should be first rinsed with sea water and then washed with 
freshwater. In drying the glassware, towels coming from the laundry 
should be avoided because of the alkali present in them. If it is neces- 
sary to wipe the glassware, cheesecloth or other soft material washed 
free from chemicals should be used. Clean dishes should be stacked on 
filter paper or a clean dry cloth and protected from dust. The practice 
of placing them upside down on the laboratory table is undesirable be- 
cause of the possibility of chance contamination. Glassware used in the 
cultivation of diatoms and other algae should be sterilized in a dry oven 
{1Y2 hours at i6o°C). 

The use of celluloid. Very often the investigator is confronted with 
the necessity of making a tank or an apparatus of a special design to be 
used for a physiological or embryological experiment. The problem is 
easily solved by using celluloid which is made in sheets about 3x5 feet, 
ranging in thickness from 0.005 to °- I2 5 i ncn - The celluloid manu- 
factured by the Dupont Company and other companies producing 
cellulose by-products may appear in the market under different trade 
names (pyralin, viscoloid, etc.). Clear as well as opaque celluloids and 
that frosted on one or both sides are available. The thin sheets are 
almost as transparent as glass but the thicker ones are less clear and 
have a slightly yellowish hue. Celluloid sheets may be cut by scissors or 
sawed with a hacksaw. After the edges have been filed and sandpapered 
the pieces may be stuck together with glue made of a thin solution of 
celluloid in acetone. This solution dries very quickly and the making 
of the apparatus presents no difficulty. 

Strips and sheets of celluloid may be bent to a desired shape by 
warming them in hot water. After being cooled they become rigid again. 



1 6 Marine Invertebrates 

Celluloid is non-corrosive and apparently insoluble in seawater. It 
has proved to be the most valuable and indispensable material for the 
physiological work carried on in the author's laboratory at the U. S. 
Bureau of Fisheries station at Woods Hole, Mass. (See note i on p. 50.) 

Seawater supply. Marine biological laboratories are usually pro- 
vided with running seawater pumped from the sea and stored in tanks. 
The installation of a satisfactory salt water system is primarily an 
engineering and architectural problem the discussion of which is beyond 
the scope of the present book. It suffices to mention here that seawater 
delivered to the laboratory tables must be pure and should not con- 
tain toxic substances. Attempts are always made therefore to avoid its 
contact with heavy, easily oxidized metals such as iron and copper by 
using lead, rubber, or rubber-lined steel pipes and nickel or rubber 
pumps. Many marine laboratories use large capacity bronze pumps 
which give satisfactory results, for the seawater comes in contact with 
the metal part of a pump only for a very short time. Celluloid has been 
recommended for pipe and fittings and is very successfully used by the 
Biological Station and Aquarium at Helgoland (Hagmeier, 1925). Sea- 
water delivered to a laboratory table through rubber-lined or celluloid 
pipes seems to be deprived of toxic substances, and may safely be used 
for physiological experiments and the cultivation of the most delicate 
marine forms. 

The investigator is sometimes confronted with the necessity of con- 
ducting his research in a locality remote from a permanent biological 
station. He can, however, easily provide himself with running seawater 
by purchasing a small centrifugal pump with an electric motor, a suf- 
ficient length of ordinary garden hose and a small tank which should be 
supported by a platform. A pump delivering 5-10 gallons a minute 
and two 5o-gallon wooden barrels for a storage tank, placed 5-6 feet 
above ground will provide a sufficient amount of running water for four 
or five aquaria. One of the barrels may be equipped with a float con- 
nected to a switch controlling the operation of the motor so that the 
pump works only when the water in the barrels reaches a certain low 
level. The whole equipment may be purchased and assembled for about 
$60.00. 

It has been the author's experience in establishing small temporary 
laboratories in various localities along the Atlantic and Gulf coasts that 
ordinary cast iron pumps are preferable to bronze ones. There is but 
little oxidation of iron when the pump is in operation and consequently 
water delivered to the laboratory is not toxic. Furthermore, water sup- 
plied under similar conditions by bronze pumps proves to be much more 
harmful to a number of marine forms, such as lamellibranch larvae, that 
are very sensitive to minute amounts of copper. 



Marine Aquaria 17 

A great number of investigators located in inland laboratories are 
interested in the possibility of maintaining marine forms in a limited 
amount of seawater, shipped to them from the nearest place on the shore. 
For this purpose water must be collected from an unpolluted locality, 
preferably off shore, and stored in a suitable container. Glass bottles 
are of course ideal but not always practicable for shipping. 

If storage in glass containers is desired the ordinary carboys for spring 
or distilled water may be used. A more practicable and cheaper way is 
to collect and ship seawater in 25 or 50 gallon paraffined oak barrels. 
After a few days of storage the organisms present in the water die and 
the decomposing organic matter depletes the oxygen content of the water, 
resulting in the accumulation of hydrogen sulphide (H 2 S). This, how- 
ever, does not render the water unsuitable for laboratory use. By 
aerating the sample for several hours all the hydrogen sulphide is driven 
out or oxidized and the sulphur is precipitated and may be filtered off. 
The author's experiments (unpublished data) show that the presence of 
sulphur is not harmful to many marine invertebrates and that it pro- 
motes the growth of Nitzschia cultures. 

Seawater in which marine forms have been kept may again be made 
suitable by filtering it through a layer of gravel and sand and aerating 
it. Increase in the concentration of salts owing to evaporation may be 
compensated for by the addition of distilled water, while the deficiency of 
calcium, phosphates, nitrates, and other salts used up by the organisms 
is restored by the proper dosage of these respective substances. The 
simplest way of controlling the concentration of salts consists in marking 
the level of water in a tank and adding distilled water whenever the level 
falls because of evaporation. Should a more accurate check be desired 
the salinity of the water may be determined by titrating the sample 
with a silver nitrate solution and finding the corresponding concentra- 
tion in Knudsen's tables. A description of the standard method of 
salinity determination may be found in Murray and Hjort (1912) and 
Oxner (1920). 

Filtration. Natural seawater may contain large quantities of sus- 
pended organic and inorganic matter which must be filtered out before 
it is supplied to the laboratory table. For this purpose some of the 
marine laboratories and aquaria have a more or less elaborate system of 
filters consisting of several layers of gravel and sand through which the 
water is allowed to pass. The institutions located far from the sea and 
dependent in their operations upon a more or less limited supply of 
water, as a rule filter the water as it leaves the aquarium tanks before 
using it again. In the laboratories located on the seashore the water is 
pumped directly into tanks in which it remains for a period of time 
sufficient for the settling of a considerable portion of sediment. The 



i8 



Marine Invertebrates 




outlet through which the water is drawn from a storage tank is always 
located several inches above the bottom. A small amount of seawater 
may be made to last a long time if the investigator regularly filters and 
aerates it. A very convenient type of filter (Fig. 19) consists of a water- 
tight wooden box with two 
pipes passing through its bot- 
tom. The long pipe serves as 
an overflow while filtered water 
runs through a short pipe after 
having passed through a layer 
of charcoal (ch), sand (sd), 
fine gravel (fg) and coarse 
gravel (eg). From a reservoir 
(not shown in the figure) the 
water is delivered to the aqua- 
ria. When in operation the 
filter is placed below the aqua- 
rium from which the water is 
drawn. (See also p. 540-) 
Sometimes experimental work requires the use of seawater entirely 
devoid of any suspended matter, either inorganic or organic. It may be 
obtained by filtering through collodion membranes (ultrafiltration), 
using fine Berkefeld filters, or by passing through a thick layer of asbes- 
tos. Water obtained by the latter method may not be free from bacteria 
but contains no plankton or micro- 
plankton. A typical arrangement is 
shown in figure 20. From a labora- 
tory faucet seawater runs slowly 
into a large Buchner funnel, the 
bottom of which is covered with a 
layer of asbestos (A) about 1 inch 
thick. A small watch glass is. 
placed on its surface to provide a 
more uniform distribution of water 
over the entire area. The funnel is 
inserted into a neck of a 2 liter 



Fig. 19. — -Filter for seawater. After 
Sachs, eg, coarse gravel; ch, charcoal; 
fg, fine gravel; sd, sand. 



To suction pump 




Fig. 20. — Filtering of water through as- 
bestos. A, asbestos; T, glass tube lead- 
ing to suction pump. 



vacuum flask connected to a 5 gal- 
lon bottle. A glass tube (T) in- 
serted in a stopper of the bottle 
leads to a suction pump. If neces- 
sary, two 5 gallon bottles may be connected in a series. Filtered sea- 
water fills up the flask and gradually is sucked into the bottle. After a 
few days of operation a compact organic film forms on the surface of the 




Fig. 21 
air blower. 



The 



IT 



Marine Aquaria 19 

asbestos and nitration becomes much more efficient. The 
rate of filtration may be regulated by the vacuum produced 
in the system. 

Aeration. The problem of aeration is easily solved in 
laboratories supplied with compressed air. To avoid pos- 
sible contamination of water by oil vapors from the pump 
or by rust which often accumulates in the pipes the air is 
washed by passing it through a wash bottle filled with 
water. In most cases the pressure in the compressor is 
greater than may be used conveniently for aeration. Pres- 
sure in the wash bottle, regulated by a number of outlets 
inserted in the rubber stopper and provided with rubber 
tubing and screw clamps, may be very accurately adjusted 
and the system used as a safety valve. 

To obtain quick absorption, air blown through water 
must be delivered in the finest bubbles. 
This is accomplished by using small 
blocks made of porous stone (filtros) 
mounted by means of DeKhotinsky ce- 
ment at the end of a glass tube. (The blocks are 
usually available at stores selling aquarium supplies 
and fishes.) Should less efficient aeration be desired 
air is blown through a capillary tubing or through a 
blower (Fig. 21). 

In laboratories not equipped with compressed air, 
aeration may be provided by a small electric pump.* 

By means of the following simple devices aeration 
may be obtained in laboratories not provided with 
electricity. Two glass cylinders about 8 inches long 
and iY 2 inches wide connected by a long vertical tube 
are mounted 4 to 5 feet apart on the wall or on a suit- 
able stand (Fig. 22). As the water runs drop by drop 
through the upper cylinder into a glass tube and fills 
the lower cylinder, air is sucked through the outlet 
(A). By regulating the level of the lower outlet (L) 
a small pressure is produced in the lower cylinder and 
air is driven through the outlet (C). After being ad- 
justed the cylinders work for a long time without any 
further attention. 

Aeration of the aquarium containing running sea- 

* Of the great variety of pumps available on the market the most convenient type for 
a small laboratory is a model manufactured by Marco Air Product Company, Blooming- 
field, N. J. This small and inexpensive instrument is sufficient to aerate 6 or 7 medium- 
sized tanks. Its operation requires very little attention. 



V 



• 



VJ 



Fig. 22. — A sim- 
ple air pump. A, 
C, and L, outlets. 



20 



Marine Invertebrates 



ivm flrrH ""fn 1' 



water may be provided efficiently by hanging a loosely fitted glass tube 
over the faucet and allowing a fine jet of water to strike the surface with 

considerable force (Fig. 23). Tiny air bub- 
bles are carried down the entire length of the 
glass tubing, the lower end of which almost 
reaches the bottom, and escape through the 
water, thereby aerating the fluid. 

A small but very efficient pump designed 
by A. E. Hopkins (1934) may be con- 
structed easily from a piece of celluloid. 
The device consists of a small, motor- 
driven, centrifugal pump enclosed within a 
chamber from which it draws water (Fig. 
24). The rotor (R) is cut from a piece of 
celluloid y 8 inch thick and is mounted on the 
shaft of y 4 inch glass tubing. The pump 
receives water through the hole (X) in each 
side and pumps it out through the tube (O) . 
A larger tube (I) leads from the aquarium 
into the pump chamber to permit continuous 
replenishment of the water. The pump is 
entirely water-lubricated and the only for- 
eign materials used in its construction 
are celluloid, rubber, and glass. 

Super saturation. Not all the forms commonly kept in marine aquaria 
require aeration. As a matter of 
fact many of them may be in- 
jured or even killed by injudi- 
cious aeration. Furthermore, 
on account of considerable 
pressure in the seawater pump, 
water may be supersaturated 
with air and become decidedly 
toxic. To avoid injury to the 
organisms such water must not 
be delivered directly to the 
tanks but should be allowed to 
stand until equilibrium with 
the atmospheric gases is estab- 
lished. If the water must be 

used immediately it should be de-aerated by allowing it to fall on an 
inclined glass, porcelain, or celluloid plate from which it runs into the 
aquarium. 



Fig. 

means 



23. — Aeration by 
of a jet of water. 




Fig. 24. — The pump for circulation and aera- 
tion of water in small aquaria. After Hopkins. 
I, tube leading from aquarium to pump cham- 
ber; O, tube for excurrent water; R, rotor; X, 
hole for incurrent water. 



Marine Aquaria 



21 



Water used in the experiments on fertilization of eggs, development 
of larvae, etc., should never be taken directly from the laboratory faucet. 
A suitable container, such as a one or two liter flask, should be filled and 
set aside for several hours before using to permit the escape of excess 
of air. For metabolic studies all the micro-organisms suspended in the 
water must be carefully removed by filtration through a suitable filter 
and the oxygen content of the sample must be standardized. 

Agitation. Agitation of water is essential for maintaining certain 

forms such as jelly fishes, small crustaceans, larvae of lamellibranch 

1 1 

1 1 

c ii 

u 




Fig. 25.— The plunger jar. After E. T. Browne. A, glass disc or plate; B, glass 
rod; C, lever; D, axis of lever; E, bar to stop downward motion of bucket; F, bucket; 
H, flexible hose; K, string; S, siphon. 

mollusks, etc., which otherwise settle on the bottom, stick to each other, 
and die. In the aquaria with running seawater the inlets and outlets may 
be arranged so as to provide efficient circulation. When the water is 
not changed for several days or weeks, some method of mechanical agi- 
tation is necessary. In a small aquarium this may be accomplished by 
blowing air through glass tubing lowered to the bottom of the tank. 
Should aeration not be desired, various kinds of stirring apparatus may 
be used. There are on the market many types of stirrers, consisting of 
glass or metal propellers rotated by means of electric motors, which may 
be adapted to this purpose. The difficulties are that in most cases the 
stirring devices are designed for use during a limited period of time and 
are not suitable for long, continuous use. A very simple and practical 
device which overcomes this difficulty was designed by Browne (1907) 
and is known as a plunger jar (Fig. 25). It consists of a glass disk or 
plate (A) suspended from one end of the lever (C), rotating along axis 



22 



Marine Invertebrates 



Qj) 



Fig. 



>6. — The overflow 
siphon. 



(D). A bucket (F) with an automatic siphon (S) is suspended from 
the other end of the lever. The bucket is filled with water delivered 
through a flexible hose (H). The size of the bucket is so adjusted that 
when it is almost full it swings down and pulls the plate (A). As soon 
as the level in the bucket is slightly above the upper level of the siphon 
the water is emptied through it and the glass plate swings down pulling 
up the bucket. A bar (E) stops the downward motion of the bucket 

whereas a piece of string (K) prevents the 
striking of the glass plate (A) against the 
bottom of the tank. The tank may be kept 
under a cover with a small hole permitting the 
passage of a glass rod (B). A plunger may 
be made of celluloid as well as of glass. The 
instrument is easy to make according to de- 
sired specifications. If several jars are used, 
one master bucket may operate all the plungers 
which are connected to the lever by means of 
a series of strings and pulleys. Plunger jars are very successfully used 
in the Plymouth laboratory in maintaining and rearing very delicate 
marine animals. 

Constant level arrangements. The simplest way to maintain a con- 
stant level of water in a tank is by inserting a horizontal pipe at the de- 
sired height in the wall. The opening of the outlet must be covered with 
a screen to prevent the escape of the animals. This arrangement may, 
however, prove unsatisfactory in many cases because of the clogging of 
the screen. Better results are obtained with a vertical overflow pipe 
passing through the bottom of a tank and protected with a metal screen 
cylinder extending above the level of the water. 
The cylinder is mounted on a tightly fitted cork or 
rubber stopper with a hole for the passage of a 
tube. The size of the mesh, of course, depends 
upon the dimensions of the organisms kept in the 
tank. The metal screen used for this purpose 
should not be corrosive in seawater. According to 
Richards (1933) screens made of pure nickel or 
of stainless steel are the least toxic. 

The desired level of running water may be 
maintained by using an overflow siphon which is 
made by bending over one arm of a U-shaped glass 
and placing it in a position shown in figure 26. 
The controlling level of the siphon may be 
changed easily by attaching a rubber tubing to the short arm and fasten 
ing the free end of it at the desired height. 



3 



Fig. 27. — The overflow 
siphon. Adapted from 
Hagmeier. 



Marine Aquaria 23 

In another type of arrangement (Fig. 27) the outside arm of the 
siphon is inserted in a small glass cylinder. A glass tube passing through 
the bottom serves as an overflow controlling the level in the tank. The 
instrument may easily be made from a large test tube by cutting off its 
bottom and inserting two rubber stoppers. A hole in the upper stopper 
serves for the escape of air. 

The following arrangement (Fig. 28) permits a careful regulation 
of the rate of flow of filtered water through a series of culture vessels. 




Fig. 28. — The arrangement used in rearing marine larvae. A, reservoir tank; B, C, D, 
E, culture jars; F, funnel with cotton filter; L, overflow siphon controlling the level 
of water in the funnel; S, overflow siphon controlling the level in culture jars. 

Tall cylindrical jars with wide lips are very convenient for this purpose. 
To insure a constant flow the water is first filtered through cotton (F) and 
siphoned from the tank (A), which is always kept full by means of an 
overflow siphon (L). The first jar (B) serves only as a supply reser- 
voir for the others (C, D, E) which are connected to one another by 
siphons. The jars are placed on boards of various thicknesses so that 
in each of them the level of the overflow (the lip of the jar) is gradually 
increasing from left to right. An automatic siphon (S) controls the 
level of water in the last jar. By this arrangement water gradually passes 
from (A) through all other jars and is discharged by the siphon (S). 
Should the latter become clogged, the water in the first jar rises and 
overflows before the culture jars, standing slightly above it, become full. 



24 



Marine Invertebrates 



A few organisms may swim back through the siphon connecting the 
second and the first jar and be lost. The opening of the controlling 
siphon may be covered with plankton silk or other fine material to pre- 
vent the escape of minute organisms. This arrangement has been suc- 
cessfully used by the author in rearing and maintaining small crusta- 
ceans, the larvae of mollusks and echinoderms, and small jelly fishes. 

Further improvement of this method was made by F. G. Walton 
Smith whose personal communication reads as follows: "Rapid clog- 
ging of the pores of this material (bolting silk) is prevented by dipping 
the mouth of the siphon under the surface of molten paraffin wax melt- 
ing at 48 ° C. and then blowing air through the other end as it is removed. 
The resulting smooth coating of wax on the fibers seems to prevent 




Fig. 29. — The current rotor. 

the entanglement of larvae and allows the filter to work efficiently for 
a much longer period than would otherwise be the case. The net is 
attached to the siphon by means of a wide rubber band, and is of such 
a nature that when worked, the openings are just small enough to serve 
to retain the larvae." Using this technique Dr. Smith had no difficulty 
in growing oyster larvae at the U. S. Bureau of Fisheries Station at 
Beaufort, N. C. 

Slow exchange of water may be obtained by using the so-called "filtros" 
block which may be installed as a partition in the aquarium, or by 
inserting a Berkefeld filter and slowly sucking the water through it. 
Stone filters become clogged very quickly and require frequent changes. 

Current rotor. This instrument is designed to change the water in 
the aquarium without losing the small organisms living in it. The es- 
sential feature of the apparatus (Fig. 29) designed by Galtsoff and Cable 
( 1933) is a cylinder (A) of 60 mesh or finer nickel screen suspended in a 
tank and rotated by means of an electric motor. Rotation of the 
cylinder when placed at one end of an oblong aquarium sets up a com- 
plex system of currents the direction of which is indicated in the accom- 



Marine A quaria 2 5 

panying illustration. Strong circular currents are formed in the imme- 
diate vicinity of the cylinder, while at the far end the water moves 
very gently. There is also a noticeable upward motion from the bottom 
of the tank. 

The speed of rotation and the corresponding strength of currents may 
be regulated by the speed of the motor, controlled through a rheostat, 
and by means of a set of pulleys of different diameters. The dimensions 
of the cylinder also affect the strength of the current produced and they 
therefore should vary according to the size of the aquarium used. The 
cylinder shown here is 4 inches in diameter and 6 inches long. About 
one inch is left above water. The tank is 25 x 15 x 14 inches. The 
bottom of the cylinder (B) 
is a celluloid disk. Non- 
corrosive material should 
also be used for the suspen- 
sion rod (C) and brace wires 
(W). The diameter of the 
pulley (P) is 12 inches. 

The water may be with- 
drawn from the tank through 
a siphon (S) the upper end 
of which is placed inside the 
revolving cylinder. When 
the cylinder is in rotation 
small organisms are never 
actually drawn against its 

wall, because the centrifugal force throws them away from it. They 
are then caught up in the circular currents and soon find themselves in 
quieter waters at the far end of the aquarium. In this manner the 
water in the tank may be changed without losing its inhabitants. 

When desirable, a constant flow of water may be supplied by placing 
the lower end of the overflow siphon (S) in a vessel (V) , adjusted so that 
the top of it is level with the water in the aquarium. The water is 
introduced from a reservoir, in which it is kept at a constant level. If 
necessary the water from a laboratory faucet may be filtered through 
glass wool to remove sediment and other foreign matter. 

Imitation of tidal movement. The arrangement imitating tidal move- 
ment of water and permitting attached forms to be subject to rhythmical 
changes in hydrostatic pressure consists of a series of tall jars in which 
the equilibrium is maintained through a system of tubes (T, Ti) (Fig. 
30). An automatic siphon (S) in jar (A) controls the level in all the 
jars. After the level in the controlling jar has reached the highest posi- 
tion the water begins to empty through the siphon drawing it from all 




Fig. 30. — The arrangement imitating tidal 
changes. A, controlling jar; S, automatic siphon; 
T, T lf glass tubes connecting the jars. 



26 



Marine Invertebrates 



H, f= 



G 



the other jars. As soon as the lowest level is reached (determined by the 
position of the short arm of the siphon) the system begins to fill up 
and the level in all the jars rises. The rate of flow of water may be 
regulated to obtain the desired tidal interval. This arrangement was 
first employed by H. F. Prytherch in a study of the effect of oil pollution 
on oysters (Galtsoff, Prytherch, et al., 1935). (See note 2 on p. 50.) 

Circulation of water in a closed system. Many organisms may be 
maintained in a limited supply of seawater if the latter is kept in circu- 
lation and is systematically filtered. Various simple devices designed 

to meet these requirements are based on 
the use of an air pump. The following 
is the description of a simple but efficient 
device (Fig. 31) designed by Burch and 
Eakin which we copy from Science 
(1934). The pump (P) (Fig. 31) is 
made from a pyrex glass test-tube, 10 cm. 
high and 1.5 cm. in diameter. A glass 
tube (A) 5 mm. in diameter is sealed to 
the side of the test-tube approximately 2 
cm. from the mouth and then bent 
parallel with the test-tube. A similar 
glass (B) is sealed to the base of the 
test-tube. The pump is placed in an 
inverted position in the reservoir and an 
exceedingly small air current is permitted 
to enter the pump through the glass tube 
(A) at the side. The exact depth at 
which the pump will give a maximum 
efficiency may be determined by experi- 
mentation; however, the pump should be 
at least 15 cm. below the water level in the reservoir. (See note on p. 50.) 
Another method developed by Cleve (quoted from Hagmeier, 1933) is 
shown in figure 32. The bottom of a tank is covered with a thick layer 
of sand (sd) through which water is sucked into a funnel (F) forming 
the lower end of the siphon, and is emptied into a small tank filled with 
sand and charcoal (ch). From a filter tank by a similar arrangement of 
siphon and funnel (Fi) the water enters into a U-shaped glass tube (U). 
Bubbles of air, blown in at point L gradually push the water into the 
horizontal tube (T) and back into the tank. By using this arrangement 
various marine forms may be kept for a very long time in a small amount 
of seawater. 

A similar arrangement for aeration and circulation of water was used 
by Browne (1907) for the cultivation of hydroids (Fig. ^t,). Air blown 




Fig. 31. — A device for water 
circulation. After Burch and 
Eakin. P, pump; A, B, glass 
tubes. 



Artificial Seawater 



27 



very gently through a tube (A) causes the circulation of water in a 
small tube (B) in which a portion of a hydroid colony is suspended. 

ARTIFICIAL SEAWATER 

From a physiological point of view seawater is a well balanced solu- 
tion of mineral salts, virtually all of them existing in the ionic form. 
Earlier analyses accounted only for the major salts, ignoring other sub- 
stances present in extremely small quantities. On the basis of the 



■*w 



fr^ 



V 



h 



ra 



ch 



m 



^-= 



- v - •. 



>V r y ; -. ■.■:■•-,,-. ■ ■■>-, v- 




Fig. 32. — The arrangement for filtration 
and circulation of water. After Cleve, 
from Hagmeier. F, Fj, funnels forming 
the upper arms of the siphons; sd, sand; 
ch, charcoal; L, place air is blown in; 
U, T, return tube. 



Fig. 33. — A device for aeration 
and circulation of water in small 
aquaria. After Hagmeier. A, 
B, tubes. 



analyses of the samples collected by the "Challenger," Dittmar gives 
the following composition of the seawater (quoted from Murray and 
Hjort, 1912) : 

table 1. Composition of seawater (according to Dittmar). 

Grams in 1000 gm. of seawater 

NaCl 27.213 

MgCl 2 3807 

MgSC-4 I0 58 

CaS0 4 l - 200 

K2SO4 °- 86 3 

CaCG-3 OI2 3 

MgBr 2 0076 

So far as the principal constituents are concerned the composition 



28 Marine Invertebrates 

of seawater salts remains more or less uniform throughout the entire 
expanse of the ocean, varying only in concentration. This may be seen 
from Table 2 (quoted from Quinton, 1912) representing the results of the 
analyses of Forchhammer (1865), Makin (1878), and Dittmar (1884), 
and expressed as percentage of total solids. 

table 2. Composition of seawater (percentage of total solids). 

Forchhammer Makin Dittmar 

NaCl 78-32 76-9IS 77-758 

MgCb 9-44 11 -4°7 10.878 

MgS0 4 6.40 4.483 4-323 

CaS04 3.94 4-226 4.070 

K2SO4 2.468 2.465 

KC1 1.69 

MgC0 3 0.290 

MgBr2 0.298 0.217 

Other 0.21 0.206 

Analytical and physiological work carried out during the last two 
decades has greatly increased our knowledge of the chemistry of sea- 
water, and has shown that certain substances, such as phosphates, 
nitrates, silica, and iron, which occur in infinitesimally small amounts, 
are indispensable to the growth and propagation of marine forms. A 
complex interrelationship between the concentration of the nutrient 
salts and the productivity of the sea is at present the principal problem 
of oceanic biology. Progress in this field of research became possible 
only after the chemists developed and perfected simple colorometric 
methods of determination of phosphates, nitrates, iron, and silica.* 

According to Thompson and Robinson (1932) the following elements 
or their compounds (written in the order of their abundance) are deter- 
minable in seawater. 

table 3. Approximate composition of seawater with a 
chlorinity of 19.00%. 

Concentration as millimols Concentration as millimols 

or milligram atoms per or milligram atoms per 

Constituent kg. of seawater Constituent kg. of seawater 

Chlorine 535-0 Bromine 0.81 

Sodium 454-0 Strontium 0.15 

Sulphate 82.88 Aluminum 0.07 ? 

Magnesium 52 .29 Fluorine 0.043 

Calcium 10.19 Silicon 0.04 

Potassium 9-6 Boron 0.037 

Carbon Dioxide 2.25 Lithium 0.015 

*The description of these methods would be out of place in the present book and the 
readers interested in these are referred to the book of Harvey (1928) and original papers 
published in the Journal of the Marine Biological Association of Plymouth and other 
oceanographical periodicals. 



Artificial Seaivater 29 

table 3. — {continued) 

Nitrate 0.014 Silver 0.0002 

Iron 0.0036 Nitrite 0.0001 

Manganese 0.003 ? Arsenic 0.00004 

Phosphorus 0.002 Zinc 0.00003 

Copper 0.002 Hydrogen Ion 0.00001 

Barium 0.0015 Gold 0.00000025 

Iodine 0.00035 

There are, however, a number of elements, such as cobalt, vanadium, 
lead, nickel, tin, caesium, and rubidium, the presence of which has not 
yet been detected in seawater but is postulated because they are found 
in the tissues of marine animals and plants or in the salt deposits left by 
the evaporation of large quantities of seawater.* 

On account of the exceeding complexity of seawater it was only after 
a great deal of effort that biologists succeeded in elaborating a formula 
for an artificial solution which possesses the same properties as the 
ocean water. There exist at present several formulae for the preparation 
of artificial seawater which may be used in experimentation with marine 
animals. 

For embryological studies on echinoderms Herbst (1903-4) employed 
the following solution: 



NaCl 




3.00 gm. 


KC1 




0.08 gm. 


MgSG-4 




0.66 gm. 


CaCl 2 




0.13 gm. 


Dist. w 


ater 


100 cc. 



To this solution 1 cc. of 4.948% NaHC0 3 must be added. 

In Van't Hoff's formula the principal salts are given in the molecular 
proportions in which they occur in the sea: namely, 100 NaCl; 7.8 
MgCl 2 ; 2.2 KC1; 3.8 MgS0 4 ; and from 1.5 to 2.2 CaCl 2 . After the 
M/i solutions of the salts have been mixed in the above mentioned pro- 
portions the solution is diluted to the same concentration as that of the 
seawater in which the animals normally lived. A trace of sodium 
bicarbonate should be added to bring the pH to 8.0-8.2. Water re- 
distilled from glass must be used. While such a solution would serve 
for experimental purposes it is not suitable for maintaining the animals 
over a long period of time. Lacking in phosphates and nitrates, it is 
obviously unsuitable for the marine algae. A more complex solution 
prepared according to McClendon's formula answers this purpose. 

*For further discussion regarding the composition of the seawater and its significance 
to the life in the sea the reader is referred to the papers of Quinton (1912), Yernadsky 
(1923), Thompson and Robinson (1932), and Galtsoff (1932, 1934)- 



30 Marine Invertebrates 

table 4. Artificial seawater (according to McClendon) . 

(All solutions are of M/i concentrations unless otherwise stated.) 

NaCl 483.65 cc. NaoSiOa 0.0025 gm. 

KC1 10.23 cc. NaoSuOo 0.003 cc. 

MgS0 4 28.55 cc. H 3 P0 4 0.002 cc. 

MgClo 25.16 cc. H3BO3 100 cc. 

CaClo 1 1 .00 cc. AloCl,; 0.01 cc. 

NaBr 0.8 cc. NH 3 0.001 cc. 

NaHC0 3 2.5 cc. LiN0 3 0.002 cc. 

H 2 373-63 ". 

It is claimed that even most delicate marine algae will live in this solu- 
tion for long periods. 

For marine forms Penn (1934) recommends the use of the following 
medium which is based for anions on the analysis of the salt content of 
the blood of certain organisms and for cations on the buffering properties 
of salts. 

table 5. Perm's medium for marine forms. 

NaCl 0.1335 N 

CaClo 0.0112 N 

KC1 0.0084 N 

NaN0 3 °-°°55 N 

NaHC0 3 °°48 N 

MgS0 4 .' 0.0040 N 

KH2PO4 000 °5 N 

NaSi0 3 trace 

NH4NO3 (For green forms only) 0.0125 N 

FeCl 3 (For green forms only) trace 

Artificial seawater is regularly used by several marine aquaria in 
Europe. For example, water in the Berlin aquarium consists primarily 
of an artificially prepared solution mixed with a small amount of natural 
seawater. For making up the large quantities of artificial seawater 
needed for the operation of a public aquarium, a simple procedure must 
be followed and no attempts made to add all the salts entering into 
the composition of natural water or to use chemically pure ingredients. 

table 6. Von Flack's formula for making large quantities of 

artificial seawater. 

Sodium chloride (NaCl) 2815 gm 

Calcium sulphate (CaS0 4 2H2O) 172 gm 

Magnesium sulphate (MgS0 4 7H2O) 320 gm 

Magnesium chloride (MgCb 6H2O) 850 gm 

Potassium chloride (KC1) 80 gm 

Magnesium bromide (MgBr2) 10 gm 

Water 100 liters 



Securing Food 31 

According to Sachs (1928), elements occurring in small amounts in 
the sea may be present as impurities in the salts even in excess of their 
concentration in natural seawater. The two formulae generally used in 
Germany are given in Tables 6 and 7. 

Sodium chloride and calcium sulphate are dissolved first in about 
30 liters of water. The solution is vigorously shaken and stirred until 
all the calcium goes into solution, then the other salts are added and 
the volume is made up to 100 liters. If the water is very hard a smaller 
amount of calcium sulphate may be used. 

table 7. Schmalz's formula for making larger quantities of 

artificial seawater. 

Sodium chloride (NaCl) 2815 gm. 

Potassium chloride (KC1) 67 gm. 

Magnesium chloride (MgClo 6 HoO) 551 gm. 

Magnesium sulphate (MgSO-i 7 HoO) 692 gm. 

Calcium chloride (CaClo HoO) 145 gm. 

Water 1 00 liters 

First dissolve all the salts excepting calcium chloride; bring up the 
solution almost to 100 liters; then add calcium chloride and water to 
make up the volume. 

Before using, the newly prepared solution should be tested on sea 
anemones or other organisms. 

METHODS OF SECURING FOOD FOR MARINE 
INVERTEBRATES 

No culture of any organism may be maintained for a long period 
if proper food is not regularly supplied. In the case of carnivorous 
animals the problem resolves itself into ascertaining the forms which 
constitute the principal food of the animal in question and in finding the 
means of keeping a good supply. Thus many organisms subsisting on 
animal plankton may be maintained for a long time if an arrangement 
is possible by which regular plankton samples may be taken and the 
desired forms obtained. Copepods or other planktonic crustaceans may 
easily be segregated from the mass of algae and other micro-organisms 
with which they are closely associated, by pouring the sample of plank- 
ton into a crystallizing dish about 10 inches in diameter and from 3 to 
4 inches high, the outside wall of which is painted black with the excep- 
tion of one vertical strip about one inch wide which should remain un- 
covered. A short time after the dish has been placed on a laboratory 
table with the open space toward the light, copepods and other crusta- 
ceans congregate at the two opposite sides depending upon the photo- 
tropic reactions which control their behavior. They may easily be re- 
moved with a pipette and fed to the animals in the tanks. 



32 Marine Invertebrates 

Organisms living on debris, or those which may be fed small pieces of 
fish or shellfish, may be kept in the laboratory tanks for long periods. 
Greater difficulty is encountered in keeping alive the plankton-feeding 
forms. Seawater in laboratory circulation contains but a small number 
of microscopic algae and may therefore be lacking in essential food ele- 
ments. On the other hand, it still may contain copepods and other 
crustaceans which may directly or indirectly be destructive to delicate 
larvae or other organisms under cultivation. It is therefore advisable to 
use filtered seawater and to provide food by adding diatoms or other 
algae. This method requires a constant supply of these forms which 
must be grown in the laboratory. 

Caswell Grave (1902) originated the method of rearing marine larvae 
by putting them in a balanced aquarium in which the diatoms growing 
on the bottom furnished an abundant supply of natural food and kept 
the water pure. This method consists of putting a liter or more of sand 
dredged from the ocean bottom in an aquarium of seawater and allowing 
it to stand several days before a window, but protected from direct sun- 
light. Under these conditions a film of diatoms develops in several 
days. The larvae, from 12 to 24 hours after fertilization, are placed in 
an aquarium of fresh water to which a dozen or more pipettefuls of the 
diatom-stocked surface sand are added. The aquarium is then covered 
and set before a window. Using this method Grave succeeded in rearing 
a number of spatangoids and sand-dollars until they had completed their 
metamorphosis, and in keeping them in a healthy and growing condition 
for three months thereafter. [See p. 557.] The capacity of the aqua- 
rium was 1 liter and the water was changed only twice during this period. 

An abundant supply of various diatoms may be raised by using the 
following method developed by Just (1928). Mud and scrapings from 
eel grass are placed together with animals and plants in jars containing 
an equal amount of seawater. The jars are covered and set aside in a 
subdued light. After a period of putrefaction the culture purifies itself 
and an abundant growth of diatoms ensues. From this stock culture 
the diatoms are removed, suspended in filtered seawater and strained 
through bolting silk. Only the diatoms that have passed through the 
silk are used for feeding. It is advisable to start several cultures at 
from 5 to 10 day intervals. 

In spite of the fact that a number of marine larvae were successfully 
reared on diatoms growing in mud or sand cultures both methods suffer 
from a certain degree of uncertainty. It is impossible to predict what 
species of diatom will develop and whether similar cultures will always 
be available. For a more critical work on the physiology of feeding 
and food requirements, pure cultures should be used. At present cultures 
of a single species of diatom may be carried on indefinitely under con- 



Securing Food 33 

trolled laboratory conditions. The method originated by Miquel (1897) 
and modified by Allen and Nelson ( 19 10, 19 14) consists of adding certain 
nutrient salts to seawater, sterilizing, and inoculating with a single 
species of diatom. Two solutions are prepared separately: 

table 8. Preparation of Miquel solution. 

Solution A. 

Potassium nitrate 20.2 gm. 

Distilled water 100 cc. 

Solution B. 

Calcium chloride (CaCb 6H2O) 4 gm- 

Sodium phosphate, secondary, 

Crystals (Na 2 HP0 4 12H2O) 4 gm. 

Ferric chloride (melted) (FeCl 3 6H 2 0) 2 cc. 

Hydrochloric acid (HO) concentrated 2 cc. 

Distilled water 80 cc. 

Dissolve calcium chloride in 40 cc. of distilled water and add the 
hydrochloric acid. In a separate beaker dissolve the sodium phosphate 
in 40 cc. of distilled water, add the melted ferric chloride, and slowly mix 
the two solutions. To prepare Miquel's solution add 2 cc. of solution A 
and 1 cc. of solution B to one liter of seawater. Sterilize by bringing 
just to the boiling point. Cool and decant or filter off the slight pre- 
cipitate, separating the amount obtained into two 1 liter flasks. Shake 
vigorously to aerate. The prepared medium is poured into sterile, short- 
necked, wide-mouthed flasks of 125 cc. capacity and is covered with 
inverted beakers. The flasks should be only about % full so tnat the 
proportion of air surface to the volume of the liquid is large. The flasks 
are inoculated by adding 6 to 8 cc. of an old culture of diatoms and are 
placed in front of a window but are protected from the direct sunlight. 
They should be shaken at least once a day. 

The Miquel's seawater may be modified by adding garden soil extract 
which is known to have a stimulating effect on the growth of diatoms 
(Gran, 1931, 1932, 1933). To prepare the extract put 500 grams of 
garden soil in a flask, add 500 cc. of water and autoclave for 20 minutes 
at 15 lbs. pressure. Filter, sterilize again, and keep in the refrigerator. 
Add 1 cc. of soil extract to each liter of prepared Miquel's seawater. 
Experiments carried out by Gran (1932) at Woods Hole show that the 
synthetic Ferri-ligno-protein compound of Waksman (1932) which in its 
chemical characteristics corresponds closely to the "humic" substances 
of the soil, gives the same stimulating effect on the growth of the plank- 
ton diatoms as the soil extract. 

Needless to say, in dealing with diatom cultures the same precautions 
must be taken as are usually observed in bacteriological work. The 
cultures grow best at about 15 to 16 C. 



34 



Marine Invertebrates 



By using Miquel's seawater Allen and Nelson (1910) were able to 
obtain persistent cultures of the following species of diatoms: Asterio- 
nella japonica, Biddulphia mobiliensis, B. regia, Chaetoceras densum, 
C. decipiens, C. constrictum, Cocconeis scutellum, Coscinodiscus excen- 
tricus, C. granii, DityUum brightwellii, Lauderia borealis, Nitzschia 

— -. closterium, N. closte- 



rium forma minutissima, 
N . seriata, Rhizosolenia 
stoltcrjothii, Skeleto- 
nema costatum, Strep- 








totheca thamensis, and 
Thalassiosira decipiens. 
Of all these species 
the cultivation of Nitz- 
schia closterium /. minu- 
tissima is more widely 
practised in many lab- 
oratories both in the 
United States and in 
Europe than that of any 
other diatom. This is 
primarily due to its 
small size and its ability 
to remain in suspension 
for a long period of 
time, these qualities 
rendering it very useful 
as food for small plank- 
tonic organisms. 

Light and temperature 
are the two principal 
physical factors which 
govern the rate of prop- 
agation of diatoms. The 
growth of cultures left in the laboratory and subject to rather wide fluc- 
tuations in temperature and intensity of illumination are greatly affected 
by these changes. Temperatures above 22 ° C. are obviously harmful to 
a Nitzschia culture and when the summer heat approaches 30 , as hap- 
pens regularly in the author's laboratory of the U. S. Bureau of Fisheries, 
they may perish. To avoid this difficulty and to keep the cultures in 
health, both temperature and illumination should be kept constant. 

The following easily constructed arrangement has been successfully 
used by the author. Culture flasks are kept in a cabinet (Fig. 34), 60" 



Fig. 34. — The cabinet for diatom cultures. A, air 
switch; C, dry cells; E, relay; F, fan; H, small 
heater; M, metastatic temperature controller; P. 
pipes to refrigeration plant; R, refrigeration unit; 
S, wall switch; T, transformer. 



Securing Food 



35 





x 32" x 11", with a door (not shown in figure) and back made of double 
glass panes with a 1 inch air space between them. The cabinet is con- 
structed of wood and insulated with celotex; the inside is painted white. 
Culture flasks are kept on wire shelves which rest on adjustable metal 
supports. A clearance of not less than % inch be- 
tween the edges of the shelves and the walls of the 
cabinet provides for better circulation of air. The 
upper shelf is occupied by the refrigeration unit (R) 
separated from the rest of the cabinet by a metal sheet 
which serves for collecting the condensation water, and 
also as a protection to the cultures placed just below it 
against excessive cold. The temperature is regulated 
by the air switch (A) connected to the electric re- 
frigeration machine (not shown in the diagram). To 
facilitate circulation of air a small electric fan (F) 
may be hooked up to the air switch. Experience 
shows, however, that when the refrigeration unit is in 
operation there is a sufficient circulation of air inside 
stand for iiiumina- the box. The temperature differences in the flasks 
tion of the diatom placed on various shelves do not exceed 2 C. and a 
temperature of from 15 to 16 C. may easily be main- 
tained in the cabinet when the room temperature does not exceed 22 ° C. 
If necessary the refrigeration unit may be 
disconnected and a heater, placed on the 
bottom shelf, turned on. The heating 
assembly consists of a small heater (H), 
fan (F), relay (E), and metastatic tem- 
perature controller (M). 

Equal illumination on both sides of the 
cabinet is provided by 18 Mazda 25 watt 
bulbs, mounted on two separate stands ~^ 
(Fig. 35) made of iron pipes and pro- 
vided with metal reflectors which are 
placed 3 feet from the glass walls. Under 
this constant, controlled illumination and 
temperature, Nitzschia cultures grow bet- 
ter and faster than when kept before the 
laboratory window. 

In a study of the effect of intensity of 
illumination on the growth of Biddulphta 
mobiliensis and Carteria sp., Schreiber 
(1927) used the following method (Fig. 
36). Cultures kept in the box were 





Q 



n 



I T 



3^, 






Fig. 36. — The method of grow- 
ing Nitzschia under artificial il- 
lumination. After Schreiber. 
O, screen made of oiled paper. 



36 



Marine Invertebrates 



illuminated from above. To avoid over-heating, light from a iooo watt 
gas-filled bulb was passed through a layer of water. A screen (O) made 
of oiled paper was inserted to provide uniform illumination. 



METHODS OF OBTAINING A CULTURE OF A 
SINGLE SPECIES 

Two methods have been employed by Miquel in obtaining cultures of 
a single species of diatom. The method of isolation consists in 

picking out an individual cell under the micro- 
scope and introducing it into a prepared 
medium. If the method of subdivision is 
used, a small quantity of water containing a 
mixture of various organisms is added to a 
prepared medium and poured out into a num- 
ber of tubes. If the operation is repeated 
many times some of the tubes may contain 
one unit of diatoms only from which a fresh 
culture may be made. 

Allen and Nelson (ioio) proposed the fol- 
lowing modifications of this procedure. One 
or two drops of plankton are added to 250 cc. 
of a suitable sterile medium and poured into 
petri dishes. The dishes should be kept under 
a constant temperature in a subdued light, in 
a place where they may be examined with a 
hand lens without moving or disturbing them. 
In the course of a few days colonies of differ- 
ent species of diatoms will be seen growing at 
different spots on the bottom. They may 
be picked up and transferred to fresh culture media. [See pp. 43, 70.] 
The isolation should be made as early as possible before all the water 
is infected by some one organism, either diatom or flagellate. Some- 
times it is necessary to repeat the process several times before one suc- 
ceeds in isolating the desired species. To facilitate the process of elimina- 
tion of flagellates and other micro-organisms Schreiber (1927) recom- 
mends the use of a device shown in figure 37. The principal part of it 
consists of a U-shaped glass tube, the middle piece of which is drawn 
into a capillary ; one arm is made longer than the other. A diatom culture 
is fed drop by drop into the long arm from which it flows through the 
capillary into the short arm. Owing to the higher velocity of the current 
in the capillary, the diatoms are carried into the left arm and accumulate 
in the lower part of it. The rate of the flow of water through the appara- 




Fig. 37. — The device used 
for the concentration of 
diatoms. After Schreiber. 
D, diatoms. 



Culture of Single Species 37 

tus should be adjusted so that the velocity of the vertical current in the 
left arm is less than the rate of sinking of the diatoms. Free swimming 
flagellates and organisms lighter than diatoms escape while the latter 
accumulate in the area (D) just above the mouth of the capillary tubing. 

Cultures of a single species of diatom obtained by the method of isola- 
tion and subsequent washing usually are contaminated with bacteria, 
the presence of which apparently does not interfere with the growth 
of the diatom. Allen designates them as "persistent" cultures, reserving 
the name "pure" only for bacteria-free cultures of a single diatom species. 
The elimination of bacteria is a very difficult and time consuming process 
which consists of repeated washings in sterile media followed by frac- 
tional subdivision. Purification of cultures by a method of differential 
poisoning was attempted by Allen ( 1914) with only a measure of success. 
Cultures of Thalassiosira gravida were treated by adding 1 mg. of 
CUSO4.5H2O to each 100 cc. After an interval of 12 minutes a fresh 
medium was inoculated with 1 cc. of the first one. In this way the num- 
ber of bacteria was materially reduced but complete sterilization was 
not obtained. 

Chlorination produced by electrolysis of water was applied also only 
with partial success (Allen, 1914). An electric current varying from 
1.7 to 1.5 amperes was passed between the two sterile carbon electrodes 
immersed in seawater. The electrolysis was continued for about 3 min- 
utes; then the water was allowed to stand for one hour. Fifty cc. of 
chlorinated water were added to an equal amount of sterile medium and 
the solution was inoculated with a small amount of a Thalassiosira cul- 
ture. In this way the number of bacteria was materially reduced. 

In healthy cultures the presence of bacteria does not interfere with 
the propagation of the diatom (Nitzschia) but as soon as conditions 
become unfavorable the bacterial growth is promoted and the diatoms 
suffer (Galtsoff, et al., 1935). The unhealthy state of such cultures is 
easily noticeable for the cells stick together forming large clumps which 
settle on the bottom and which sink almost immediately after stirring. 
Microscopical examination of a stained preparation showed that every 
Nitzschia cell was covered with a large number of bacteria closely ad- 
hering to its body. 

In old cultures Nitzschia has a tendency to develop teratological forms. 
This condition may be remedied by subculturing and it usually disap- 
pears in a short time. 

Natural seawater enriched by the addition of nutrient salts appears to 
be the best medium for the cultivation of diatoms. It is of interest that 
according to Allen (1914) Thalassiosira failed to grow in artificial sea- 
water to which nitrates, phosphates, and iron were added according to 
Miquel's method. Excellent results were obtained, however, when less 



38 Marine Invertebrates 

than 1% of natural seawater was added to the culture medium. Arti- 
ficial seawater used in Allen's experiment comprised only the six principal 
salts (see Dittmar's analysis of seawater, page 27) and apparently was 
deficient in some other growth-promoting substance which is present in 
natural seawater. 

Other solutions than that of Miquel have been recommended by 
several investigators for the cultivation of diatoms and green forms. 
Schreiber (1927) gives the following formula: 

table 9. Medium for cultivation of diatoms and green forms. 

. (Schreiber) 

Potassium nitrate (KNO3) 0.2 gm. 

Dipotassium hydrogen phosphate (K2HPO4) .... 0.1 gm. 

Potassium silicate (K^SiOa) 0.01 gm. 

Ferric sulphate (Fe 2 [S04].3) °-°°5 gm- 

Redistilled water 50.00 cc. 

This solution is added to 950 cc. of filtered seawater and the medium is 
sterilized by steam at about ioo° C. If during the heating a precipitate 
(calcium carbonate) is formed, the solution should be allowed to stand 
several weeks until a sufficient amount of C0 2 has been absorbed from 
the air and the calcium salts redissolved. 

Although diatoms constitute the most essential food element of a great 
number of plankton-feeding larvae, other food organisms should not be 
neglected. Mixed cultures of various green flagellates may be obtained 
from jars in which the algae are allowed to putrify. These cultures once 
obtained may be carried on for a long time by inoculating Miquel's or 
other media. Their usefulness in feeding marine larvae should be 
ascertained, however, by experimentation. 

Isolation of a green flagellate and its maintenance in a bacteria-free 
culture presents less difficulty than does the isolation of a diatom. Using 
standard bacteriological technique, German investigators succeeded, for 
instance, in obtaining bacteria-free pure cultures of Carteria sp., a phyto- 
monadine flagellate very common in the North Sea (Schreiber, 1927). 
The sample of plankton is first centrifuged in sterile seawater and set 
aside and the Carteriae, which are positively phototropic, are separated 
from the rest of the planktonic organisms and placed on gelatin or agar 
plates, from which they are subcultured. 

Sperm of marine algae is often used as food for small lamellibranch 
larvae which are unable to ingest diatoms. At Woods Hole the sperm 
of Ulva may be obtained during the summer. Freshly collected plants 
are left exposed overnight on pieces of filter paper. The next morning the 
leaves are put in a shallow crystallizing dish filled with water and set 
in a place exposed to strong light. In a short time large masses of green 
sperm may be pipetted from the side having the greatest illumination. 



Culture of Single Species 39 

In order to insure an ample supply of food, the cultivation of diatoms 
or other algae should be started at least two weeks before the time of the 
experiment to be conducted with marine larvae. Between the seasons of 
experimental work with marine larvae, a small number of stock cultures 
of their food should be maintained in the laboratory. 

Bibliography 
Allen, E. J. 1914. On the culture of the plankton diatom Thalassiosira gravida, 

Cleve, in artificial seawater. J. Mar. Biol. Assoc. 10:417. 
Allen, E. J., and Nelson, E. W. 1910. On the artificial culture of marine plankton 

organisms. Ibid. N. S. 8:421. 
Bond, R. M. 1933. A contribution to the study of the natural food-cycle in aquatic 

environments. Bull. Bingham Oceanog. Coll. 43, Art. 4:1. 
Browne, E. T. 1898. On keeping Medusae alive in aquarium. J. Mar. Biol. Assoc. 

5:186. 

1907. A new method of growing Hydroids in small aquaria by means of a 

continuous current tube. Ibid. N. S. 8:37. 

Burch, A. B., and Eakin, R. M. 1934. A device for water circulation. Science 

80:563. 
Dittmar, W. 1884. "Challenger" reports. Physics and Chemistry 1. 
Forchhammer, G. 1865. On the composition of seawater in the different parts of 

the ocean. Philos. Trans. 155. 
Galtsoff, P. S. 1932. The life in the ocean from a biochemical point of view. J. 

Wash. Acad. Set. 22:246. 

1934- The biochemistry of the invertebrates of the sea. Ecol. Monog. 4:481. 

Galtsoff, P. S., and Cable, L. 1933. The current rotor. Science 77:242. 
Galtsoff, P. S., Prythercii, H. F., Smith, R. O., and Koehring, V. 1935. The 

effects of crude oil pollution on oysters in Louisiana waters. Bull. U. S. Bur. Fish. 

48:143. 
Gran, H. H. 1931. On the conditions for the production of plankton in the sea. 

Cons. Perm. Intern. Pour V Exploration de la Mer. Rapp. et Proces-Verbaux des 

Reunions. 75:37. 

1932. Phytoplankton methods and problems. J. du Cons. Intern. Pour 

V Exploration de la Mer. 7:343. 

1933. Studies on the biology and chemistry of the Gulf of Maine. Biol. Bull. 

64:159. 
Grave, C. 1902. A method of rearing marine larvae. Science 15:579. 
Hagmeler, A. 1925. Neue Aquarium-einrichtungen der Staatlichen Bioligischen 

Anstalt Auf Helgoland. Intern. Revue d-gesamt Hydrobiol. u. Hydrographic 

12:405. 
1933. Die Ziichtung verschriedener Wirbelloser Meerestiere in Abderhalden's 

Handbuch d. Biol. Arbeitsmethoden Abt. 9 T. 5, 1:465. 
Harvey, H. W. 1928. Biological Chemistry and Physics of the seawater. Cambridge 

University Press. 194 pp. 
Herbst, C. 1903-4. Ueber die zur Entwicklung der Seeigellarven Notwendigen 

anorganischen Stoffe, ihre Rolle und ihre Vertretbarkeit. Arch. f. Mech. 17:306. 
Hopkins, A. E. 1934. A mechanism for the continuous circulation and aeration of 

water in small aquaria. Science 80:383. 
Johnstone, J. 1908. Conditions of life in the sea. Cambridge University Press. 

332 PP. 
Just, E. E. 1928. Methods of experimental embryology with specific reference to 
marine invertebrates. The Collecting Net. 3:1. 



40 Marine Invertebrates 

Makin, C. J. S. 1878. On the composition of the Atlantic Ocean. Chem. News 

77:155-156, 171-172. 
McClendon, J. F. 1917. Physical Chemistry of Vital Phenomena. 
Miquel, P. 1890-1893. De la culture artificielle des Diatomees. Le Diatomiste 

1:73; 93:121; 149:165. 
Murray, J., and Hjort, J. 1912. The depths of the ocean. Macmillan Co. 821 pp. 
Oxner, M. 1920. Manuel Pratique de l'analyse de l'eau de Mer. I. Chlorination 

par la methode de Knudsen. Bull. d. 1. Comm. Intern, p. I'explor. Scientifique de 

la Mer Mediterranee. 36 pp. 
Penn, A. B. K. 1934. Physiological media for fresh water and marine protozoa. 

Science 80:316. 
Quinton, Rene. 191 2. L'Eau de Mer Milieu organique. Paris, Masson et Cie. 

503 PP- 
Richards, O. W. 1933. Toxicity of some metals and Berkefeld filtered seawater 

to Mytilus edulis. Biol. Bull. 65:371. 
Rogers, C. G. 1927. Textbook of Comparative Physiology. McGravv Hill Co. 

635 PP- , . 

Sachs, W. B. 1928. Meerwasser Aquarium in T. Peterfi, Methoden der Wissenschaft- 

lichen Biologie, 2:232. 
Schhler, J. 1933. Ueber kultur und Methoden beim Studium der Meerespflangen 

in Abderhalden's Handbuch d. Biol. Arbeitsmethoden. Abt. 9 T. 5, 1:181. 
Schreiber, E. 1927. Die Reinkultur von marinem Phytoplankton und deren 

Bedeutung fii die Erforschung der Produktions fahigkeit des Meereswassers. 

Wiss. Meeresuntersuchungen Abt. Helgoland, 16, 10:1. 
Schubert, A. 1930. Entwicklung des Nannoplanktons in Rohkulturen mit she- 

benden Seewasser. Wiss. Meeresuntersuchungen, Abt. Helgoland, 18, 2:1. 
Thompson, T. G., and Robinson, R. I. 1932. Chemistry of the Sea. Bull. Nat. 

Res. Council 85, Oceanography, p. 95. 
Vernadsky, W. J. 1923. La Composition chimique de la Matiere vivante et la 

chimie de l'ecorce terrestre. Revue General des Sciences Pures et Appliques, 34:42. 

1924. La matiere vivante et la chimie de la Mer. Ibid. 35:5. 

1924. La Geochimie, 404 pp. Librairie Felix Alcan, Paris. 

Waksman, S. S., and Iyer, K. R. N. 1932. Synthesis of a humus-nucleus, an im- 
portant constituent in soils, peats and composts. J. Wash. Acad. Sci. 22:41. 

COLLECTING AND REARING TERRESTRIAL AND 
FRESHWATER INVERTEBRATES 

F. E. Lutz, P. S. Welch, and J. G. Needham 

MUCH that has been stated in the preceding pages by Dr. Galtsoff 
for marine invertebrates is equally applicable to freshwater forms. 
It will suffice, therefore, if we merely add some notes and suggestions 
concerning methods more applicable to inland aquatic, terrestrial, and 
aerial invertebrates. 

COLLECTING AND HANDLING LIVING SPECIMENS 

For collecting and transporting the larger inland invertebrates (cray- 
fishes, clams, the larger snails, dragonfly nymphs, diving beetle larvae, 
etc.), the small seines, traps, bait pails, and live boxes of com- 



Collecting and Handling 



4i 




Fig. 38. — The apron net. 



merce are everywhere available. Also, for getting small animals out of 
their places of hiding, there are shovels and sifters, rakes and hoes, axes 
and chisels, etc. Some commercial tools especially devised for collecting 
purposes have been illustrated in the preceding article. 

In the following pages we will describe only a few of the most useful 
and most generally applicable devices for collecting and handling in- 
vertebrates. Others will be found in the articles which follow, where 
their special uses will be indicated. Many others may be found by our 
readers if they will consult Pe- 
terson's Manual of Entomolog- 
ical Equipment and Methods, 
which, although prepared pri- 
marily for work on insects, 
contains much that is equally 
applicable to the handling of 
other invertebrates. 

Aquatic animals. A dip net 
(Fig. 1) is perhaps the most 

widely used tool for collecting in freshwater. It must be stout enough 
to stand hard usage and its mesh must be fine enough to retain the ani- 
mals desired. 

The best single tool for collecting the larger aquatic invertebrate 
animals is the apron net (Fig. 38). It is so shaped at the front that it 
may be pushed through beds of weeds or under bottom trash. Its wide- 
meshed cover allows the animals to enter while keeping out the weeds 
and coarser trash. A final push through the water lands the catch at 
the rear where it is easily accessible for picking over by hand. 

The smaller animals that are mixed with the trash in the net may best 
be found by dumping its contents into a white pan of water where they 
will at once reveal their presence by their activity. They may be taken 
from the water most easily and without injury on a lifter made from a 
strip of wire cloth by infolding its edges. 

An apron net is equally satisfactory for scraping up and sifting bot- 
tom mud and sand to obtain burrowers. It may be used for collecting 
insects and other animals from among loose stones in rapid streams 
by setting it edgewise against the bottom facing up stream and stirring 
the stones above it. The animals that are dislodged by the stirring 
will be swept by the current into the net. Old leaf drifts caught in the 
edges of the current may be stirred in the same way to get the animals 
hiding in them, but more stirring and over-turning of the leaves will be 
necessary to dislodge them. 

The small kitchen strainers for sale in any 10-cent store, if securely 
attached to handles, are good for dipping small animals from pools. 



42 Land and Freshwater Animals 

Small aquatic organisms may sometimes be easily collected and con- 
centrated by the use of the following simple field or laboratory device. 
Attach a piece of rubber tubing of appropriate size and length to the 
stem of an ordinary, medium-sized glass funnel. Stretch across the open 
end of the funnel a piece of India linen, bolting cloth, grit gauze, or 
similar material, the mesh of which is such that the organism desired will 
not pass through, and hold in place with a rubber band or cord. Fill a 
pail with water. Place the funnel, wide end down, in the pail but leave 
the longer part of the rubber tubing outside the pail, the free end 
extending below the level of the bottom. Apply momentary suction 
at the free end of the rubber tubing to convert the latter into a siphon 
which will drain the water through the gauze-covered funnel to the 
outside. 

By pouring the water containing the desired organisms into the pail 
the water is gradually eliminated through the funnel but the organisms 
are retained. This process may be continued until the desired concentra- 
tion of the organisms in the pail is reached. 

A cover of wire cloth of wider mesh may be placed over the pail to 
exclude from the catch all larger animals and coarser trash. 

For collecting plankton a standard plankton net may be used (Fig. 
15). If samples for qualitative study only are wanted, a simpler, less 
expensive, and less cumbersome net is more practical. It may be made 
by anyone and consists only of a regularly tapered bag of silk bolting 
cloth attached by a topband to a rather heavy circular rim of non-rusting 
metal. The bag may be 2 or 2% feet deep and its bottom should allow 
easy eversion for the removal of the catch. A cord is attached to the 
rim for towing. 

Removal of the catch in such a net may be facilitated by inserting a 
vial of appropriate size and shape into a small hole at the end of the 
bag. Held in place by a rubber band or a stout thread, such a container 
may be removed easily after the collection is completed. 

If made of No. 25 standard silk bolting cloth, the net will retain all 
but the minutest of the organisms (nannoplankton), but when drawn 
through the water it will clog quickly, pushing much water aside without 
straining it. No. 12 cloth, while not retaining things so small, will strain 
more water and yield a bigger catch of the forms more generally useful 
in the zoological laboratory. 

Small aquatic animals may be taken up on a lifter if not too delicate, 
but they should not be exposed to the air for any considerable length of 
time. In general the more delicate among them are better transferred by 
means of a pipette, without exposure to the air. A hand bulb on a tube 
may be used for the larger entomostracans. 

For isolating single unicellular algae for the production of pure 



Collecting and Handling 43 

cultures the late Dr. A. Brooker Klugh* devised a plunger pipette that 
is a great help in picking out cells of the smallest sizes. We copy in full 
his description and figure of it. 

"This instrument (Fig. 39) consists of: (1) a piece of thin, soft glass tubing 
drawn to a capillary tube at one end; (2) a glass plunger drawn from a piece of 
glass rod to sufficient fineness to fit the capillary tip, and with a flattened knob at 
the other end; (3) a piece of rubber tubing which is placed so as to project beyond 
the glass tube. 

"In the figure the capillary tube is, for the sake of clearness, shown as relatively 
coarse, but in practice this tube should have an inside diameter of 80 micra or less. 

"These parts are so adjusted that the end of the plunger inside the capillary is 



Plwger. W*w 



Plwupr 

Head . 




Fig. 39. — The plunger-pipette. After Klugh. 

about 1 mm. from the end of the capillary, while the knob rests against the rubber. 
This is accomplished by inserting the plunger (which should be made with the fine- 
drawn portion longer than required), and cutting off the part which projects through 
the capillary, then making the fine adjustment by moving the rubber slightly 
upwards. 

"The manner of using this instrument is as follows: A drop or two of water 
containing some of the organisms it is desired to isolate is placed on a slide, the 
organism located, and examined with the 4 mm. objective and a xio, or higher, 
ocular. The desired organism is then located under the 16 mm. or 8 mm. objective. 
The pipette is held with the thumb and second finger just in front of the rubber, 
while the plunger-head is pressed with the first finger so that the end of the plunger 
projects from the tip of the capillary The end of the plunger is brought against the 
organism and the pressure of the first finger released, when the resiliency of the 
rubber withdraws the plunger and the organism is drawn into the end of the 
capillary. (If other organisms, or debris, lie close against the desired organism, they 
may be knocked away by shooting the plunger in and out by the pressure of the 
forefinger.) The organism is then transferred in the pipette to a hollow-ground 
slide containing a drop of the culture medium, and is ejected by a pressure on the 
plunger-head. It is then examined under high power to see that it is absolutely free 
from foreign organisms, picked up with the pipette as before, and transferred to the 
culture-vessel. 

"The chief advantages of this instrument are: 

"1. The plunger does away with the drawing in of undesirable material by 
capillarity. 

"2. The plunger may be employed to clear other organisms away from the organ- 
ism to be isolated. 

"3. The instrument is quick and certain in operation. 

"4. It is easily portable. 

"5. It is simple and requires no special attachments. 

* /. Roy. Micr. Soc. for 1922, p. 2b^ 



44 Land and Freshwater Animals 

"6. It is inexpensive, and several can be kept on hand ready for instant use in 
case of breakages." 

A feather, trimmed at tip and edges to the shape and degree of pliancy 
required, is very useful for holding specimens without injury in any 
desired position for examination, also, for moving delicate specimens 
about. Fish culturists have long used a feather for picking over trout 
eggs on the screens in hatching troughs. For cleaning it is much better 
than a camel's hair brush. Its hooked barbicels catch and lift the dirt 
instead of smoothing it down. 

Certain aquatic animals suffer greatly when carried to the laboratory 
in ordinary collecting receptacles partly or wholly filled with water. 
These may be transferred from the natural habitat to the laboratory 
buried in wet sphagnum moss or placed between layers of cloth or paper 
towels thoroughly soaked with cold water. Towels so used should be 
spread on the bottom of a collecting container and protected from the 
light and heat of the sun. The animals should be restored to a proper 
environment as quickly as possible, and closely watched for a time in 
order that injured specimens may be promptly removed. 

Aerial insects. For collecting flying insects an air net is needed. 
Many kinds will be found advertised in the catalogues of dealers in ento- 
mological supplies. The standard insect net is made with a bag of some 
kind of netting (No. ooo silk-bolting cloth, voile, bobbinet, marquisette, 
cheesecloth, etc., according to one's choice) 12" to 18" in diameter, 
rounded at the bottom, and a convenient arm-length deep. The bag is 
attached by a topband to a circular rim of stiff wire, affixed to a light 
strong handle some three feet long.* 

The net must be used with care to avoid injury to delicate specimens, 
and still greater care must be exercised in handling and carrying and 
caging them after capture. It is bad treatment of living animals to 
dump them in numbers into a bare glass or tin container where they 
cannot get a foothold except by clawing at one another. 

For collecting living specimens of leafhoppers, flea beetles, and other 
small and very agile insects that are prone to jump out of a collecting 
bottle every time it is opened, Mr. Milton F. Crowell of North East, 
Pennsylvania,** suggests fastening a small cone of wire cloth, open apex 
downward, inside the mouth of a collecting vial. The cone serves as a 
baffle. He uses a 1" x 3" shell vial. 

"The cone was made by taking a small square of screen and forcing it into the 

♦Directions for making nets and other entomological collecting apparatus may be found 
in Elementary Lessons on Insects by J. G. Needham, pp. 177-184 (C. C. Thomas, Pub- 
lisher, Springfield, 111., 1928) and in Fieldbook of Insects by Frank E. Lutz, pp. 7-14 and 
PI. 3 (G. P. Putnam's Sons, Publisher, New York City, 1935) • The latter also gives 
some information on rearing methods. 

**/. Econ. Ent. 21:633, 1928. 



Collecting and Handling 



45 



opening of the vial by applying pressure to the center of the screen. When the cone 
was thus shaped, a small hole was cut in its apex. The cone was then placed in the 
vial again and the corners of the screen trimmed off. The cone can then be forced 
a little way into the vial, remaining in place by the spring-action of the bent wire 
against the side of the glass. Care should be taken not to make the cone so small 
that it will drop out. 

Insects that seek the light on emergence from the pupal stage are 
easily collected in a very simple trap. 
They are placed in a dark box before 
emergence. A hole is bored in one 
side of the box, and the open end of 
a glass vial is fitted into the hole. 
Light entering only through this hole 
attracts the insects to enter the vial, 
from whence they are easily removed. 
This is adequate for most minute 
parasites, but for larger and livelier 
insects, such as screw-worm flies,* a 
fruit jar with a cone-shaped baffle 
guarding against return of the flies to 
the box, may replace the vial. 

For picking up minute beetles by 
suction Frank J. Psota of Chicago 
devised an efficient aspirator (Fig. 
40) which he has described** as 
follows: 




Fig. 40.- — A suction-pump collector. 
After Psota. Two longitudinal sec- 
tions through suction-pump collector 
and (to the right) a cross-section 
through the same above the middle, 
looking upward. 



"The apparatus is shown in the accom- 
panying figures: A, is a cork with center 
hole; B, a glass tube 4 inches long, 1% inch 
in diameter, and z /i inch thick ; C : cork of 
type similar to A; D, glass tubing bent 
in S-shape; this curve is very important 
because it destroys a straight path for in- 
sects and dust; E, glass tubing % inch in 
diameter with enlarged edges on both sides of the cork; F, rubber tubing which is 
of the desired length (usually 20 to 30 inches), with mouthpiece on one end, the other 
is slipped over the glass near the cork; G, short piece of rubber tubing which prevents 

♦This is the device used successfully by Mr. D. C. Parmann at the Laboratory of the 
U. S. Bureau of Entomology at Uvalde, Texas. A rim-capped fruit jar is used, with the 
center of the cap left out. The rim is fixed inside a large hole bored in the side of the 
rearing box, and the jar is screwed into it. A cone of wire cloth with an opening at its 
apex large enough to admit the flies, is so shaped that its base is held firmly between the 
jar and the screw cap when these are put together, the apex projecting into the jar. 
Thus the escape of the flies is prevented, until the baffle is removed. 



**Ent. News 27:23, 1916. 



4 6 



Land and Freshwater Animals 



the glass tube from breaking when insects are collected on or around solid objects 
and in crevices ; H, silk netting which is stretched over the end of the tube and tied 
with thread sealed with wax in order to prevent it from fraying ; this netting 
prevents the entrance of dust particles into the tube. 

"The end of the rubber tube G is placed near the objects desired, such as small 
beetles, shells, or any small specimens, which are then drawn into the main chamber 
through the glass tube D, by the suction which is created by a sharp inhalation at 
the end of the rubber tube F. 

"Specimens in the main chamber may be emptied into a cyanide jar by removing 
the bottom cork C, which is pushed into the tube for only about one-third of its 
length." 




Fig. 41. — Beamer's aspirator. Courtesy of Ward's Natural Science Establishment, Inc. 



Another form of aspirator (Fig. 41) employing the same principles 
was devised by Dr. R. H. Beamer of the University of Kansas. 

CAGES AND SHELTER 

In the maintenance of many kinds of aquatic invertebrates a "rearing 
raft" may prove to be very useful. A floating raft or platform of 
appropriate size is anchored in water suitable for the purpose. Sus- 
pended beneath this platform are cages so located and so constructed 
that they maintain the contained animals in approximately the con- 
ditions of the native habitat, and make convenient their examination 
and observation by the investigator. 

One of the most generally useful, most easily constructed, and least 
expensive of cages is the pillow cage (Fig. 42). It is made from a 
single square of woven wire cloth by doubling and closely folding two 
opposite edges to form a cylinder, and then in like manner cross-folding 
the ends. The folds must be crimped tightly and evenly. A square 
yard of the cloth quartered makes four cages of the size most commonly 
used for insects. Cages made from small-wire cloth may be folded with 




Fig. 42. — A pillow cage. 



Cages and Shelter 47 

the fingers but larger and stronger ones will require a small tinker's 
folding tongs. A woven edge should form the top, so that there be no 
wire ends to prick the fingers on opening and closing the cage. 

Such a cage is very adaptable. It may be used for carrying home a 
catch, since its walls afford a foothold, and its crevices at the ends afford 
hiding places. It may be hung in 
a tree or buried under trash or im- 
mersed in a pond, to hold hiber- 
nating animals in safety from 
their predatory enemies. It may 
be used for distributing parasitic 
insects in a grove by placing para- 
sitized pupae within it and hang- 
ing it in a tree; the mesh will then 
have to be of a size to permit the 
escape of the parasites, while re- 
taining their injurious host insects. 
When rearing the insects that feed 
on a growing plant one end may be 
fitted over a flower pot contain- 
ing the plant. 

When used as a transformation cage for aquatic insects such as 
dragonflies, stoneflies, and mayflies, it should be set aslant in the water 
with only the lower end immersed and plenty of room above for ex- 
panding wings. If any adults chance to fall back into the water, the 
sloping sides will facilitate their crawling out again. 

Hollow-ground slides capped with a cover glass are often used as 
rearing cages for organisms of microscopic size. Dr. Marshall Hertig 
(Science 83: no, 1936) has suggested a method of making them in any 
desired shape or size. 

"The essential apparatus for turning out these laboratory-made slides is an 
electric motor (that of an electric fan will serve), a flexible shaft provided with a 
chuck or "handpiece" into which may be fitted any of the dentist's arsenal of 
burrs, drills and abrasive devices. Of these the most generally satisfactory for 
grinding glass are the abrasive wheels, which consist of small disks of carborundum 
or other material mounted on a mandrel, and which are available in a variety of 
diameters, thicknesses and degrees of abrasiveness. Abrasive "points," i.e., small 
carborundum spheres, cones and cylinders, may also be used, but are much less rapid 
than the abrasive wheels on account of their small diameter and hence low velocity 
of grinding surface. 

"The process of grinding a depression consists merely of placing a drop of water 
on the slide and applying the abrasive instrument. Very little spattering occurs. 
The most rapidly ground depression is the slot made by the edge of the carborundum 
wheel. A cavity of this shape is desirable for elongate specimens. By moving the 
wheel while grinding, a depression of almost any size and shape may be made, and 



48 Land and Freshwater Animals 

rotating the slide on a turn-table produces a circular concavity similar to that 
of the ordinary hollow-ground slide." 

CAGE MANAGEMENT 

In the maintenance of animal cultures the big factors are food, 
temperature, humidity, and sanitation. 

Feeding and watering. Proper feeding is of first importance in the 
maintenance of animal cultures. Errors are easily made in both amount 
and kind of food. The amount should be all that will be completely 
consumed. Over-feeding is a more common error than under-feeding. 

Watering is often as important as feeding, and he who reads the 
articles that follow will find many ingenious devices for supplying drink 
while avoiding fatalities from drowning in the pool. For the larger 
animals commercial chick-watering and rat-watering devices are avail- 
able. For smaller ones there is such provision as a wide-mouthed bottle 
filled with water and closed by placing over its mouth a petri dish lined 
with a sheet of filter paper, the whole then inverted and set on the 
cage floor. (See p. 433.) Very small insects are supplied both food 
and drink from open test tubes of capillary smallness filled with liquid 
and placed where accessible within the cage. 

Temperature and humidity. For ordinary rearing work accurately 
stabilized temperature and humidity are rarely needed, since the animals 
are well inured to a considerable range of both. But there is much need 
for the exercise of judgment in the location of cages with respect to 
both these factors. 

Cultures which require a certain uniformity of surrounding tempera- 
ture may be handled in the following ways if regular temperature control 
apparatus is not available or is not required. For organisms not re- 
quiring light an under-ground cave, a subterranean room, or some similar 
below-ground space may afford a fairly regular temperature throughout 
the year. For organisms requiring light, culture jars may be placed in 
another container through which a stream of water from some constant 
source is running. Such provision is often very satisfactory providing, 
of course, that the water passing through this improvised jacket is rela- 
tively uniform and of the desired temperature. Workers in lakeside 
laboratories may sometimes secure the desired uniform temperature by 
suspending culture containers on ropes hung from a float or buoy in the 
deep water of a lake. The temperatures at different depths may be 
determined in advance by the use of an appropriate recording ther- 
mometer. Since the deeper waters may change little if at all over the 
desired period, this method is often a convenient one, and it offers the 
possibility of selecting temperatures in the vertical temperature gradient. 
Modern refrigeration with thermostatic control has made possible the 



Cage Management 49 

determination of optimum temperature and the limits of tolerance for 
many species, and has provided the means for maintaining any constant 
temperature that may be needed for careful experimental work. 

Insufficient humidity is a most frequent cause of failures of cultures 
of terrestrial animals. In the following articles will be found many means 
of adding moisture such as sprinkling, burying in the ground, placing 
inside the cage water-holding stuffs (filter paper, sphagnum moss, peat, 
sponges, paper towels, fresh green leaves, etc.), conducting water inside 
by means of wicks, dispersing it from porous earthenware containers 
by capillarity, allowing evaporation from open pans, spraying it in the 
cage, etc.* One of us (Lutz) even in keeping scorpions gives them a 
shower bath from a bulb spray every day. Although they are desert 
creatures they normally hide in relatively moist places except at night 
or when the air outside their hiding places is moist. 

On the other hand, too much moisture may be fatal. It frequently 
brings disaster by favoring the growth of molds. 

Sanitation. Keeping cages clean is very important. Sterilization of 
containers by heat (steam, autoclave, etc.) and by chemicals is practiced 
in many ways, and sterilization of food, drink, and shelters as well. 
Wooden tubs are prepared for use by charring the interior in a flame. 
Small containers are closed against infection by sealing the lids with vase- 
line. Excreta and uneaten food require prompt removal. Cages of cer- 
tain kinds may be washed, or they may be supplied with removable bot- 
toms, or the floor may be covered with absorbent stuffs like sawdust or 
sphagnum. 

Sick specimens must be isolated to prevent the spread of infection. 

Note: The following note concerning Pablum has been sent us by William LeRay 
and Norma Ford, University of Toronto. "As a food for many invertebrate 
and vertebrate animals, we are now using a pre-cooked cereal, devised in the 
Research Laboratories of the Department of Pediatrics, University of Toronto, 
and sold under the name of Pablum by Mead Johnson & Co., Evansville, Ind. 

"Insects which take dry food, such as ants, flour moths, etc., are fond of it ; 
burrowing crayfish grow as rapidly on it as when living outdoors ; it is excellent for 
earthworms, as well as for fish, birds, etc. To thrushes this cereal is particularly 
acceptable. 

"Pablum consists of wheatmeal, oatmeal, wheat embryo, yellow cornmeal, pow- 
dered beef bone, dried yeast, powdered dehydrated alfalfa leaf, and sodium chloride. 
It has not only high nutritive value, but also furnishes substantial amounts of 
vitamins A, B, E, and G, and essential mineral elements, calcium, phosphorous, iron, 
and copper." 

* For chemical methods consult Spencer, Hugh M., Laboratory methods for maintaining 
constant humidity. Internal. Critical Tables 1:67-68, 1926. 



50 Land and Freshwater Animals 

ADDENDA 

THE progress of technique during the past year has made it desirable 
to add the following notes. The information they contain is es- 
sential, and it brings the methods down to date. 

Note i (supplemental to p. 16) — Extensive use of celluloid in the 
laboratory may be objectionable on account of its inflammability. To 
avoid fire hazards a so-called "Plastocele" can be used. This product 
manufactured by the DuPont Viscoloid Company, Arlington, N. J., 
closely resembles celluloid in appearance and general properties but has 
great advantage over it in being extremely slow burning and difficult to 
ignite. For dissolving and cementing plastocele a mixture of equal 
amounts of methyl acetone and methyl cellosolve is used. 

p. s. G. 

Note 2 (supplemental to p. 26) — The operation of the siphon can 
be greatly improved by blowing a bulb at the end of the upper arm of 
it and boring a small hole one-half inch above the tip of the lower arm. 

p. s. G. 

Note 3 (supplemental to p. 26) — -The water is forced by air bubbles 
into a Wolff bottle which serves to maintain an even flow into the 
aquarium. The overflow through the automatic siphon is carried into 
the top of the gravel and sand filter from which it returns to the 
reservoir. 

p. s. G. 



Phylum I 

Protozoa, Class Mastigophora 



GROWTH OF FREE-LIVING PROTOZOA 
IN PURE CULTURES 

R. P. Hall, University College, New York University 

INTRODUCTION 

THE pure-culture technique offers certain definite advantages in the 
maintenance of free-living Protozoa in the laboratory. As a depend- 
able source of material for class use, bacteria-free cultures far surpass in 
value the usual hay infusions. Thus, a number of species of flagellates 
and ciliates have been maintained in our laboratory over periods ranging 
from two to six years, with an abundant supply of each type always avail- 
able. In a suitable medium, bacteria-free cultures remain viable for 
several months; hence frequent transfers are unnecessary for the 
maintenance of stock cultures. The technique is simple and requires rela- 
tively little equipment and no more than a rudimentary knowledge of 
bacteriological procedures. Bacteria-free cultures are even more valu- 
able as a source of material for experimental studies. In physiological 
investigations the advantages of the elimination of bacteria in the precise 
control of experimental conditions are obvious. Biochemical investiga- 
tions, impossible a few years ago, may now be carried out on free- 
living Protozoa with almost the same facility as in the case of bacteria. 
In short, the establishment of bacteria-free cultures opens to the pro- 
tozoologist, physiologist, and biochemist a wide field of investigation 
which promises to add much to our knowledge of the morphology, life 
history, and metabolism of Protozoa. 

Two general methods have been followed in the growth of Protozoa in 
the absence of other living micro-organisms. In the first, the organisms 
have been washed free of bacteria and grown in sterile peptone solutions 
or similar media. In the other method, used particularly for ciliates, 
the Protozoa have first been freed from bacteria and then placed in 
suspensions of killed bacteria or other micro-organisms. Ciliates and 
other Protozoa have also been grown by various workers on single strains 
of living bacteria, yeasts, algae, and small Protozoa. 



52 Phylum Protozoa 

Several investigators (Parpart, 1928; Luck and Sheets, 1931; Hether- 
ington, 1934) have described relatively simple methods for washing 
Protozoa free from bacteria. The method of Parpart, a method which 
offers few technical difficulties, has been used successfully in our labora- 
tory and elsewhere. In this method it is a comparatively easy matter to 
free the Protozoa from bacteria, although it is often very difficult to estab- 
lish thriving cultures of the bacteria-free organisms. 

MEDIA FOR CHLOROPHYLL-BEARING FLAGELLATES 

For obvious reasons, the chlorophyll-bearing flagellates are more 
easily established in bacteria-free cultures than is any of the other groups 
of free-living Protozoa. This is evidenced by the fact that approximately 
40 species of green flagellates have been isolated in such cultures, most of 
the strains being maintained at the present time in the laboratory of 
Professor E. G. Pringsheim (Pringsheim, 1928a, 1930). The composi- 
tion of the various media, so far as inorganic constituents are concerned, 
is based upon past experience in the growth of algae and green flagellates. 
With the establishment of bacteria-free cultures, peptones or other organic 
sources of nitrogen have usually been added to the media for the mainte- 
nance of rich cultures. Several of the media which have been tried in our 
laboratory and have given good results with different species of Euglenida 
and Phytomonadida are listed in Table 1. 

table 1. Media for chlorophyll-bearing flagellates. 

Media 

Constituents ABC D E F 

NH4NO3 0.5 gm 1.0 gm. 

KNO3 0.5 gm 0.5 gm. 

Tryptone 2.5 gm 2.0 gm. . . .5.0 gm. 

Glycine 1 -99 6 gm. 

K 2 HP0 4 0.209 gm. 

KH2PO4 0.5 gm. . . .0.5 gm. . . .0.2 gm. . . .0.25 gm. . .0.5 gm. 

MgS0 4 0.1 gm. . . .0.1 gm. . . .0.2 gm 0.25 gm. . .0.25 gm . .0.048 gm. 

NaCl 0.1 gm. . . .0.1 gm 0.1 gm. 

KC1 0.2 gm .... 0.25 gm. 

FeCI 3 Trace Trace Trace Trace 

Sodium acetate 2.5 gm 2.0 gm 1 .48 gm. 

Dextrose 2.0 gm. 

Distilled water . ...i.ol 1.0 1 1.0 1 1.0 1 0.0 1 1.0 1 

Medium A, adjusted to pH 7.0, has given very good results (Loefer, 
1934) with Chlorogou'nim euchlorum and C. elongatum, and should be 
equally satisfactory for other species of Chlamydomonadidae. Medium 
B, at pH 7.0, has supported growth of Chlorogonium and C. elongatum 
(Loefer, 1934) in darkness for more than a year, the flagellates retaining 



Mastigophora 53 

their chlorophyll throughout the period of culture. The same medium 
has been used for Haematococcus pluvialis, Colacium vesiculosum, and 
five species of Euglena, in the maintenance of stock cultures in light. 
Medium C at pH 7.0 supports good growth of Euglena gracilis, according 
to the findings of Dusi (1930) and results obtained in our own laboratory. 
Ammonium phosphate or ammonium sulphate may be substituted for 
ammonium nitrate. Medium D is the formula used by Lwoff and Lwoff 
(1929) for Chlamydomonas agloejormis and Haematococcus pluvialis 
and by Lwoff and Dusi (1929) for Euglena gracilis in darkness, except 
that Difco tryptone is substituted for the peptone used by the French in- 
vestigators. In our laboratory, this medium has given excellent results 
with stock cultures of six species of Euglenida, Haematococcus pluvialis, 
and two species of Chlorogonium. Medium E was used by Jahn (1931) 
for Euglena gracilis and by Hall (1933) for E. anabaena and E. deses. In 
addition, the medium is satisfactory for other species of Euglena, and for 
Haematococcus and Chlorogonium. Growth is less abundant than in 
media containing sodium acetate, but the cultures remain viable for 
several months. Medium F, at pH 7.0, supports good growth of 
Chlorogonium elongatum, C. euchlorum, and Haematococcus pluvialis. 
The same medium supports slow growth of the colorless Chilomonas Para- 
mecium, as reported by Mast and Pace (1933) and confirmed in our 
laboratory. 

In addition to the various liquid media, certain types of solid media 
have proved useful in our laboratory, particularly for the growth of stock 
cultures over long periods and also in the preparation of cultures for 
shipment. In the preparation of a solid medium we have added either 
Difco dehydrated dextrose-agar or starch agar to one of the media listed 
above; for a semi-solid medium, one-half or less of the usual amount of 
dehydrated agar is added to the liquid. Such media may be tubed and 
then slanted or not, as preferred. Slant or stab cultures may be sealed 
with melted paraffin, and will usually remain healthy for several months. 

MEDIA FOR COLORLESS FLAGELLATES 

In general, the addition of peptone in suitable amounts to one of the 
media (A-F) listed above will provide a satisfactory medium for growth 
of the colorless Phytomastigophora. The formulae have been varied 
somewhat by different workers, however, as indicated in Table 2. 

Medium G was used by Pringsheim (1921) for growth of Polytoma 
uvella, and good results were obtained also by Lwoff ( 1932 ) . The same 
solution, in Pringsheim's laboratory, was made the base of an agar medium 
which supported good growth of Polytoma. Medium H is a gelatin 
medium developed by Lwoff (1932) for Polytoma uvella; with the omis- 
sion of gelatin, the same formula gives a very satisfactory liquid medium 



54 Phylum Protozoa 

table 2. Media for colorless flagellates. 

Media 



Constituents G H I J 

Glycine 2.0 gm. 

Peptone 2.0 gm 10. o gm 10.0 gm. 

Gelatin iS°-° g m - 

NH4NO3 0.5 gm. 

Dextrose 2.0 gm. 

Sodium acetate 2.0 gm 2.0 gm 2.0 gm. 

K2HPO4 0.2 gm 2.0 gm. 

KH2PO4 0.25 gm 1.5 gm. 

K2CO3 2.0 gm. 

KC1 0.25 gm. 

MgS0 4 0.1 gm 0.25 gm 0.25 gm. 

Distilled water 1.0 1 1.0 1 1.0 1 

Tap water 1.0 1 

table 3. Media for ciliates. 

Media 

Constituents K L M N P 

Peptone 10. o gm . . . 10. o gm. . . 10-30 gm 5.0 gm. 

KH 2 P0 4 2.0 gm. 

K2HPO4 0.02 gm. 

Na 2 HP0 4 0.01 gm. 

NaCl 0.5 gm. . . .4.0 gm 0.003 g m - • °- 02 S m - 

KC1 0.01 gm. 

MgSC"4 0.01 gm 0.025 g m - • 0.0045 g m • °-° 2 § m - 

CaCl. 0.01 gm 0.001 gm. 

KNO3 0.5 gm 0.0013 gm. 

(NH 4 ) 2 P0 4 0.05 gm. 

FeCU 0.001 gm. . .0.0002 gm. 

CaS0 4 0.015 g m - 

Ca(N0 3 ) 2 0.2 gm. 

FeS0 4 Trace 

Distilled water . .1.0 1 1.0 1 1.0 1 1.0 1 1.0 1 1.0 1 

for the same species. Lwoff adjusted the pH to 8.0 with K2CO3. 
Medium I was used by Jahn (1933) for the growth of stock and experi- 
mental cultures of Chilomonas Paramecium. Medium J has been used 
in our laboratory for maintaining stock cultures of Astasia ocellata (Jahn 
strain), Astasia sp. (Jahn strain), and Chilomonas Paramecium. 

MEDIA FOR CILIATES 

The growth of Colpidium striatum and C. campylum in various types 
of media has been investigated by Elliott (1935a), and that of Glau- 
coma ficaria and G. piriformis by Johnson ( 1936). Results obtained by 



Mastigophora 55 

these investigators, as well as our general experience in the maintenance 
of stock cultures, indicate that simple media, containing one of the Difco 
peptones (tryptone particularly), are altogether suitable for growth of 
the four species mentioned. Other investigators have previously used 
more complex media for Colpidium and Glaucoma, but it seems that, ex- 
cept in special cases, little is to be gained by the use of complex salt 
solutions in addition to peptone. Occasionally, however, such simple 
media may fail to support growth of the ciliate; Glaser and Coria ( 1935) 
have recently confirmed their earlier reports that a special medium is 
necessary for growth of Paramecium whereas a relatively simple medium 
is suitable for Chilodon and Trichoda. Some of the simpler formulae, 
which have been tried in our laboratory and found satisfactory, are listed 
in table 3. 

Medium K was used by Lwoff ( 1929) in his earlier work with Glaucoma 
piriformis. Although the medium was very satisfactory, Lwoff later 
concluded that such a complex medium is unnecessary and accordingly 
substituted medium L (Lwoff, 1932) in his later investigations. Medium 
M was used by Elliott (1935, 1935a) for Colpidium striatum and C. 
campylum. Difco tryptone was found to be superior to a number of other 
commercial peptones, although Difco proteose-peptone and neopeptone 
were fairly satisfactory. Tryptone gave best results with Colpidium in 
concentrations of 1.0-3.0%. Johnson (1936) has found that a 1.5% 
trjrptone medium is optimal for growth of Glaucoma ficaria, while Loefer 
(1936a) has found it necessary to reduce the concentration to 0.5% for 
Paramecium bursaria. Formula N was devised by Pringsheim (1928) 
for P. bursaria. In Loefer's (1936a) experience, the addition of Difco 
tryptone (0.5%) or better, Difco proteose-peptone (0.5%), results in a 
medium very satisfactory for the growth of this ciliate in bacteria-free 
cultures. Medium O was made up by Loefer (1936a) on the basis of 
tap water analyses, since it had been found that tap water (New York 
City) gave excellent results in the cultivation of several species of Pro- 
tozoa. Either tryptone or proteose-peptone is added to the salt solution 
in a concentration of 0.5%. Medium P was used for Paramecium bur- 
saria by Pringsheim (1928, p. 310) ; with tryptone or proteose-peptone 
added, fhis formula gives a medium satisfactory for growth of the species 
in bacteria-free culture. 

For another medium which has given good results with several bacteria- 
free strains of ciliates, the reader is referred to the work of Glaser and 
Coria (1935). Since this medium requires blood serum in its prepara- 
tion, it will be less generally useful than the simpler media described 
above, but it should be quite useful in laboratories in which the necessary 
technical facilities are available. 



56 Phylum Protozoa 

MEDIA FOR SARCODINA 

Acanthamoeba castellanii has been grown by Cailleau ( 1933) in a liver 
extract-serum medium sterilized by filtration, and later (1933a) in a 
peptone medium of the following composition: peanut peptone (Vaillant), 
3.0'; ; sodium chloride, 0.4% ; magnesium sulphate, 0.001% ; monopotas- 
sium phosphate, 0.001%. Other peptones found to be satisfactory were: 
pancreatic stomach peptone, pancreatic peptone of spleen, and peptic 
peptone of "delipo'ide" liver, all three being Vaillant preparations. With 
peptone in a concentration of 3.0%, growth of Acanthamoeba was ob- 
tained through successive transfers, while low concentrations of peptone 
were found to be unsatisfactory, as reported previously by Lwoff ( 1932 ) . 

GROWTH OF PROTOZOA ON KILLED MICRO-ORGANISMS 

The growth of several species of amoebae on killed bacteria was re- 
ported by Tsujitani (1898) and Oehler (1916, 1924, 1924a). Re- 
views of early and later investigations are to be found in papers by 
Wulker (1911) and Sandon (1932), respectively. Among the ciliates, 
Colpoda steiniiwas grown by Oehler ( 19 19) on killed bacteria and yeasts; 
Glaucoma scintillans, on dead bacteria by E. and M. Chatton (1923); 
Paramecium caudatum, on dead yeasts and bacteria by Glaser and Coria 
(1933); and Glaucoma ficaria by Johnson (1936a) on dead bacteria, 
yeasts, and flagellates. Negative results have been obtained by a number 
of other workers. The literature has been reviewed by Sandon (1932) 
and Johnson (1936a). 

GROWTH OF PROTOZOA ON SINGLE SPECIES 
OF MICRO-ORGANISMS 

The maintenance of pure-line cultures of Protozoa on single species of 
living bacteria, yeasts, or other micro-organisms has been accom- 
plished by a number of investigators, particularly Oehler (1919, 1924, 
1924a). More recent investigations include those of Geise and Taylor 
(1935), Loefer (1936), and Johnson (1936a). The literature has been 
reviewed by Sandon (1932) and Johnson (1936a). Such methods are 
much more reliable than the use of the common "infusion" cultures and, 
while they do not offer the precision of the bacteria-free technique, the 
procedures are relatively simple and are often worth while in view of 
the good results obtained. 

GROWTH IN RELATION TO pH AND OTHER FACTORS 

The observations of a number of workers have shown that the pH of 
the medium is an important factor influencing the success or failure 
of bacteria-free cultures of Protozoa. The findings of Dusi (1930a), 
Jahn (1931), Elliott (1933), Dusi (1933, i933a), Hall (1933), Loefer 



Mastigophora 57 

(1935), and Johnson (1936) show that growth of various species of flag- 
ellates and ciliates in bacteria-free cultures is influenced to a marked 
degree by the pH of the medium. The literature on this subject has been 
reviewed by Loefer (1935). Not only is there a direct relationship 
between pH and growth in the simpler media, but there are also indirect 
relationships to be considered. For example, it is known (Elliott, 1935a) 
that sodium acetate accelerates growth of Colpidium in a medium near 
pH 7.0, but is decidedly toxic in media below pH 5.5, although the latter 
pH is near the optimum for growth of the ciliate in acetate-free media. 
Observations in progress on several species of Euglena indicate that 
similar relationships hold for the green flagellates. Elliott (1935) has 
noted a similar relationship between pH of the medium and the accel- 
erating effects of carbohydrates on growth of Colpidium. Near the neu- 
tral point, little or no acceleration was observed with certain carbo- 
hydrates, whereas in the acid range growth was decidedly stimulated by 
the same carbohydrates. 

The concentration of both organic and inorganic constituents of the 
medium may also have an important bearing on the maintenance of 
bacteria-free cultures, as indicated by the observations of Cailleau 
(1933a), Elliott (1935a), Loefer (1936a), and Johnson (1936). It is 
known that the addition of various substances — such as dextrose and 
other carbohydrates, sodium acetate and other salts of the lower fatty 
acids, yeast extract, etc. — to media listed above will increase the growth 
of a number of species of Protozoa. Our observations show that the 
factor of concentrations is important in the determination of such acceler- 
ating effects, since an amount in excess of the optimal concentration may 
even inhibit growth of the Protozoa. This question has been discussed in 
detail by Loefer (1936a). 

References 

For the culture of many colorless and pale green flagellates see p. 177. 
Order Chrysomonadida, Family Ochromonadidae 
For the culture of Ochromonas see p. 62. 

Bibliography 

Cailleau, R. 1933. Culture d'Acanthamoeba castellani en milieu liquide. C. R. 
Soc. Biol. 113:990. 

1933a. Culture d'Acanthamoeba castellani sur milieu peptone. Action sur 

les glucides. Ibid. 114:474. 

Dusi, H. 1930. La nutrition autotrophe d'Euglena gracilis Klebs aux depens de 
quelques azotes inorganiques. Ibid. 104:662. 

1930a. Les limites de la concentration en ions H pour la culture de quel- 
ques Euglenes. Ibid. 104:734. 

1933. Recherches sur la nutrition de quelques Euglenes. 



I. Euglena gracilis. Ann. Inst. Pasteur 50:550. 
1933a. Recherches sur la nutrition de quelques Euglenes. 



58 Phylum Protozoa 

II. Euglena stellata, klebsii, anabaena, deses et piscijormis. Ibid. 50:840. 
Elliott, A. M. 1933. Isolation of Colpidium striatum Stokes in bacteria-free 

cultures and the relation of growth to pH of the medium. Biol. Bull. 65:45. 
1935- Effects of carbohydrates on growth of Colpidium. Arch. j. Protist. 

84:156. 

1935a. Effects of certain organic acids and protein derivatives on the growth 



of Colpidium. Ibid. 84:472. 
Geise, A. C, and Taylor, C. V. 1935. Paramecium for experimental purposes in 

controlled mass cultures on a single strain of bacteria. Ibid. 84:225. 
Glaser, R. W. 1932. Cultures of certain Protozoa, bacteria-free. J. Parasit. 19:23. 
Glaser, R. W., and Coria, N. A. 1930. Methods for the pure culture of certain 

Protozoa. J. Exper. Med. 51:787. 

1933- The culture of Paramecium caudatum free from living micro- 
organisms. J. Parasit. 20:33. 

1935- The culture and reactions of purified Protozoa. Amer. J. Hyg. 



21:111. 
Hall, R. P. 1933. On the relation of hydrogen-ion concentration to the growth 

of Euglena anabaena var. minor and E. deses. Arch. f. Protist. 79:239. 
Hetherington, A. 1934. The sterilization of Protozoa. Biol. Bull. 67:315. 
Jahn, T. L. 1931. Studies on the physiology of the euglenoid flagellates. 

III. The effect of hydrogen-ion concentration on the growth of Euglena gracilis 
Krebs. Ibid. 61: 387. 

1933- Studies on the oxidation-reduction potential of protozoan cultures. 

I. The effect of pH on Chilomonas Paramecium. Proto plasma 20:90. 

Johnson, D. F. 1936. The isolation of Glaucoma ficaria Kahl in bacteria-free 
cultures, and growth in relation to pH of the medium. Arch. f. Protist. 86:263. 

1936a. Growth of Glaucoma ficaria Kahl in cultures with single species 

of other micro-organisms. Ibid. 86:359. 

Loefer, J. B. 1934. The trophic nature of Chlorogonium and Chilomonas. Biol. 
Bull. 66:1. 

1935- Relation of hydrogen-ion concentration to growth of Chilomonas 

and Chlorogonium. Arch. /. Protist. 85:209. 

1936. A simple method for the maintenance of pure-line mass cultures of 



Paramecium caudatum on a single species of yeast. Trans. Amer. Micr. Soc. 

55:255- 

1936a. Isolation of a bacteria-free strain of Paramecium bursaria and con- 



centration of the medium as a factor in growth. (Submitted for publication.) 
Luck, J. M., and Sheets, G. 1931. The sterilization of Protozoa. Arch. f. Protist. 

75:255. 
Luck, J. M., Sheets, G., and Thomas, J. O. 1931. The role of bacteria in the 

nutrition of Protozoa. Quart. Rev. Biol. 6:46. 
Lwoff, A. 1929. Milieux de culture et d'etretien pour Glaucoma piriformis (Cilie) . 

C. R. Soc. Biol. 100:635. 
— 1932. Recherches biochimiques sur la nutrition des Protozoaires. Le pou- 

voir synthese. Monographies de I'Institut Pasteur (Paris, Masson et Cie.), 158 pp. 
Lwoff, A., and Dusi, H. 1929. Le pouvoir synthese d'Euglena gracilis cultivee 

a l'obscurite. C. R. Soc. Biol. 102:567. 
Lwoff, M., and Lwoff, A. 1929. Le pouvoir de synthese de Chlamydomonas 

agloejormis et d'Haematococcus pluvialis. Ibid. 102:569. 
Mast, S. O., and Pace, D. M. 1933. Synthesis from inorganic compounds of starch, 

fats, proteins and protoplasm in the colorless animal, Chilomonas Paramecium. 

Protoplasma 20:326. 



Cryptomonadidae 59 

Oehler, R. 1916. Amobenzucht auf reinem Boden. Arch. f. Protist. 37:175. 

1919. Flagellaten- und Ciliatenzucht auf reinem Boden. Ibid. 40:16. 

1920. Gereinigte Ciliatenzucht. /bid. 41:34. 

1924. Weitere Mitteilungen iiber gereinigte Amoben- und Ciliatenzucht. 

Ibid. 49:112. 

1924a. Gereinigte Zucht von freilebenden Amoben, Flagellaten, und Ciliaten. 



Ibid. 49:287. 
Parpart, A. K. 1928. The bacteriological sterilization of Paramecium. Biol. Bull. 

55:ii3. 
Phelps, A. 1934. Studies on the nutrition of Paramecium. Arch. j. Protist. 

82:134. 
Pringsheim, E. G. 1921. Zur Physiologie saprophytischer Flagellaten (Polytoma, 

Astasia und Chilomonas). Beitr. Allg. Bot. 2:88. 
1928. Physiologische Untersuchungen an Paramecium bursaria. Arch. j. 

Protist. 64:289. 

1928a. Algenreinkulturen. Eine Liste der Stamme, welche auf Wunsch 



agbegeben werden. Ibid. 63:255. 

1930. Neue Chlamydomonaceen, welche in Reinkultur gewonnen wurden. 



Ibid. 69:95. 
Sandon, H. 1932. The food of Protozoa. Publ. Fac. Sci. Egypt (Cairo, Misr-Sokar 

Press), 187 pp. 
Tstjjitani, J. 1898. Ueber die Reinkultur der Amoben. Zentralbl. f. Bakt., I, 

24:666. 
Wulker, G. 1911. Die Technik der Amobenzuchtung. Ibid. 33:314- 

Order cryptomonadida, 
Family cryptomonadidae 

CULTIVATION OF PROTOZOA 

John P. Turner, University of Minnesota 
Chilomonas sp. To each 100 cc. of pond water* (previously freed from 
Protozoa by heating to 70 C.) add 4 grains of wheat. Inoculate with 
Chilomonas from old culture or from wild stock after isolating them 
with a fine pipette [see pp. 43, 7°] • Within a week the culture should be 
cloudy with Chilomonas, and this serves as a valuable source of food for 
many larger forms.- Chilomonas may be obtained from almost any 
pond where there is decomposing vegetation, especially if that be allowed 
to stand in water in the laboratory for a few days with wheat added. 

Amoeba proteus. To each 100 cc. of pond water (previously heated 
to 70 C. if a "pure" culture is wanted) add 2 grains of wheat and a few 
drops of Chilomonas culture to serve as food. A day or two later inocu- 
late with the Amoeba (often found on dead lily pads, etc., in shallow 
water). Keep between 15 and 25 C. for best results, and disturb as 
little as possible. Cultivate in water less than an inch deep. 

* Note: In most parts of the country tap water from city mains will grow Protozoa 
satisfactorily if allowed to stand for a few days after being drawn. In many cases pond, 
lake, spring, or stream water is better. 



60 Phylum Protozoa 

Actinosphaerium eichhorni. To each ioo cc. of pond water add 4 grains 
of wheat, some Chilomonas and, if available, any other small Protozoa. 
Inoculate with Actinosphaerium. Culture in a flat, shallow dish. Con- 
siderable search of small, permanent ponds may be necessary before 
Actinosphaerium are found, but by repeated inoculation of a number of 
rich mixed cultures with each sample taken, one should soon discover 
them. 

Arcella vulgaris. Both wheat and hay infusions are good, but a mix- 
ture is best. To 100 cc. of pond water add 2 grains of wheat and % 
gram of hay. Inoculate with Chilomonas if available, although Arcella 
will grow by feeding merely on the decomposing infusion. After two or 
three days add Arcella which may be found on the bottom of many old 
cultures or in the bottom ooze from any shallow pond. Isolate them from 
the ooze and other Protozoa with a fine pipette [see p. 43] and place 
them in a watch glass with a drop of water. See that there are no other 
Protozoa nor worm eggs present that might develop and feed on the 
Arcella. Then place them in the culture in a shallow dish. 

Euplotes patella. To a liter of water add 5 grams of timothy hay, 
10 halves of yellow split peas, and 10 grains of wheat. Heat to boiling 
point and set aside until the next day. Inoculate with Chilomonas which 
serves as food. In a few days these will be abundant and the culture is 
ready for the Euplotes. When kept in such medium, Euplotes will multi- 
ply to incredible numbers. 

Blepharisma sp. Follow directions given for Euplotes patella but 
dilute medium with equal parts of pond water before adding Blepharisma. 

Spirostomum ambiguum and 5. teres. Putrid cultures are best. Rich 
wheat and hay infusions give excellent results. To each 100 cc. of water 
add 5 grains of wheat and 5 or 10 2-inch lengths of timothy hay stalks. 
Inoculate with Chilomonas for food and after a day or two add the 
Spirostomum. If available, bed the bottom of the dish with old, thor- 
oughly washed sphagnum moss. They should multiply to such numbers 
as to make large white blotches on the moss. 

Stentor coeruleus. Fill a battery jar % to % full of a culture of Para- 
mecium prepared as follows: One liter of lake water is brought to a 
boil and a handful of timothy hay added; as soon as all the hay has been 
submerged in the boiling water the heat is turned off and, on the follow- 
ing day, the infusion is diluted with an equal portion of lake water and 
inoculated with the Paramecia. In a week or so the Paramecia are suffi- 
ciently abundant. Then add a gram or two of timothy hay and inoculate 
with Stentor. If kept near a window but not in direct sunlight, the 
Stentor should become very abundant. With occasional (every 2 or 3 
weeks) additions of small amounts of raw, dry hay, rich cultures will 
last for months. 



Euglenidae 61 

References 

For the culture of Chilomonas see also pp. 62, 63, and 136. 

For the culture of Chilomonas paramedian see pp. 53 and 113. 
Order Phytomonadida 

For the culture of various members, of this order see p. 52. 
Family Chlamydomonadidae 

For the culture of Chlamydomonas see p. 53. 

For the culture of Haematococcus pluvialis see p. 53. 

For the culture of Chlorogonium euchlorum and C. elongation see p. 52. 

For the culture of Carteria see pp. 35 and 38. 
Family Volvocidae 

For the culture of Volvox see references on p. 72. 



Family polytomidae 



POLYTOMA CULTURES* 

Josephine C. Ferris, University of Nebraska 

PUT 2 70 grams of chicken-size bone meal into a muslin bag and tie 
compactly so that the whole mass is in a firm ball. Cover with water 
and bring to a boil. Pour off water and allow the bag to cool for 2 hours. 
Boil 1 gram of ground timothy hay for 10 minutes and allow to cool. 
Pour cooled hay solution into small battery jar and add enough cold 
sterilized water to make 1200 cc. Lift cooled bag of bone meal with 
sterilized forceps into jar and inoculate with Polytoma. Keep at about 
1 7 to 20 C. Every 48 hours repeat the above procedure and use all the 
scum of the old culture in inoculating the new culture. Such a culture 
may be maintained many months or even years. Removing the surface 
film of Polytoma with a sterilized spoon and centrifuging yields con- 
centrated masses of these Protozoa. 

References 

For the culture of Polytoma see also p. 53. 

For the culture of Parapolytoma satura see pp. 113 and 116. 

Order euglenoidida, Family euglenidae 

CULTURING EUGLENA PROXIMA 

J. A. Cederstrom, University of Minnesota 

Euglena proximo, may be cultured very readily for laboratory use in 
filtered rain water (100 cc.) to which is added 1 cc. of one day old 
pasturized milk from which the cream has been removed completely. 
Keep the culture at ordinary room temperatures of from 6o° to 68° or 

* See Biol. Bull. 63:442. 1932- 



62 Phylum Protozoa 

70 F. Place the culture near a window; diffused light is quite satis- 
factory. From three to six weeks are needed for the development of the 
culture. This time will depend on the density of the population desired. 
Ordinary half-pint milk bottles make satisfactory containers, and ab- 
sorbent cotton stoppers, which will admit air, may be used to keep out 
dust and spores of other forms of life. Good results have been obtained 
in culturing other species of Euglena in the same manner. 
. A pure culture of Euglena may be obtained by selecting under the 
microscope with a fine pipette [see p. 43] individual specimens of the 
desired species. A clean culture is very valuable and, once it is obtained, 
it may be perpetuated indefinitely if care is taken to prevent contamina- 
tion. Obviously, only sterilized pipettes should be used in removing 
specimens from the culture for class use. 

References 

For the culture of Euglena spp. see pp. 53, 63, and 71. 

For the culture of Euglena deses see p. 53- 

For the culture of Euglena anabaena see p. 53. 

A CULTURE MEDIUM FOR FREE-LIVING FLAGELLATES* 

THE following culture method, which has been tried out for two years, 
may be of use to other laboratories. Whole wheat is weighed into 
5 gm. lots, which are then put into large test tubes and 25 cc. of tap 
water added. These are then plugged with cotton, capped with lead 
foil, and autoclaved at 15 pounds' pressure for 2 hours, which very 
thoroughly macerates the wheat. Tap water is again added up to 50 cc, 
and desired percentages of this fluid are used after shaking. After open- 
ing a tube it is necessary to sterilize again in an Arnold sterilizer, as 
bacterial growth is vigorous in the mixture. However, a tube may be used 
day after day, if sterilized daily. 

Various percentages of this mixture afford a very good medium for 
many Protozoa. Bacterial feeders such as Chilodon, Paramecium, 
Oicomonas, and others thrive on it. Ochromonas, Chilomonas, and several 
of the smaller Euglenas (E. gracilis [see also pp. 53 and 82], E. quar- 
tana, and E. mutabilis) have been grown in abundance in various dilu- 
tions. There are several species of Amoeba which likewise occur or are 
capable of being cultured in large numbers. It has proved best, how- 
ever, for Entosiphon and Peranema. Both of these forms are easily 
grown in quantities sufficient for classroom use; isolation cultures of the 
former have been carried over a year on this medium. In general 
it seems much better than cracked boiled wheat, which is often used. 

♦Reprinted from Science 65: 261, 1927, by James B. Lackey, U. S. Public Health 
Service. 



Euglenidae 63 

CULTURE OF SOME FLAGELLATES AND CILIATES 
Paul Brandwein, Washington Square College, New York University 

Euglena. To 100 cc. of a modified Klebs' solution (Solution B)* in 
a white glass battery jar add 40 rice grains (boiled 5-10 minutes) and 
900 cc. of distilled water. The foregoing medium is allowed to stand for 
about five days. The jar is then placed in indirect sunlight (the direct 
rays of the sun should not strike this culture for more than an hour a 
day), and inoculated with Euglenae three times (10 cc. of a dense 
Euglena culture) at three day intervals. If an old Euglena culture is 
available the organisms may be found encysted on the sides of the vessel 
and it is of great advantage to inoculate the cysts along with the free 
Euglenae. Starting ten days after the initial inoculation, growth may be 
accelerated by adding (three times, at weekly intervals), 25 cc. of Solu- 
tion B and 10 mg. of the tryptophane powder. The further addition of 
5 grains of boiled rice each month will serve thereafter to maintain the 
culture. Large ciliates and rotifers are detrimental. 

Another technique, applicable to several Protozoa involves the use of 
an egg yolk-distilled water medium. 

Chilomonas. A thin smooth paste is prepared by grinding 0.5 gm. of 
the boiled yolk of a fresh hen's egg with a small amount of distilled water. 
This is added to 500 cc. of distilled water and the mixture, after standing 
two days, is inoculated with the original Chilomonas culture. If such a 
culture is not available, spontaneous inoculation will occur if the culture 
jar is left uncovered, since cysts of Chilomonas seem to be omnipresent. 

Paramecium, Colpidium, Colpoda, Euplotes. These ciliates have done 
well on the egg yolk medium when Chilomonas is provided as prey. Start 
a Chilomonas culture as previously directed and inoculate with 10 cc. 
of a culture of the desired ciliate, three times, on the 4th, 6th, and 8th 
day after starting. Dense maximum growth has usually been obtained 
in two weeks and subculturing has been necessary about every month. 

Didinium. The organism will thrive exceedingly well when introduced 
into one of the Paramecium cultures described above. As the Para- 
mecium diminishes in one culture a fresh one should be available for 
inoculation with the Didinium. In fact it is advisable to keep 
Paramecium cultures in a separate room ; otherwise it is difficult to avoid 
contamination with Didinium. 

Vorticella. A modification of the method found useful for Paramecium 
is necessary here. The medium of % gram of mashed hard-boiled egg 
yolk in 750 cc. of distilled water is permitted to stand for two days; it 
is then filtered through cotton, 100 cc. of the filtrate are added to a 

♦Modified Klebs' Solution (Solution B): KXO :J .25 gm., MgS0 4 .25 gm., KH 2 P0 4 
.25 gm., Ca(N0 3 ) 2 1 gm., Bacto-Tryptophane Broth (powder) .010 gm. (Digestive 
Ferments Co.), distilled water to make 1000 cc. 



64 Phylum Protozoa 

finger bowl and the Vorticellae are introduced; great numbers of the 
animals will be found clinging to the glass surface within two weeks. It is 
advisable to subculture every three weeks. 

Stentor. This organism may be cultured by two methods; namely, 
that found useful for Amoeba, and that described for Vorticella, except 
that certain modifications must be made here. The rice-agar with 50 cc. 
of Solution A [see footnote on p. 73] is permitted to stand for two days in 
a finger bowl ; at this point Chilomonas is added together with as many 
Stentors as possible. Or, the medium of y 2 gram of mashed hard-boiled 
egg yolk in 750 cc. of distilled water is permitted to stand for three days, 
filtered, and inoculated with Chilomonas and the Stentors. (About 5 cc. 
of a heavy Chilomonas culture are necessary for the inoculation in both 
cases.) It is desirable to subculture every month. 

Stylonychia and Oxytricha. For these hypotrichs and certain others 
the presence of Chilomonas seems advantageous. Before introducing the 
desired ciliate, allow some 30 cc. of Solution A in a rice-agar bowl, 
inoculated with 10 cc. of a Chilomonas culture [see p. 63] to stand 
about four days. Swarming cultures have usually been obtained within 
two weeks. 

References 

For a Phacus-like organism see p. 69. 

For the culture of Colacium vesiculosum see p. 53. 

For Euglenamorpha see p. 69. 
Family Astasiidae 

For the culture of Astasia sp. and A. ocellata see p. 54. 

For the culture of Peranema see pp. 62 and 136. 
Family Heteronemidae 

For the culture of Entosiphon see pp. 62 and 177. 

Order protomonadida, 
Family trypanosomatidae 

A SIMPLE METHOD FOR CULTURING TRYPANOSOMA 

LEWISI 

Reed O. Christenson, University of Minnesota 

IT is not always feasible, on short notice, to obtain living trypanosomes. 
Laboratory cultures should be established and maintained. This can 
be done easily by using the following method which we have found both 
simple and reliable. 

Wild rats are caught by setting a number of spring-traps near their 
holes or runways. The captives are killed and opened. A small amount 
of blood is taken from the heart, using clean pipettes, and this is dropped 



Polymastigida 65 

into cotton-stoppered vials half full of sterile 0.75% saline solution. The 
vials are taken to the laboratory, a drop of the fluid placed on a slide, 
covered, and examined. Under low power ( 16 mm. objective) the para- 
sites cannot be seen, but if they are present their movements cause violent 
agitation of the corpuscles, thus enabling a tentative diagnosis to be made. 
They are readily seen under high power (4 mm. objective) if the light 
is carefully regulated. About 5 to 10% of the wild rats carry infections. 

To establish the strain in laboratory animals the fluid from positive 
vials is drawn into a clean hypodermic syringe, and intraperitoneal inocu- 
lations are made into partly grown laboratory rats, about 0.5 cc. being 
given each one. Occasionally an animal will be found to be immune. 
This fact necessitates the infection of several so as not to lose the infec- 
tion. About seven days later a test is made. The tip of the tail is cut off 
and a drop of blood allowed to mix with a small drop of saline solution 
on a slide. It is then covered and examined microscopically. 

By the end of the second week the trypanosomes have usually reached 
their maximum number and have begun to decrease. By the fourth or 
fifth week (occasionally somewhat longer) they may have disappeared 
entirely from the blood. Before this time it is necessary to inoculate 
new, non-immune animals. An infected rat is etherized; the needle of a 
hypodermic syringe partly filled with saline solution is inserted into the 
heart; and some blood is drawn out. This is injected directly into the 
peritoneal cavity of the new animals. By repeating the transfer of in- 
fective serum at the proper times the strain may be maintained for in- 
definite periods. 

Occasionally it is desired to keep living trypanosomes for a few days in 
the serum-saline mixture. This may be accomplished by placing the 
container in a refrigerator (about 18 C.) where they will live for a week 
or longer. 

References 

For the culture of Oicomonas see p. 62. 
Family Bodonidae 

For the culture of Embadomonas see p. 88. 

Order polymastigida 

NOTES ON CULTURING CERTAIN PROTOZOA AND 
A SPIROCHAETE FOUND IN MAN 

M. J. Hogue, University of Pennsylvania Medical School 

MEDIA USED 

Locke-Egg medium. One hen's egg is thoroughly shaken with glass 
beads in a flask. Add 200 cc. of Locke solution (sodium chloride 0.9 gm., 



66 Phylum Protozoa 

calcium chloride 0.024 gm., potassium chloride 0.042 gm., sodium bi- 
carbonate 0.02 gm., dextrose 0.25 gm.). Heat over a hot water bath for 
15 minutes keeping the medium in - constant motion by revolving the 
flask. Filter through cotton with a suction pump. Put about 6 cc. of 
the filtrate into each test tube. Autoclave the tubes for 20 minutes under 
15 pounds' pressure (Hogue, 1921a). 

Ovomucoid medium. The white of one hen's egg is thoroughly shaken 
with glass beads in a glass flask. Add 100 cc. of 0.7% sodium chloride 
solution. Cook this for half an hour over a hot water bath, keeping the 
contents of the flask in constant motion by revolving the flask. Filter 
through cotton, using a suction pump. Put about 6 cc. of the filtrate in 
each test tube. Sometimes one loopful of the egg yolk is added to each 
test tube. Autoclave the tubes for 20 minutes under 15 pounds' pressure 
(Hogue, 1921a). 

Sodium chloride sheep serum water. To a flask containing 100 cc. 
of 0.85% sodium chloride solution which has been sterilized in the auto- 
clave for 15 minutes at 15 pounds' pressure add 10-15 cc. of sterile sheep 
serum water. This is prepared by diluting one part of sheep serum with 
three parts of distilled water and sterilizing in an Arnold steam sterilizer 
for one hour at ioo° C. for three successive days. Pour the sterile sodium 
chloride sheep serum water into sterile test tubes. It will have a pH 
of 7-7.4 (Hogue, 1922a). 

Sodium chloride pig serum water. This is made in the same way as 
sheep serum water except that one part of pig serum is diluted with four 
parts of distilled water. It will have a pH of from 6.8-7.4 (Hogue, 
1922a). 

Sodium chloride sheep serum water modified. This modification is 
used for Trichomonas buccalis. Take equal amounts of 0.85% sterile 
sodium chloride solution and sterile sheep serum water. Put 5 cc. in a 
test tube. This has a pH of 7.7. Just before using, add a small amount 
of saliva taken from a person who is not infected with T. buccalis (Hogue, 
1926). 

Deep cultures. "Deep cultures" are used in order to keep organisms 
for a long time without transferring them (Hogue, 1922a, 1922b, 1926, 
1933). These are made by putting 15 cc. of the culture to be used into 
150 mm. test tubes, inoculating them at the bottom of the tube with a 
long pipette and then covering the medium with a layer of sterile paraffin 
oil. This prevents evaporation but does not make the cultures anaerobic. 
Under these conditions the animals do not divide so rapidly and do not 
form such large quantities of waste products. In these "deep cultures" 
they live for weeks or months, depending on the species. 



Polymastigida 67 

SPECIES CULTURED 

Retortamonas intestinalis (Waskia or Embadomonas) grew well on 
both Locke-egg and ovomucoid, forming cysts in each medium. On the 
Locke-egg it lived from 3-10 days. One culture lived 17 days on the 
ovomucoid. The cultures were grown at 35 C. (Hogue, 1921a). 

A new variety of Retortamonas found in man grows well on sodium 
chloride sheep serum water. It has been kept at room temperature (20- 
33 C.) for over 3 years and has survived one heat wave of 39. 5 C. 
It is transferred every 7 days. "Deep cultures" of it are kept at room 
temperature. In them the organisms live from 8-9 months. One culture 
lived over 12 months (Hogue, 1933). 

Trichomonas buccalis grew on a sodium chloride sheep serum medium 
especially rich in sheep serum and containing a small amount of human 
saliva. On this medium they were cultured for over 7 months. Transfers 
were made every 2-3 days, though they lived about 5 days on this 
medium at 30 C. In "deep cultures" they lived from 30-60 days 
(Hogue, 1926). 

Trichomonas hominis grew well on Locke-egg, ovomucoid and sodium 
chloride sheep serum water. In the early work the first two media were 
used and the cultures were incubated at 35 C. They lived about 6 days 
on the Locke-egg and from 6-10 days on the ovomucoid (Hogue, 1921b). 
In later work sodium chloride sheep serum water has been used. They 
have been incubated at 36 C. and at room temperature. In "deep 
cultures" these organisms live from 35-66 days at 35 C. The most 
favorable pH for rapid multiplication is pH 7.2-8.4 with pH 8 as an 
optimum. If it is desirable to keep the cultures longer pH 7.2-7.4 is 
better. Here they divide more slowly (Hogue, 1922a). 

Spirochacta eurygyrata grew well on Locke-egg, ovomucoid, and 
sodium chloride pig serum water. On ovomucoid and Locke-egg they 
lived from 8-12 days. The best results were obtained with sodium 
chloride pig serum water with pH 7. These were incubated at 35 ° C. 
In "deep cultures" of sodium chloride pig serum w T ater they lived from 
60-90 days. One culture contained active spirochaetes 127 days after 
inoculation (Hogue, 1922b). 

References 
For the culture of Trichomonas see also pp. 69, 88, and 91. 
For the culture of Tritrichomonas see p. 118. 
For the culture of Enteromonas see p. 88. 
For the culture of Monocercomonas see p. 129. 
For the culture of Chilomastix see p. 88. 

Bibliography 
Hogue, M. J. 1921a. Waskia intestinalis : Its cultivation and cyst formation. 
J. A. M. A. 77:112. 



68 Phylum Protozoa 

1921b. The cultivation of Trichomonas hominis. Amer. J. Trop. Med. 

1:211. 

1922a. A study of Trichomonas hominis, its cultivation, its inoculation into 



animals, and its staining reaction to vital dyes. Johns Hopkins Hosp. Bull. 

33:437- 

■ 1922b. Spirochaeta eurygyrata. J. Exper. Med. 36:617. 

1926. Studies on Trichomonas buccalis. Amer. J. T-rop. Med. 6:75. 

1933. A new variety of Retortamonas (Embadomonas) intestinalis from 



man. Amer. J. Hyg. 18:433. 

FROG AND TOAD TADPOLES AS SOURCES OF 
INTESTINAL PROTOZOA FOR TEACHING PURPOSES* 

R. W. Hegner, School 0) Hygiene and Public Health 

MANY teachers of protozoology and invertebrate zoology use frogs 
for the purpose of obtaining intestinal Protozoa for class use, 
but it does not seem to be generally known that the tadpoles of frogs 
and toads are even more valuable than the adults as sources of material. 
Unfortunately tadpoles are most abundant late in the spring and in 
early summer when classes are usually not in session, but two species 
of frogs that are more or less common throughout the United States pass 
two or more seasons in the tadpole stage and hence are available in 
the autumn and, in the southern part of the country, at any time of the 
year; these are the green frog, Rana clamitans, and the bullfrog, R. 
catesbiana. A breeding place once found will serve as a source of supply 
year after year. 

Sample tadpoles should be collected some time before the class meets so 
as to determine the incidence of infection and numbers present of the 
various species of Protozoa, since these vary from year to year. The 
specimens for class use may be collected several days before they are 
needed but should not be kept more than a week or two since they tend 
to lose their infections under laboratory conditions. The writer has 
found dishes about 10 inches in diameter and 3 inches deep containing 
a quart of tap water to be suitable for about 20 tadpoles each. The 
dishes should not be covered with glass plates, but the water should be 
changed every day or two. Tadpoles may be killed very quickly, as 
adult frogs usually are, by destroying the brain and spinal cord with a 
heavy needle. The ventral body wall may then be opened from the 
anterior to the posterior end. The intestine is coiled within the body 
cavity, being several hundred millimeters in length. The rectum, or 
posterior portion of the alimentary tract, is tightly coiled and is separated 
from the intestine by a constriction. The different species of intestinal 
Protozoa are rather definitely distributed within the intestine and rectum. 

* Reprinted from Science 56:439, 1922, with slight changes by the author. 



Amoebidae 69 

The anterior portion of the intestine is inhabited by a flagellate, Giardia 
agilis [see also p. 89] ; in various parts of the intestine and rectum 
Endamoeba ranarum may be found; the rectum is the principal habitat 
of three genera of flagellates, Trichomonas [see p. 67], Hexamitis, and 
Euglenamorpha, and of several green flagellates resembling members of 
the genera Euglena and Phacus. To study any of these in the living 
condition, the part of the digestive tract containing them should be 
teased out in a drop of 0.7% salt solution and covered with a cover glass. 
Any of the species mentioned may be found with low magnification, such 
as obtained with a 16 mm. objective and a number 5 ocular. 

Nyctotherns cordijormis is a very large ciliate that is often found in 
the rectum of tadpoles. It appears to be a scavenger and resembles 
Paramecium in structure and in its primary life processes. Opalina 
ranarum is also a large ciliate. It and other species of Opalina are 
frequent inhabitants of the rectum of tadpoles. 



Class Sarcodina, Order amoebae a, Family 

AMOEBIDAE 

PROTOZOAN CULTURES* 

George R. La Rue, University of Michigan 

NATURAL POND CULTURES 

THESE cultures should be made by the methods of Hyman, Jennings, 
and others. The plant material collected should not be restricted to 
Ceratophyllum and Elodea, but should include any vegetable matter, 
e.g., old lily pads and stems, cat-tails (especially if decay has com- 
menced), decaying leaves of trees, grass and sedges, etc., from pools, 
ponds, marshes, bogs, ditches, and rivers. Chara and clean Spirogyra 
and the brown mat of algae from the surface of a dam or of stones are 
of value, if not too much is taken and if to this is added hay or other 
materials to furnish food for bacteria. Decaying Sphagnum from 
sphagnum bogs is valuable. Mud or ooze from the bottom of ponds or 
pools which get the drainage from pastures or barnyards is also good. 
In collecting this vegetable material always bring some of the water 
from the same situation. Most Protozoa are sensitive to changes in 
water. Treated water is particularly bad for some species and should 
never be used until it has stood for some days in an open vessel or tank. 

* The beginner would do well to read the article by Hyman {Trans. Micr. Soc. 
44:216, 1925) to secure some ideas concerning general methods. He should also read 
other articles, some of which are listed at the end of these directions. 



70 Phylum Protozoa 

Place natural pond cultures in shallow glass or earthenware dishes 
and keep covered to prevent evaporation. If pond scums and Euglena 
are wanted or if these are needed for the food of any desired protozoan, 
set the cultures in diffuse light, never in direct sunlight. Label each cul- 
ture. Use one pipette for each culture. This precaution is of great 
importance in examining the subcultures. 

Examination of cultures. At intervals each culture should be ex- 
amined carefully. In making the examination, take samples of scum, of 
clear liquid, scrapings from sides of vessel, from the light side and from 
the dark side, from the vegetation, and from the bottom. These situa- 
tions may furnish different forms because some forms swim freely while 
others creep. Some require much oxygen while others can exist on less. 

Subcultures. If at any time any culture is yielding a large number of 
desirable species such as Paramecium, Euglena, Amoeba, etc., sub- 
cultures should be made as follows: Clean thoroughly short stender 
dishes or finger bowls. Place in these filtered cistern or distilled water. 
Do not use raw tap water. If neither cistern nor distilled water is 
available, filter some water which has stood for a long time in an aqua- 
rium or tank. Such water seems to have lost many of its noxious prop- 
erties. Place a few straws of clean hay in the water and allow it to stand 
36 to 48 hours before the desired Protozoa are placed in it. In place of 
dry hay, the hay may be boiled in distilled or cistern water and the hay, 
not the hay water, added to the water in the dish. Boiling the hay will 
sterilize it so that sterilization of the hay in an autoclave is probably 
unnecessary. Allow to stand before inoculating. In such cultures bac- 
teria will develop and on these Paramecium and certain other forms feed. 

PURE CULTURES 

Isolation of particular species is not an entirely simple matter. How- 
ever, individual Protozoa may frequently be secured by using a mouth 
pipette. This is a glass tube with a finely drawn tip in one end of a 
rubber tube 12 to 15 inches long with a short piece of glass tubing to 
place in the mouth in the other end. Place a small dish of the culture 
containing the organism sought on the stage of the binocular microscope 
and put the glass tube in the mouth. When a desired individual is found 
bring the point of the pipette near it and suck on the tube. With some 
practice the protozoan may be caught and may be put on a slide in a 
small drop of water and examined. In this way individual Protozoa 
may be isolated for the inoculation of cultures and by washing them in 
one or several changes of water the worker can be reasonably sure that 
other species are not present. ^^J 

Amoeba. In culturing Amoeba tap water may be harmful and must 



Amoebidae 71 

not be used. Distilled water is very good. Boil a little hay and add the 
hay (not water) to distilled water in several short stender dishes (2% 
to 4 in. diam.), or finger bowls. Inoculate the cultures with Euglena, 
diatoms, and other algae, small colorless flagellates, or some small ciliates 
like Colpidium [see p. 107]. When these cultures are going well 
inoculate with material containing plenty of good-sized Amoebae. If the 
food organism is an alga the Amoeba culture must have diffuse light. 
If ciliates are the food organism darkness will be suitable but not neces- 
sary. When such cultures are once established add dry or boiled hay 
at intervals of a few days to a few weeks and add distilled water to 
make up for evaporation. Reinoculate cultures if they do not show 
numbers of good Amoebae. 

The following method of culturing large Amoebae has proved very 
successful. Thoroughly wash and rinse finger bowls and fill % full of 
distilled water. Add 6 or 8 grains of rolled wheat, rolled oats, or rice. 
Rice is best. Label, cover, and mark level of water on label. Inoculate 
at once with Amoebae from a good culture, taking material from the 
bottom of the dish and examining it to ascertain that Amoebae are 
actually being taken. At intervals of a week or two fill cultures up to 
mark, using distilled water. After culture is well established add a few 
kernels of rice or flakes of wheat or oats occasionally. Removal of a 
part of the water from time to time and the addition of fresh distilled 
water stimulates reproduction. 

Euglena. Euglena thrives best in water having considerable organic 
material. For this reason good cultures may be made in manure solu- 
tions. Horse or cow manure is boiled in spring water or distilled water. 
These solutions should be made and allowed to stand 36 to 48 hours 
and then inoculated with Euglena. Old hay cultures if left in diffuse 
light almost invariably end in being almost pure cultures of Euglena. 
In collecting Euglena in nature seek it in barnyard pools, or pasture 
pools which receive considerable organic material. It may often be 
found among the algae of ponds even though no green scum is found on 
the surface. Any green scum on the surface of a pond, or green slime 
on decaying vegetation in ponds or streams, is almost certain to have 
large numbers of euglenoid forms in it. 

Besides hay and manure solutions, rice in water yields good cultures. 
In using rice, boil 7 or 8 grains in a pint of distilled water or old water 
from a tank, put in a broad dish and allow to stand until a bacterial 
scum has formed. Then inoculate. Euglena cultures come on slowly, 
and must be started 4 to 8 weeks before they are needed for study. In 
old cultures Euglenae usually encyst on the surface of the dish, and they 
may be kept many months in the encysted condition. Covered cultures 
may be good for months and since they may be revived after encystment 



72 Phylum Protozoa 

takes place by adding fresh manure solution, they serve as admirable 
sources of material for inoculating new cultures. 

Paramecium. Hay cultures are perhaps the most common type but 
Paramecium will thrive in almost any medium which does not become 
too sour and which will grow plenty of bacteria. Cracked wheat has 
been used; also rice, rolled wheat or oats, bread, and malted milk. The 
latter makes a good bacterial culture; such cultures do not last long 
but they are easily inoculated into new cultures. If wheat is used, add 
boiled wheat to distilled water at the rate of 30 or 40 kernels to a gallon 
of water. Rolled wheat or oats should be used raw, and at the begin- 
ning not more than 10 flakes to a pint of water. Bread should be used 
sparingly. 

Bibliography 

Dawson, J. A. 1928. The culture of large free-living Amoebae. Amer. Nat. 62:453. 
Hyman, L. 1925. Methods of securing and cultivating Protozoa. 

I. General statements and methods. Trans. Amer. Micr. Soc. 44:216. 
— 1931. Methods of securing and cultivating Protozoa. 

II. Paramecium and other ciliates. Ibid. 50:50. 

Jennings, H. S. 1903. Methods of cultivating Amoeba and other Protozoa for 

class use. /. A ppl. Micr. and Lab. Methods 6:2406. 
Kofoid, Charles A. 1915. A reliable method for obtaining Amoeba for class 

use. Trans. Amer. Micr. Soc. 34:271. 
La Rue, G. R. 1916. Notes on the collection and rearing of Volvox. Ibid. 35:151. 
1917- Notes on the culturing of microscopic organisms for the zoological 

laboratory. Ibid. 36:163. 

191 7. Further notes on the rearing of Volvox. Ibid. 36:271. 



Smith, B. G. 1907. Volvox for laboratory use. Amer. Nat. 41:31. 

Turtox Service Leaflet No. 4. The care of Protozoan cultures in the laboratory. 
General Biological Supply Co. 

Turtox Protozoa Booklet. General Biological Supply Co. 

Turtox Biology Catalog and Teachers' Manual. General Biological Sup- 
ply Co. 

Welch, M. W., 1917. The Growth of Amoeba on a solid medium for class use. 
Trans. Amer. Micr. Soc. 36:21. 



CULTURE OF SOME FRESHWATER RHIZOPODA 

Paul Brandwein, Washington Square College, New York University 

Amoeba* The following method has been notable in giving a larger 
proportion of successful cultures which achieve a very dense maximum 
growth in 3-4 weeks and do not require subculturing for 8-10 weeks. It., 

♦Chalkley. H. W., 1930, Science 71:441 and Pace, D. M., 1933, Arch. f. Protist. 
79:133 have previously reported two methods for Amoeba. Our medium differs little 
from Chalkley's, but the method in its entirety has given better results. 



Amoebidae 73 

differs from previous methods chiefly in the use of agar* and the slight 
modification of the salt content of the culture solution. 

Prepare finger bowls by covering the bottom with a thin (1-2 mm.) 
sheet of agar. This is done by pouring a warm, filtered, aqueous 0.75% 
solution of powdered agar into each bowl. While the agar is still soft 
imbed 5 rice grains, evenly spaced. The finger bowls and pipettes are 
previously washed thoroughly in hot water, and the rice heated (10 
minutes) in a dry test tube immersed in boiling water — two necessary 
precautions against contaminating organisms. 

About 50 Amoebae, together with 10 cc. of the medium in which 
they have previously been growing, are introduced into each bowl and 
then 30 cc. of the general culture solution (Solution A) f are added. 
Thereafter, every three days, 20 cc. of solution A are added to each 
bowl until the total volume is 80-90 cc. 

When maximum growth has been attained and a culture shows signs 
of waning, it may be replenished by adding 10 cc. of solution A and 1 
rice grain (preheated). 

In a day or two after starting a culture the agar layer becomes de- 
tached from the bottom of the vessel and the Amoebae grow in layers on 
its upper and lower surfaces and also on the glass surface. 

In about 2 months, it is advisable to subculture by dividing the con- 
tents of each bowl, exclusive of the rice-agar, into four parts, pouring 
each into a freshly prepared finger bowl, and adding an equal quantity 
of solution A. From here the procedure is the same as before. 

If the original source of Amoebae is limited, as is the case when they 
are collected from the fieldj, it is necessary to modify the method 
slightly, by starting the cultures in Syracuse dishes instead of finger 
bowls. This apparently gives a better initial concentration of the 
Amoebae and makes the change of culture conditions less abrupt. 

The Syracuse dishes are prepared with an agar film in which 2 rice 
grains are imbedded. Introduce the available Amoebae with 4 cc. of 
the water in which they were collected and 4 cc. of solution A. In suc- 
cessful cases there will be a rapid proliferation and when 200-300 animals 

*The use of an agar layer on the bottom of the dishes was originally suggested by 
Dr. R. Chambers for the purpose of anchoring the rice grains, about which the Amoebae 
tend to congregate. Mr. M. Sheib, working for Dr. Chambers, has been very successful 
with this method. The writer came upon it independently, and is of the opinion that the 
augmented growth is due to the increased surface available to the Amoebae for securing 
their prey, although some component, added via the agar, may be involved. 

fGeneral culture solution (Solution A) — NaCl 1.20 gms., KC1 0.03 gms., CaCU 
0.04 gms., NaHC0 3 0.02 gms.; phosphate buffer solution having a pH 6.9-7.0, 50 cc; 
distilled water to 1000 cc. For use dilute this 1:10. This solution maintains a fairly 
constant pH of about 7.0 and serves well not only for Amoeba, but also for general use. 

Jin ponds from beds of Vaucheria, Hydrodictyon, from the under side of Castalia, 
Lemna, and Spirodela leaves. 



74 Phylum Protozoa 

are present, the culture (minus the rice-agar) is added to a rice-agar 
bowl with 30 cc. of culture solution A. The steps from here are the 
same as before. 

The cultures were maintained at 19-2 2 ° C. by stacking the finger 
bowls in a sheet metal container, placed in a sink through which tap 
water was kept circulating to the level of the highest finger bowl. Even 
better growth has been obtained at 17-19 C, but it was easier to main- 
tain the former temperature. 

Arcella. The foregoing technique has also been very successful for 
this organism. In this case, however, temperature control is not neces- 
sary. 

It is detrimental to have Stentor, Paramecium, large hypotrichs, 
Philodina, or Stenostomum in cultures of Amoeba or Arcella. A culture 
in which these organisms have gained ascendance should be discarded 
and precautions should be taken against contamination of other cultures. 
Chilomonas and Colpidium while not detrimental in moderate popula- 
tions, should not be allowed to proliferate to the point where a culture 
becomes cloudy with them. Mold usually grows on and about the rice 
and does not seem detrimental although at times it is annoying since it 
is hard to disentangle the Amoebae from the mycelium. 

Actinosphaerium. For this organism best success has been obtained 
by insuring the presence of Paremecium, Stenostomum, or Philodina, 
which forms, apparently, serve as prey. To 30 cc. of solution A in a 
rice-agar bowl add 30 cc. of a dense culture of either of these animals 
and inoculate with 5-10 Actinosphaeriae. Maintain in diffuse light and 
replenish with more of the above mentioned organisms as required. 
Prolific cultures have usually been obtained in about two weeks. 

AMOEBA 

William LeRay and Norma Ford, University of Toronto 

THE fact is not generally appreciated that Amoebae are found in 
clean water, not foul. Two forms of Amoebae {A. proteus type) are 
used in our department. One of these, an unusually large and active 
form, possessing many pseudopodia, is found in association with catfish 
and newts (Ameiurus nebulosus and Triturus viridescens). Another 
with fewer pseudopodia and more regular in outline is taken with the 
Brook Stickleback (Eucalia inconstans). The latter form we find the 
more satisfactory for class work. 

The Brook Sticklebacks are collected in the backwash pools of rivers 
or streams. Two or three of these small fish are placed in a bowl con- 
taining 2000 cc. of pond water and fed upon Daphnia or enchytraeid 
worms. Gradually ooze forms on the bottom of the bowl and in about 
two weeks this ooze will be found to contain Amoebae. 



Amoebidae 75 

A culture of Amoebae is then set up in bowls containing 600 cc. of 
pond water. The bottom of each bowl is sprinkled with exceedingly 
fine sand which has been carefully washed and sifted through bolting 
cloth. The individual grains of sand should be scarcely larger, and for 
the most part smaller, than a single Amoeba. To each bowl is added 6 
grains of boiled wheat. (The development of the culture may be 
hastened by using boiled brown rice in place of the wheat, although the 
latter culture does not last so long.) Some ooze from the fish bowls is 
now introduced and the culture kept at about 73 ° F. 

In order that the Amoebae may thrive, fungus must grow on the grains 
of wheat. If this fails to appear, it may be obtained by adding a bit of 
dead worm or dead fly from some other culture. 

Within twelve days there should be an abundance of the Amoebae. 
Large numbers may be present as early as six or seven days, and usually 
so in eight days. A culture lasts about a month, but new ones should 
be started while the Amoebae are still active and healthy. Old cultures 
should be made over completely. 

The fine sand in the culture is most helpful in picking the Amoebae 
out of the bowl for microscopic study, since a little sand taken into a 
pipette will have the organisms attached to it. The sand also forms an 
excellent support for the coverslip, allowing the Amoebae to move about 
freely. As part of the habitat of the culture the grains of sand afford a 
cover under which the organisms can hide. In an undisturbed culture 
the Amoebae will be found contracted and resting either under the grains 
or close beside them. If disturbed by jarring or by a beam of light the 
Amoebae glide or "walk" away from this protecting cover. 

References 
For the culture of Amoeba see also pp. 62, 134, 136, and 177. 

STOCK CULTURES OF AMOEBA PROTEUS* 

IN the course of experiments it has been necessary to maintain cultures 
of Amoeba proteus in stock. The writer endeavored to find a medium 
that made requisite a minimum amount of attention. The effort in this 
direction met with considerable success. In view of the wide use of 
Amoeba of the proteus type in biological research and elementary in- 
struction in biology, a culture medium that is simple, reproducible, and 
extremely reliable will be of general interest. The medium used is as 
follows: 

NaCl 0.1 gr. 

KC1 0.004 gr. 

CaClo °-°o6 gr. 

H2O 1000 cc. (glass distilled) 

♦Reprinted with slight changes from an article in Science 71:442, 1930, by H. W. 
Chalkley, U. S. Public Health Service. 



76 Phylum Protozoa 

Two hundred to 250 cc. of this solution is put into a finger bowl or 
glass crystallizing dish 8 or 10 cm. in diameter and to each of such dishes 
is added 4 or 5 grains of polished rice (any brand carried at the corner 
grocery is suitable) . The cultures thus prepared are immediately seeded 
with 50 to 100 Amoebae, covered with glass plates to prevent evaporation 
and entry of dust, and then left, preferably in a dark cool place, to 
develop. Such cultures will produce a fine crop in from 2 to 4 weeks and 
so far in some 30 to 40 cultures the writer has had only one or two 
failures. Out of five cultures that were set up as a test, three cultures 
one year old had ample numbers of Amoebae; the other two died out in 
eleven months. 

These five cultures during their existence were deliberately neglected. 
No detritus was removed. Rice was added only when it was discovered 
that none was apparent in the culture. Water too was added to compen- 
sate for evaporation with no attempt at regularity, say, on the average of 
once a month. The temperature variation was from 19° to 2 8° C. 

In other words, the cultures were subjected to as careless handling as 
if in the hands of a somewhat below par student assistant, but they sur- 
vived. 

M. E. D. 
References 
For the culture of Amoeba proteus see also p. 59. 

THE CULTURE OF 
AMOEBA PROTEUS LEIDY PARTIM SCHAEFFER 

D. L. Hopkins, Duke University and D. M. Pace, Johns Hopkins University 

THE Amoebae used in the development of the following culture 
methods correspond to those designated by Schaeffer as Amoeba 
proteus Pallas (Leidy) in 1916 and as Chaos diffluens Mueller in 1926. 
We shall follow Mast and Johnson (1931) and retain the generic name, 
Amoeba, but shall follow Schaeffer (19 16) in separating Amoeba proteus 
(Leidy) into two species, Amoeba proteus Leidy partim Schaeffer and 
Amoeba dubia Schaeffer. While our experience has been limited to 
Amoeba proteus, it is probable that the culture methods described would 
also be satisfactory for Amoeba dubia. 

Collection and culture. Amoeba proteus is not found in nature as 
frequently as are smaller Amoebae. When found it will usually be in 
ponds or pools where there is neither too great nor too little organic 
material, where there are no excessively swift currents, and in water that? 
is not too alkaline. If, with a spoon or dipper, surface water containing 
a considerable amount of organic material is collected, taken to the 
laboratory, poured into tall slender jars and allowed to settle it will be 



Amoebidae 77 

found that the heavy organic material will settle in a thick layer on the 
bottom, and that the Amoebae will settle on top of this layer. They 
then may be removed with a pipette to a shallow dish and examined. 

Amoeba proteus collected in this way may be cultured simply by 
placing a suitable spring or pond water in finger bowls or other shallow 
dishes to depth of about 2 cm., adding 3 to 4 grains of wheat, or 5 to 6 
grains of polished rice, or 5 to 6 one-inch stems of timothy hay, and 
then inoculating with Amoebae from the collection. The Amoebae in 
these cultures will become very abundant in from 3 to 4 weeks. It 
should be noted here that Dawson (1928) gives an elaborate method 
for obtaining cultures from freshly collected material consisting of a 
gradual dilution of the collected water with distilled water. This process 
of dilution in our experience has not been necessary. Cultures obtained 
from freshly collected material are likely to contain in addition to 
Amoebae various organisms such as small Crustacea, rotifers, and va- 
rious Protozoa. Since none of these organisms, except the cryptomonad, 
Chilomonas, are necessary as food, and since others such as the Crustacea 
probably feed on the Amoebae ; it is well to take steps to eliminate these 
unnecessary organisms. A. proteus feeds on a variety of organisms, but 
Chilomonas alone is entirely adequate. 

Freeing Amoeba cultures oj contaminating organisms and establish- 
ing clone cultures. Sterilize some spring water by autodaving for 15 
minutes at 15 pounds' pressure, or merely by bringing to a boil. Allow 
to cool. Place sterile spring water in three or four sterile, chemically 
clean Syracuse watch glasses, and a finger bowl. The water in the 
finger bowl should have a depth of about 2 cm. Add to the finger bowl, 
3 to 4 grains of wheat, 5 to 6 grains of rice, or 5 to 6 pieces of timothy 
hay stems about 1 inch long. The wheat, rice, or hay should be auto- 
claved dry 15 minutes at 15 pounds' pressure, or placed dry in a test 
tube, then the test tube placed in a beaker of boiling water for about 15 
minutes. Now with a sterile capillary pipette and under a binocular 
microscope, select active Amoebae, pass them one at a time through the 
dishes of sterile spring water. If it is desired to obtain a clone culture 
(culture containing Amoebae all of which have descended from a single 
parent) put only one washed Amoeba into the finger bowl culture. To 
obtain merely a clean culture it is advisable to put into the finger bowl 
culture 25 or more Amoebae. Now add to the culture in the finger bowl 
containing washed Amoebae a drop of culture fluid containing only 
Chilomonas [see pp. 59 and 63] and bacteria. The best way to do 
this is to take small drops of the old culture fluid, examine them carefully 
under high power selecting only those drops which show only Chilomonas 
and bacteria to be present, rejecting all others. It is best to place these 
drops each on a small sterile coverslip, and then if the drop is found 



78 Phylum Protozoa 

satisfactory merely drop the coverslip into the culture bowl. Now cover 
the cultures carefully with a glass cover or stack the finger bowls. In 
either case be sure that the covers are sterile and chemically clean. It 
is always wise to set up several cultures to insure against mishap. If 
successful the cultures will become more or less abundant in from four to 
six weeks. To subculture proceed as before except that it is not now 
necessary to wash the Amoebae, unless observation has shown that they 
have become contaminated with undesirable organisms. 

The salt content of culture media. While almost any uncontaminated 
freshwater will support growth it was found by Pace (1933) during a 
detailed study of the relation of salts to growth and reproduction that 
a more certain way of obtaining the proper salt concentration for abun- 
dant growth and reproduction is by using a synthetic spring water of a 
definite composition and concentration determined by experiment to be 
optimum for growth and reproduction. The following solutions are 
recommended: 

(1) (2) 

NaoSiC>3 15 mg 100 mg 

NaCl 12 mg 12 mg 

Na 2 S0 4 6 mg 6 mg 

CaClo 6.5 mg 6.5 mg 

MgClo 3.5 mg 3-5 mg 

FeCl3 4 mg 4 mg 

Dist. Water 1000 cc 1000 cc. 

Sufficient HC1 to give a pH of 7.0 to 6.8. 

We shall designate (1) as "dilute artificial spring water" and (2) as 
"concentrated artificial spring water." The difference between the (1) 
and (2) solutions is in the concentration of sodium silicate. The first 
solution is recommended when attempting to culture Amoebae recently 
collected, since the concentration is nearer that of most natural fresh- 
waters than is the second. However, when Amoebae have been cultured 
in a dilute medium they may be transferred to the more concentrated 
solution and more rapid growth and reproduction will be obtained. In 
fact the second solution is a solution in which optimum growth and 
reproduction was found to take place. A concentration between those of 
these two solutions in which the sodium silicate is 25 mg. per liter in- 
stead of 15 mg. or 100 mg. per liter, results in a much slower rate of 
reproduction. No serious difficulty will arise if the total salt concentra- 
tion of (1) is a little less or that of (2) is a little greater, but the con^, 
centration of (1) must not be a little greater, or that (2) a little less 
than that indicated. Reproduction is much better when sodium silicate 
is present than when it is replaced by some other salt. Chalkley (1930) 
and Hahnert (1932) give salt mixtures which allow good growth and 



Amoebidae 79 

reproduction, but the "concentrated artificial spring water" just given 
has proven superior in our experience. 

Hydrogen-ion concentration oj cultures. Amoeba proteus will culture 
successfully at widely varying hydrogen-ion concentrations. Good cul- 
tures have been obtained at pH values anywhere between 4.0 and 8.5. 
A pH value of 6.6 to 6.8 is generally considered to be optimum. The 
optimum pH value, however, is probably dependent upon the salt con- 
tent of the culture. Therefore for ordinary purposes it is unnecessary 
to take any steps to control the hydrogen-ion concentration of the cul- 
tures. The great majority of cultures set up as described above will 
come to an equilibrium at a favorable hydrogen-ion concentration. 

The maintenance of a constant hydrogen-ion concentration for special 
purposes is rather difficult. The addition of enough buffer salts to main- 
tain a constant pH value brings the total salt concentration to a value 
too high for growth of the Amoebae. However, Hopkins and Johnson 
(1928) by a gradual addition of buffer salts were able to adapt the 
Amoebae to the increased salt concentration so that a constant pH value 
was maintained, and at the same time Amoebae grew and reproduced. 
This procedure should be as follows: To a new culture in which Amoebae 
have begun to grow and reproduce add each day about 0.5 cc. of Clark 
and Lubb's phosphate buffer of the desired pH value for each 100 cc. 
of culture fluid until a total of 5 cc. of the buffer has been added for 
each 100 cc. of culture fluid. 

Hopkins (1926) used a feeding method for maintaining a constant 
hydrogen-ion concentration. Amoeba cultures when first set up as 
described above first become more acid in reaction and then gradually 
return to the alkaline side of neutrality. If, when in the return of a 
culture to alkalinity a given pH value is reached which is desired to be 
maintained as a constant value, one adds a certain amount of fresh 
sterilized hay or wheat infusion daily, the acid tendency of the fresh 
infusion will oppose the alkaline tendency of the culture. By measuring 
the hydrogen-ion concentration daily and adding fresh infusion accord- 
ingly it is possible to maintain the concentration within a range of 0.2 pH 
units. 

Temperature. As long as the temperature in the culture is below 25 
C. they remain in good condition. If the temperature is too low 
growth is retarded. It is necessary to freeze them before they are 
seriously injured. About 22 ° C. is perhaps optimum for growth. In 
summer when room temperature goes above 25 C. it is advisable to keep 
cultures in a cool basement room or, better, in a cold room where the 
temperature is maintained at the desired level. 

Light. Direct sunlight is injurious. Growth is just as good in ab- 
solute darkness as in any light intensity. 



80 Phylum Protozoa 

Using methods described above the senior author has kept Amoeba 
proteus in continuous culture since 1923, a total of 12 years, and the 
Amoebae are still in excellent condition. 

Bibliography 

Chalkley, H. W. 1930. Stock cultures of Amoeba. Science 71:442. 

Dawson, J. A. 1928. The culture of large free living Amoebae. Amer. Nat. 62: 

453 

Hahnert, F. H. 1932. Studies on the chemical needs of Amoeba proteus: A cul- 
ture method. Biol. Bull. 62:205. 

Hopkins, D. L. 1926. The effect of certain physical and chemical factors on loco- 
motion and other life processes in Amoeba proteus. J. Morph. and Physiol. 

45:97- 
Hopkins, D. L., and Johnson, P. L. 1928. The culture of Amoeba proteus in a 

known salt solution. Biol. Bull. 56:68. 
Mast, S. O., and Johnson, P. L. 193 1. Concerning the scientific name of the 

common large Amoeba, usually designated as Amoeba proteus (Leidy). Arch. /. 

Protist. 75:14. 
Pace, D. M. 1933. The relation of inorganic salts to growth and reproduction in 

Amoeba proteus. Ibid. 79:133. 
Schaeffer, A. A. 1916. Notes on the specific and other characters of Amoeba 

proteus Pallas (Leidy). A. discoides spec. nov. and A. dubia. spec. nov. Ibid. 

37:204. 
1926. Taxonomy of the Amebas with description of thirty-nine new marine 

and fresh water species. Cam. Inst. Wash. Dept. Mar. Biol. 24: i. 

CULTURING AMOEBA PROTEUS AND A. DUBIA 

H. R. Halsey, Columbia University 

BOTH Amoeba proteus and A. dubia are found in freshwater ponds 
and streams among aquatic plants such as Cabomba or Elodea, or 
in the debris on the bottom of such bodies of water, particularly among 
rotten leaves. Large individuals are often found in considerable numbers 
among Sphagnum. 

Place small amounts of such material in finger bowls or large petri 
dishes, cover with spring water or with water from the source, and add 
2 or 3 grains of uncooked rice, or an equal number of one-inch lengths 
of boiled timothy hay stalks. Do not place too much of the material in 
a single dish. This results in decay, and in the appearance of large 
numbers of bacteria which cause the death of any Amoebae that may be 
present. 

Amoebae will appear in considerable numbers in successful cultures 
within a week or ten days. The decaying organic material is then 
removed and the Amoebae cultured by the following method. Make*a 
hay infusion of 8 one-inch lengths of timothy hay stalks in 100 cc. of 
spring water, boil for 10 minutes and allow to stand for 24 hours. At 
the end of this time add large numbers of small Protozoa such as Col- 



Amoebidae 81 

pidium [see pp. 63 and 108] and Chilomonas [see pp. 59 and 63] to 
the medium. Allow this to stand for two or three days before using. 
The Amoebae multiply rapidly on this medium so that the bottom of 
the culture dish is soon covered with them. 

These cultures should be examined weekly. Amoeba proteus and A. 
dubia have been observed to injest 50 to 100 Chilomonas within 24 
hours. This results in the rapid disappearance of the food organisms 
from the culture. If the food organisms become few in number pipette 
off half of the culture medium and add an equal amount of fresh pro- 
tozoan hay infusion. At the same time add 2 grains of uncooked rice, 
boiled wheat, or 4 one-inch lengths of boiled timothy hay stalks per 
50 cc. of culture medium. 

The Amoebae in such cultures divide rapidly up to 13 divisions per ten 
day period as judged from organisms kept in isolation. The cultures 
may be run successfully for as long as six months. They may be kept 
at room temperature even during the summer months though it is best 
to use refrigeration if the temperature reaches oo° F. or higher. 

Subcultures are made by pipetting half of the material on the bottom 
of the old culture into a new dish and adding an equal amount of pro- 
tozoan hay infusion and food material. 

This culture method may be varied somewhat, but the following facts 
should be kept well in mind. Large numbers of bacteria in a culture 
tend to cause the death of the Amoebae ; therefore other Protozoa should 
be present in the medium. The use of large amounts of organic ma- 
terial such as boiled rice, wheat, or cracked wheat should be avoided. 
These ferment very readily, causing the death of most of the Amoebae. 
The depression period observed by many investigators is due to this 
cause. If the protozoan hay infusion with a small amount of food is 
used about 90% of the cultures will be successful. This compares with 
only 50% of bacterial cultures in the author's experience. In the former 
case there is no depression period, in the latter a depression period of a 
month is not unusual. 

CULTIVATION OF MAYORELLA (AMOEBA) BIGEMMA 
ON EUGLENA GRACILIS 

John C. Lotze, Ohio State University 

THE food of Mayorella bi gemma is listed by Schaeffer (1918) in his 
original description as "flagellates, ciliates, diatoms, rhizopods, 
nematodes, vegetal tissue, etc." Botsford (1922) reported the culturing 
of Mayorella bigemma in solutions of beef extract. Taylor (1929) 
stated that Mayorella bigemma may easily be cultivated under the same 
conditions as Amoeba proteus. 



82 Phylum Protozoa 

In 1929, the author found a few amoebas of this species in an old hay 
infusion culture of mixed Protozoa. A clone culture was successfully 
established and maintained with good results by using Euglena gracilis 
as the source of food. The amoebas have been cultured by this method 
for a period of over four years. 

Distilled water, well water, and a synthetic well water were used in the 
preparation of culture media. Natural well water proved to be the 
most satisfactory. The source of this water was a well on the campus of 
Ohio State University. The synthetic well water was more satisfactory 
than distilled water ; however, fair results were obtained with the latter. 

Solutions of aminoids* were used in the cultivation of Euglena gracilis. 
A solution of 0.04% aminoids was very satisfactory and had no ap- 
preciable effect upon the amoebas introduced into such cultures. 

Pyrex Erlenmeyer flasks were used exclusively. The best results were 
obtained by using 125 cc. flasks with 75 cc. of culture medium. The 
medium was first made up in a liter flask ; then the proper amount was 
poured into each of the smaller flasks. The flasks were then loosely 
stoppered with sterile cotton and placed on a hot plate. When the con- 
tents just came to a boil, the flasks were removed and allowed to cool. 
Immediately thereafter, the medium was inoculated with Euglena. This 
procedure was very effective in eliminating contamination of the cultures 
by other Protozoa. 

Although it was found that the amoebas could be introduced into the 
cultures with the euglenas, the number of amoebas produced in such 
cultures was never as great as it was when the euglenas were given a good 
start beforehand. Satisfactory results were also obtained when amoebas 
were introduced into Euglena cultures in which the euglenas were passive 
and many were enveloped in gelatinous sheaths. 

A few cultures were maintained for a long period of time by the 
addition of euglenas previously concentrated with a centrifuge. Food 
was added only when that of a culture had become scarce. These addi- 
tions compensated for the loss of water from a culture due to evaporation. 
One culture has been maintained by this method since the spring of 1929. 

Another satisfactory method consisted of adding aminoids to an 
amoeba culture when the euglenas contained in it had become scarce. 
An amount of aminoids sufficient to make a 0.04% solution was roughly 
estimated and added to the culture. An addition of a very large amount 
of aminoids was always followed by a rapid increase in the number of 
bacteria in the culture, and finally, the death of the amoebas. 

No attempt was made to control the hydrogen-ion concentration in 

* Either beef or milk aminoids, commercial products of the Arlington Chemical Co 
Yonkers, N. Y. 



Amoebidae 83 

the amoeba-Euglena cultures. The distilled water used had a pH value 
of 6.0 to 6.2 and synthetic well water made from this was 7.1. The pH 
of the well water was always about 7.2. A 0.04% solution of aminoids, 
freshly made up, sterilized by boiling, and thoroughly cooled for an 
hour or two had a pH of about 6.9 to 7.2. In the course of 3 or 4 weeks 
the pH of such cultures gradually rose to 8.0 or 8.2 and sometimes to 
8.4. This was true regardless of whether they were Euglena or Euglena- 
amoeba cultures. After the cultures had attained this pH, they main- 
tained it without appreciable fluctuation. 

Light conditions favorable for the cultivation of Euglena gracilis were 
not detrimental to the amoebas. All cultures were kept near a north 
window to protect them from direct sunlight. 

The amoebas were easily cultured at ordinary room temperatures. 
Best results were obtained when the temperature was 75 ° F. or slightly 
above. However, cultures were maintained without attention over longer 
periods of time at lower temperatures. 

Bibliography 

Botsford, E. F. 1922. Rhythms in the rate of reproduction of Amoeba bigemma. 

Proc. Soc. Exper. Biol, and Med. 19:396. 
Schaeffer, A, A. 1918. Three new species of amoebas: Amoeba bigemma nov. 

spec, Pelomyxa lentissima nov. spec, and P. schiedti nov. spec. Trans. Amer. 

Micr. Soc. 37:79. 
Taylor, M. 1929. Some further observations on Amoeba proteus. Nature 123:942. 

THE CULTURE OF FLABELLULA MIRA 

D. L. Hopkins, Duke University, N. E. Rice, Brenau College, and 
H. E. Butts, Wellesley College 

A MARINE amoeba, Flabellula mira, named and described by 
Schaeffer (1926), has been found only in the waters around 
Florida. The amoebae used in the development of the following culture 
methods were collected at Tortugas, Florida, in 1929, and have been 
maintained in culture at the Duke University Zoological Laboratory 
since then, making a total of six years in culture. 

These amoebae may be collected from their natural habitat easily. 
With a pipette collect some seawater containing a little seaweed or debris 
from tidal pools or shallow places over a reef. Bring it into the labora- 
tory, pour into petri dishes, dilute considerably with sterile seawater, 
and add 6 grains of wheat to each petri dish. In three or four days the 
amoebae will have become abundant on the bottom of the dish and on 
the surface film. It is very probable that more than one species of 
amoebae will be present and will develop in these cultures. There are 
two or three amoebae of the genus Flabellula from which it is very diffi- 
cult to distinguish Flabellula mira. It is therefore necessary to study 



84 Phylum Protozoa 

them for some time in pedigreed or clone cultures to be sure you have 
the right species. To obtain these amoebae in clone cultures proceed 
as follows: 

With a sterile capillary pipette pass an amoeba through three or four 
changes of sterile seawater in sterile depression slides. From the last 
depression slide transfer it to a small drop of sterile wheat infusion in 
seawater (prepared by boiling about 30 grains of wheat in 100 cc. of 
seawater for about two minutes). Now with sterile vaseline seal the 
coverslip over the depression slide with the drop hanging down from 
the under side of the coverslip. Prepare several hanging-drop clone 
cultures in this way. When several amoebae have developed, sterilize 
the upper surface by washing with absolute alcohol. Allow the alcohol 
to evaporate completely, break the seal with sterile forceps, remove the 
coverslip and drop it with the amoeba-side up into sterile seawater in a 
sterile petri dish. Add 6 grains of wheat sterilized by autoclaving 15 
minutes at 15 pounds' pressure. The amoebae will remain attached to 
the coverslip and thus will easily be found when desired. They will 
become abundant within a week. If evaporation is prevented, the 
amoebae will remain viable for weeks. The cultures, however, come to 
a condition of maximum population in about seven days. They should 
be subcultured every two weeks. 

To subculture the pure clones sterilize wheat, petri dishes, seawater, 
and inoculating pipettes; place seawater and 6 grains of wheat, in a 
petri dish ; then with the pipette draw some of the amoebae up from the 
bottom of an old culture and add them to the new culture medium. 
Growth will proceed as before. 

These amoebae feed on bacteria; therefore there is little difficulty in 
obtaining food organisms. Sufficient bacteria, except in rare cases, are 
carried with the amoeba to inoculate the new cultures even after four 
or five washings. 

The salt content of cultures. F. mira is remarkable in its ability to 
adapt itself to great variations in the salt content of its medium. It may 
be cultured readily and indefinitely in the following modification of the 
artificial seawater of McClendon, Gault, and Mulholland (1917): 
Substance • grris. 

CaClo 1.220 

MgCl 2 .6H 2 5.105 

MgS0 4 .7H 2 7.035 

KC1 0.763 

NaCl 28.340 

NaBr.2H 2 0.082 

NaHC0 3 0.210 

Distilled water 1000 cc. 

Butts (1935) has made a study of the effects of salts on the growth 



Amoebidae 85 

and reproduction of this form. It may be cultured in any concentration 
from a five-fold dilution to a concentration so high that some of the 
salts begin to precipitate. The optimum concentration is about 10% 
distilled water and 90% seawater. The salt ratios may be altered con- 
siderably and still the amoebae will grow and reproduce. 

The hydrogen-ion concentration. The hydrogen-ion concentration 
seems to have but little influence on reproduction. It is possible to cul- 
ture F. mira at any concentration between pH 3.0 and 9.0. Reproduc- 
tion, however, seems to be more rapid in the ranges pH 8.0-9.0, and pH 
5.0-7.0, than in the range pH 7.0-8.0. In artificial or natural seawater 
cultures the hydrogen-ion concentration remains fairly constant, varying 
within the range pH 8.0 to pH 8.4. 

The culture of F. mira on solid media. Rice (1935) has made use 
of silica gel and Bacto agar plates in culturing F. mira. The Bacto agar 
method may be described as follows: Place 100 gm. wheat in 1000 cc. 
of artificial seawater and autoclave for 20 minutes at a pressure of 15 
pounds and a temperature of 125 C. Strain through a cheesecloth, 
and then filter through a coarse filter paper. Now add 15 gms. Bacto 
agar and enough artificial seawater to bring the volume back to 1000 cc. 
Autoclave again at the same temperature and pressure for 30 to 60 
minutes. The resulting agar medium is then tubed, plugged with cotton, 
sterilized again, and set aside for future use. When ready to culture the 
amoebae on this medium, melt the agar in a tube, pour into a sterile 
petri dish, cover, and allow to cool and solidify. Then transfer a small 
drop of liquid culture medium containing amoebae and associated bac- 
teria to the center of the plate, or scatter small drops about over the 
entire surface of the plate. The water will be absorbed by the agar 
and the amoebae and bacteria will grow and develop on the surface. 
The bacteria and consequently the amoebae will become very abundant 
and concentrated. Often the amoebae become so thick that they form 
a sort of epithelial layer over the surface of the agar. To subculture, 
amoebae and bacteria are transferred with a platinum loop to freshly 
poured plates. 

The culture of F. mira on pure strains of marine bacteria. By the 
proper bacteriological methods isolate bacteria from good cultures of F. 
mira. Then pour artificial seawater-wheat-extract-agar plates and inocu- 
late only in the center with amoebae and associated bacteria. Now with 
a platinum loop make a smear of the previously isolated marine bac- 
terium in a circle about 1 cm. from the center inoculated with amoebae. 
Some of the amoebae in traveling the 1 cm. distance from the point of 
their inoculation over a sterile surface lose contaminating bacteria so 
that when they enter the smear of the pure strain of bacteria they are 
sterile. (Some, however, may carry bacteria even this distance.) They 



86 Phylum Protozoa 

immediately feed on these bacteria and multiply rapidly. As soon as 
they have become numerous they should be transferred to a new plate 
and checked by the usual bacteriological methods for contaminations. 
If contaminants are present, the process should be repeated. F. mira 
may be cultured indefinitely on pure strains of bacteria. 

The organic composition oj the medium for F. mira. We have used 
wheat mainly as the original source of organic material. However, a 
variety of substances are adequate, such as i % solutions of sucrose, other 
sugars, or soluble starch; and 0.2% solutions of different amino acids 
and mixtures of amino acids and sugars in either liquid or solid media 
made up in artificial seawater. 

Light and temperature. F. mira may be cultured in any light in- 
tensity between bright sunlight and complete darkness and at any tem- 
perature between io° C. and 42 ° or 43 ° C, the optimum being between 

25° and 35° C. 

Bibliography 

Butts, H. E. 1935. The effect of certain salts of seawater upon reproduction 
in the marine amoeba Flabellula mira Schaeffer. Physiol. Zool. 

McClendon, J. J., Gatjlt, C. C, and Mulholland, S. 1917. The hydrogen-ion 
concentration, CO2 tension, CO2 content of seawater. Cam. Inst. Wash. Dept. 
Mar. Biol. 11:21. 

Rice, N. W. 1935. The nutrition of Flabellula mira Schaeffer. Arch. f. Protist. 

85:350. 
Schaeffer, A. A. 1926. Taxonomy of the amoebas with description of thirty- 
nine new marine and freshwater species. Cam. Inst. Wash. Dept. Biol. 24:1. 

VALKAMPFIA CALKINSI AND V. PATUXENT* 

OYSTERS obtained from markets sometimes yield Valkampfia cal- 
kinsi and V. patuxent from the intestinal tract. These two may 
be cultured easily on ordinary agar plates, yielding abundant parasitic 
material. Without considerable familiarity it is almost impossible to 
distinguish living leucocytes from the parasitic amoebae. 

M. E. D. 
References 

For the culture of Endamoeba blattae and E. thomsoni see p. 128. 
For the culture of Endamoeba ranarum see p. 69. 

NOTES ON VARIOUS MEDIA USED IN THE CULTURE 
OF INTESTINAL PROTOZOA 

Charles A. Kofoid and Ethel McNeh, University of California 

IT IS very important to keep in mind that there are two reasons for 
culturing intestinal Protozoa. The first is to aid the diagnostician 
in determining their presence or absence in the stool specimen. It is not 

♦Abstracted from an article in Science 78:128, 1933, by C. M. Breder, Jr., New 
York Aquarium, and R. F. Nigrelli, New York University. 



Amoebidae 87 

necessary in this case to keep the organism in culture over any long 
period of time. 

The second purpose is to preserve a constant supply of culture mate- 
rial over long periods for animal inoculation, metabolism experiments, or 
immunological studies. Therefore a medium which may be successful 
for the first purpose may be entirely unsatisfactory for the second. With 
this in mind we will give a brief discussion of the following media: Boeck 
and Drbohlav (1925), Dobell and Laidlaw (1926), Kofoid and Wagener 
(1925), Tanabe and Chiba (1928), Cleveland and Collier (1930), Craig 
(1930), Deschiens (1930), St. John (1932). 

Of these, the first three have been so well tested that there is little 
need of discussing them further. They are all satisfactory in maintain- 
ing cultures over long periods of time. We prefer the L. E. A. medium 
of Boeck and Drbohlav to their L. E. S. medium. We feel, however, 
that the growth of Blastocystis and bacteria is less in the L. E. B. 
medium of Kofoid and Wagener (1925). Particularly is this noticeable 
in the culture of Dientamoeba jragilis. We have found, too, that the 
optimum pH for Dientamoeba is lower than for Endamoeba histolytica 
(about pH.6.6). 

Of the others we have found that St. John's medium is excellent in 
producing a sudden increase in rate of multiplication and is thus useful 
for classroom studies as well as for diagnostic purposes. It has also the 
advantage of being relatively inexpensive and easy to prepare. But we 
have never been able to keep the amoebae in this medium longer than 
several weeks. 

St. John, 1932. 

SLANTS LIQUID 

None Heart muscle (Bacto Beef Heart Dehydrated) is extracted 

in a modified Locke's solution by boiling for one hour. 

Heart muscle i gram 

Locke's solution * iooo cc. 

The extract is filtered through filter paper and auto- 
claved at 15 lbs. pressure. "Ralston's" whole wheat 
flour is added. 

Craig's media (as he himself states) are primarily to aid the diagnos- 
tician, and not for the maintenance of cultures by transfer. 

♦Locke's solution: 

Sodium chloride 9.00 gm. 

Calcium chloride 0.24 gm. 

Potassium chloride 0.42 gm. 

Sodium bicarbonate 0.20 gm. 

Dextrose 2.50 gm. 

Distilled water 1000 cc. 



88 Phylum Protozoa 

Craig, 1930. 

i 2 3 

Inactivated Inactivated Inactivated human 

human blood human blood blood serum i part 

serum i part serum i part 0.85% NaCl 7 parts 

Locke's solution 7 parts Ringer's solution 7 parts 
Rice starch is added. 

All are sterilized by Berkefeld filtration. 

Tanabe's medium is satisfactory for a certain length of time, but as 
yet does not justify its use in preference to the Boeck-Drbohlav medium 
or one of its modifications. However, we feel that their effort to simplify 
the medium is a step in the right direction. 

Tanabe and Chiba, 1928. 

SLANTS LIQUID 

Agar 10 grams 5% rabbit serum in Ringer's. 

Asparagin 1 gram Rice starch is added. 

Ringer's solution 1000 cc. 

We have not had as much success with Cleveland's liver-infusion-agar 
medium as was hoped. We agree with him that it seems to be almost 
specific for Endamoeba histolytica, but we find two objections to its use: 
1 ) Blastocystis multiplies very rapidly in it, and, 2 ) production of gas 
by bacteria displaces the agar slants to an annoying degree. The first 
objection seems to us of considerable importance where only fresh smear 
examinations are made. 

Cleveland and Collier, 1930. 

SLANTS LIQUID 

Liver Infusion Agar Horse serum 1 part 

(Digestive Ferments Co.) 30 grams 0.8% NaCl 6 parts 

Na2HP04 3 grams 

Water 1000 cc. 

Rice flour is added. 

We have not, as yet, had opportunity to duplicate all details of 
Deschiens' medium, but it is probable that the results would be some- 
what similar to those of Tanabe. 

Deschiens, 1930. 

SLANTS LIQUID 

Agar 20 grams Locke — Ringer's solution or 

NaCl 5 grams Physiological salt solution 

Beef extract 2-5 grams 

Water 1000 cc. 

Powdered fish muscle or powdered beef muscle is added to each tube. 

Trichomonas, Chilomastix, Enteromonas, and Embadomonas grow 
well in the L. E. A., L. E. B., or L. E. S. media. Giardia has not as yet 



Amoebidae 89 

been cultured, but recent experiments indicate that some sort of tissue 
extract in which the oxygen supply is kept constant will eventually be 
successful. 

Miss Bonestell (working in this laboratory) has found that the various 
species of Trichomonas multiply exceedingly rapidly in rat embryo ex- 
tract, such as is used for tissue cultures. 

Bibliography 

Boeck, W. C.j and Drbohlav, J. 1925. The cultivation of Endamoeba histolytica. 

Amer. J. Hyg. 5:371. 
Cleveland, L. R., and Collier, J. 1930. Various improvements in the cultivation 

of Entamoeba histolytica. Ibid. 12:606. 
Craig, C. F. 1930. The cultivation of Endamoeba histolytica. In Hegner and 

Andrews: Problems and Methods of Research in Protozoology. 532+ ix pp. 

Macmillan Co., New York. 
Deschiens, R. 1930. Culture de l'amibe dysenterique et nutrition de cette amibe 

dans les cultures. Extrait du ier Congres Internal, de Microbiol., Paris, pp. 1-4. 
Dobell, C.| and Laidlaw, P. P. 1926. On the cultivation of Endamoeba histolytica 

and some other entozooic amoebae. Parasit. 18:283. 
Kofoid, C. A., and Wagener, E. H. 1925. The behavior of Endamoeba dysenteriae 

in mixed cultures of bacteria. Univ. Calif. Publ. Zool. 28:127. 
Kofoid, C. A., and McNeil, E. 1931. The advantages of Locke's blood medium 

in the culture of parasitic Protozoa of the digestive tract. Amer. J. Hyg. 

15:315- 
St. John, J. H. 1932. A new medium for the culture of Endamoeba histolytica. 

Amer. J. Trop. Med. 12:301. 
Tanabe, M., and Chiba, E. 1928. A new culture medium for Endamoeba 

histolytica. Acta Medicinalia in Keijo, 11 :i. 

IN VIVO CULTIVATION OF INTESTINAL PROTOZOA 
IN PARASITE-FREE CHICKS* 

Robert Hegner, School of Hygiene and Public Health 

AS EVERY one who has attempted experiments with animal parasites 
L in laboratory animals knows, one of the greatest difficulties is to 
secure parasite-free animals for infection purposes. Chicks offer a num- 
ber of advantages: they may be obtained at any time of the year; they 
are very inexpensive; they are free from animal parasites when they 
hatch from the egg; they may be maintained in the laboratory free 
from animal parasites without difficulty and at low cost ; and they may 
be inoculated very easily per os or per rectum with material containing 
animal parasites. It seems evident that greater precautions are necessary 
to prevent contamination under ordinary laboratory conditions with 
Coccidia than with Amoebae, flagellates, or ciliates. 

Besides being parasite-free and easily maintained in this condition, 
chicks are favorable for experimental studies because one may obtain 

* Reprinted from Science 69:432, 1929, with slight changes by the author. 



qo Phylum Protozoa 

samples from the cecum, where intestinal Protozoa seem to be almost 
entirely localized, without killing the birds or resorting to surgical opera- 
tion. The contents of the cecum are evacuated from time to time and 
this material may be distinguished easily from the intestinal contents 
passed in the form of feces. The fecal material is usually compact and 
dark in color, whereas the cecal contents are more liquid and yellowish 
in color. The best way to obtain cecal material seems to be to give the 
chicks fresh food and water early in the morning and then place them 
under glass dishes on paper towels. Here they may easily be watched 
until cecal material is passed. Some of the chicks will not evacuate 
their cecal contents for several hours, but most of them will deposit the 
desired material within a few minutes. 

The method of procedure followed was usually as follows. The Pro- 
tozoa to be inoculated were obtained either from cultures grown in test 
tubes or from fecal material. If from the former, a more concentrated 
inoculum was sometimes prepared by centrifuging the culture medium 
and pouring off most of the supernatant fluid. If the trophozoites of 
Protozoa were located in fecal material, this mass was diluted with 
normal saline solution and passed through cheesecloth to remove all 
coarse particles that might otherwise clog the passage through the tube 
used for inoculation. Protozoan cysts may be secured in large numbers 
by any of the concentration methods devised for this purpose. A simple 
method is to stir up the infected material in several liters of water in a 
tall, narrow cylinder; allow the cysts to settle to the bottom, which re- 
quires about 30 minutes, then pour off most of the supernatant fluid, fill 
the cylinder with water, stir thoroughly and allow the cysts to settle 
again. After this has been repeated several times the cysts are well 
washed and concentrated. 

A s cc. Luer syringe to which was attached a rubber catheter shortened 
to a length of about 10 cm. was used for inoculating material into the 
chicks. Most of the chicks were about 4 days old, although older birds 
were used for studies of age resistance. The amount of inoculum de- 
pends on the age (size) of the chick. From 2 to 4 cc. of material may 
be injected into the crop of a 4-day-old chick by lubricating the catheter 
with vaseline, inserting it down the throat with one hand while the bird 
is held in the other, and then slowly pushing down the plunger of the 
syringe. Similarly from 1 to 3 cc. may be injected into the rectum. The 
catheter should be inserted about 2 or 3 cm. The anal opening should 
be held closed with the fingers for a few seconds after the catheter is 
removed. Material injected into the rectum appears to find its way 
immediately into the cecum. 

The results of introducing intestinal Protozoa from man and other 
animals into chicks have been prepared for publication elsewhere 



Arcellidae 91 

(Hegner, 1929). They indicate that infections may be set up easily in 
the cecum with a number of species of Amoebae, flagellates, and ciliates. 
Some of the infections continued for over 6 months and apparently would 
have remained indefinitely. Among the Protozoa used were Trichomonas 
hominis from the human intestine and T. buccalis from the human 
mouth. These were maintained in chickens for over 4 months when the 
experiments were terminated. 

One of the most interesting results of the experiments was the dis- 
covery that the chick may be used as a sort of in vivo test tube for the 
cultivation of intestinal Protozoa. For example, cecal material from a 
guinea-fowl which was found by the ordinary smear method to contain 
a very few trichomonads was injected per rectum into chicks. Two days 
later large numbers of trichomonads, Chilomastix, and Endolimax 
amoebae were present in cecal material evacuated by the chicks. The 
trichomonads appeared to belong to two or three different species. On 
the third day the trophozoites of a large Endamoeba were found. 

This work indicates that Protozoa too few in number to be found in 
smears made from the cecal contents of birds such as guinea-fowls, ducks, 
and geese grow and multiply so rapidly when inoculated into parasite- 
free chicks that they may not only be demonstrated without difficulty 
but may be secured in sufficient numbers to prepare permanent slides for 
the detailed study of their morphology. Data already obtained by the 
use of fecal material from other animals inoculated into chicks suggest 
that this method of cultivating intestinal Protozoa in vivo in chicks may 
be extended to include species from other types of animals, especially 
mammals. 

Bibliography 

Hegner, R. W. 1929. Transmission of intestinal Protozoa from man and animals to 
parasite-free fowls. Amer. J. Hyg. 9:529. 



Order testacea, Family arcellidae 

A METHOD OF CULTURING ARCELLAE 
E. D. Miller, University of Virginia 

A WHEAT medium has been found to be very well adapted for cul- 
turing the Arcellae. This is prepared by bringing 300 cc. of 
distilled water to a boil and introducing 15 selected grains of wheat, 
after which the material is set aside to cool. After 24 hours all except 
5 grains are removed. The medium is not used for culture purposes for 
another 24 hours. During the 48 hours sufficient bacteria have accumu- 
lated to serve as food material. 



92 Phylum Protozoa 

There is a tendency toward an over-accumulation of bacteria both 
in the cultures and in the stock supply. The stock supply may be filtered 
through several layers of cheesecloth to remove excess bacterial accumu- 
lations. The material in the culture dishes may be poured off frequently 
and some of the filtered material added. 

CULTURING ARCELLAE 

Bruce D. Reynolds, University oj Virginia 

ARCELLAE may be cultured in a hay infusion as follows: 
Place 10 grams of clean timothy hay in a clean pyrex beaker 
containing 250 cc. of distilled water (pH 6.8). Heat to boiling point 
and allow to boil slowly for 5 minutes, then strain through two thick- 
nesses of cheesecloth and store in quantities of about 3 cc. in small, ster- 
ile, hard glass test tubes. The tubes should then be plugged with cotton 
and placed in boiling water for 15 minutes. After 2 days they should be 
subjected again to boiling for the same period of time in order to kill 
any bacteria which may have escaped the first sterilization by being in 
the spore stage. The medium in these tubes constitutes the stock solu- 
tion and will keep for months without deterioration provided evaporation 
does not take place. 

In making up the culture medium take 1 part of the stock solution 
and add to it 9 parts of distilled water (pH 6.8), giving a 10% hay 
infusion. After the tube containing some of the stock solution has 
been opened and a part of its contents used the remainder should be 
discarded. 

Using the medium prepared in the above manner Arcellae may be 
cultured in hollow-ground slides over a long period of time in a constant 
medium. The culture medium should be changed every day or two and 
the depression slides should be kept in a moist chamber placed in sub- 
dued light. 

References 

For the culture of Arcellae see also pp. 74 and 134. 
For the culture of Arcella vulgaris see p. 60. 



Family difflugiidae 



METHODS OF CULTURING TESTACEA 

A. B. Stump, University of Virginia 

Difflugia oblonga (D. pyrijormis), small varieties [see also p. 136] ; 
D. lobostoma and D. constricta, small varieties; and Lesquereusia 
spiralis may be cultured almost indefinitely by using a number of the 






Foraminifera 93 

green algae for food, such as Spirogyra, Zygnema, Mougeotia, and 
Oedogonium. 

Culturing may be carried on in almost any type of container, though 
the shallow types usually give best results. 

Spring, pond, or tap water may be used, with preference in the order 
named. The pH should be between 6 and 7.3. If small containers, 
depression slides, etc., are used over long periods of time, the water must 
be changed frequently (every 2 to 3 days). Larger cultures in petri 
dishes may be run a week to 10 days. 

Cultures should be kept out of direct sunlight and below 22 C. if 
possible. Small amounts of fine sand should be provided in cultures of 
D. oblonga, D. constricta, and D. lobostoma for shell construction. 



Order foraminifera 

CULTURE METHODS FOR MARINE FORAMINIFERA 
OF THE LITTORAL ZONE 

Earl H. Myers, Scripps Institution of Oceanography 

ALTHOUGH the Foraminifera are universally distributed in the sea 
l and have been the subject of investigations for more than 200 
years, comparatively little is known concerning their methods of repro- 
duction, or of those physiological factors which limit the geographical 
and bathymetric distribution of these organisms (Myers, 1934). Con- 
fusion in the systematic designation of species is frequently due to 
their changing morphology which is the result of an alternation of gen- 
erations, growth stages, a response to environmental conditions, or 
parasitism. In any attempt at a natural classification of these poly- 
morphic forms, it is necessary to recognize their genetic relationship, and, 
where it is possible, their biological explanation should be determined. 
The solution of many of these problems can best be approached by means 
of laboratory cultures. Many species of the littoral zone may be main- 
tained in cultures with a minimum of effort and without the use of 
running seawater. Therefore, this field of investigation offers a splendid 
opportunity for original work to anyone who has access to the sea and 
has acquired a reasonable amount of skill in microscopic technique. 

In selecting a problem in this field, time is an important factor to take 
into consideration, due to the low rate of reproduction in this group as 
compared to other Protozoa. Growth in the majority of polythalamous 
species is a discontinuous process, because of an alternation of a vegeta- 
tive phase with the addition of each newly formed chamber. In small 
species of Discorbis, where the test is composed of a continuous series of 



94 Phylum Protozoa 

from 14 to 19 graduated chambers, from 3 to 12 hours are required for 
the addition of each new chamber. Under optimum conditions _ an 
individual will mature usually in from 19 to 23 days, and at that time 
produce from 30 to 40 young by multiple fission. In the larger species 
of Elphidium, in which the test consists of from 40 to 50 chambers, it is 
doubful whether more than two generations occur annually. Since the 
life span determines the frequency with which one might expect to 
encounter individuals in a state of reproductive activity, it will be less 
difficult to obtain cytological evidence in support of a proposed life cycle 
in small quickly maturing species, and a more abundant supply of ma- 
terial will become available in a given time. 

The following method of collecting and maintaining these organisms 
in culture has proven satisfactory for species of Discorbis, Pyrgo, Trilo- 
culina, Bulimina, Patellina, Spirillina, and Robulus, and should be satis- 
factory for small species found within the limits of the intertidal zone. 

An abundant supply of living Foraminifera may usually be obtained 
by washing seaweed or eel grass vigorously between the hands and allow- 
ing the organisms, sand grains, and other bits of debris to settle through 
a piece of bolting cloth into the bottom of a glass vessel. A convenient 
glass bucket for this purpose is made from a 10- x 14-inch battery jar 
provided with a rope handle and covered with canvas for protection. 
The bucket should be equipped with a tubular net 8 inches deep 
attached to a wooden hoop that will rest on the upper rim of the bucket. 
The vertical sides of the net should be made of unbleached muslin and 
the flat bottom of No. 00 bolting cloth. 

After allowing the organisms about one minute to settle, the water 
should be decanted. Repeated washing by decantation^ will free the 
collection from silt and a considerable amount of organic debris that 
would decompose later. If several collections are to be made the material 
must be transferred to another container. A set of glass refrigerating 
dishes 6 inches in diameter and 2 inches deep that stack one on top of 
the other is convenient for this purpose. A carrying rack should be 
provided for the dishes and, where the collecting ground is some distance 
from the laboratory, it is advisable to control the temperature by packing 
with ice. 

The rate of mortality is high in newly collected material, and we have 
found at La Jolla that certain species that will survive at room tempera- 
ture for more than a year do not reproduce until the temperature is 
lowered to 18 C. or less. Therefore, it is well to sort the material after 
the Foraminifera have become acclimatized to laboratory conditions. 

Crowding of newly collected material should be avoided at all times. 
Not more than 5 cc. of the washings containing the Foraminifera should 
be placed in each of a number of 4-inch round-bottomed finger bowls 



Foraminijera 95 

filled with seawater, or about 20 cc. of this material may be added to a 
1 o-inch crystallizing dish. The dishes should be covered to prevent 
excess evaporation and contamination. 

The water should be changed twice a day for the first few days, and 
after that, once a day for a period of about two weeks. By that time 
many Foraminifera will have crawled up the sides of the dishes and, if 
a suitable substrate of diatoms has developed, several species should 
have become established and reproductive activity begun. 

To establish persistent cultures of a single species, dishes should be 
prepared with a suitable substrate of diatoms before the isolation and 
transfer of the Foraminifera. Pure cultures of Nitzschia [see p. 34], 
Navicula, or similar diatoms may be used for this purpose, or substrate 
material may be used which has been taken from a dish in which the 
Foraminifera have become established and in which the diatom substrate 
is thin, uniform, and free from filamentous algae. If the latter method 
is employed and no new material is added to contaminate the culture, a 
single species of diatom will usually dominate the substrate in a short 
time. 

The day after a foraminifer has reproduced asexually, the young, 
which in some species number 200 or more, remain in the vicinity of 
the parent tests. In establishing subcultures, several thousand individ- 
uals may be transferred in a minimum of time by selecting these groups. 
If this method is employed the age of the organisms will be known and 
a maximum number of individuals of any stage of development will be 
available at a given time. A convenient mouth pipette for handling 
these organisms has been described (Myers, 1933). [see also pp. 43 
and 70.] 

After the cultures are established, it is advisable to change the water 
occasionally to compensate for evaporation and to replace nutrient mate- 
rial removed by the organisms. A strong stream of water directed 
against the sides of a dish by means of a glass syringe equipped with a 
large rubber bulb will remove accumulated debris and help maintain a 
thin clean substrate. 

Before seawater is added to a culture, the water to be added should be 
filtered through a porcelain base Berkefeld or a sintered glass filter of 
suitable porosity. Growth of diatoms may be influenced by controlled 
illumination. When a subdued north light does not produce a suitable 
growth, a few drops of a saturated solution of potassium nitrate may be 
added to each culture or Allen and Nelson's (1910) modification of 
Miquel's solution may be used. [See p. 33.] 

Temperature is a limiting factor in the distribution of species. In 
cultures of Patellina corrugata there is a difference of only 4 to 5 C. 
between the optimum and the upper thermal limit at which this species 



g6 Phylum Protozoa 

can exist. Therefore, in any attempt to culture Foraminifera it is neces- 
sary to avoid temperatures above the mean that prevails in the sea during 
the summer months. 

By employing the simple precautions herein described cultures of a 
number of species of Foraminifera have been maintained for from one to 
three years, and on several occasions have been sucessfully transported 
over land a distance of more than 500 miles. 

References 
Order Heliozoa 
For the culture of Actinophrys see p. 136. 
For the culture of Actinosphaerium see p. 74- 
For the culture of Actinosphaerium ekhhomi see p. 60. 

Bibliography 

Allen, E. J., and Nelson, E. W. 1910. On the artificial culture of marine 

plankton organisms. /. Mar. Biol. Assoc. 8:421. 
Myers, E. H. 1933. A mouth pipette and containers for smaller organisms. 

Science 77:609. 
1934- The life history of Patellina corrugata, a foraminifer. Ibid. 79:436. 

Class Sporozoa, Order coccidiomorpha 

MAINTENANCE OF LABORATORY STRAINS OF AVIAN 
PLASMODIUM AND HAEMOPROTEUS 

Clay G. Huff, University of Chicago 

SINCE methods for the growth of avian parasites of the genera 
Plasmodium and Haemoproteus in the absence of living cells of 
their hosts have not been worked out, these forms must be maintained 
in one of their hosts. While successive generations of Plasmodium from 
man have been "cultured," this has only been accomplished by the 
daily addition of fresh erythrocytes, so that, strictly speaking, this 
amounts to culture in vivo. A method of cultivation has not been worked 
out which is successful in maintaining the strain over long periods of 
time. The invertebrate hosts of all of the species of avian Plasmodium 
whose life cycles have been worked out are culicine mosquitoes [See p. 
376]. Avian species of Haemoproteus are transmitted by parasitic flies 
belonging to the Hippoboscidae [See Huff, p. 446]. 

STRAINS OF AVIAN PLASMODIUM 

All strains of avian malaria of the genus Plasmodium may be main- 
tained by inoculating blood from the infected into a normal bird. 
P. relictum and P. cathemerium may be easily transmitted from bird to 



Coccidiomorpha 



97 



bird by means of various culicine mosquitoes, the best vectors being 
Culex pipiens and C. jatigans [see Huff, p. 386] . The mosquitoes must 
engorge on the infected bird at a time when gametocytes are present in 
the blood. Their bites are infectious for other birds after 8 to 14 days 
depending upon the species of the parasite and the temperature of the 
environment. Plasmodium circumflexum does not infect these common 
mosquitoes but it has been shown by Reichenow (1932) to be trans- 
mitted by Theobaldia anmdata. While several of the species of culicine 
mosquitoes become infected in small percentages when fed on birds with 
infections of Plasmodium elongatum and P. rouxi, no complete trans- 
mission has yet been effected by any of them. 




Fig. 43. — Mosquito-proof cage. 



Unless there is some special reason why mosquitoes are preferred as 
the means of transmission, all of these strains may best be maintained in 
canaries and passed when desired by blood inoculation. Infections pro- 
duced by any of these species of parasites go through an acute stage 
which is followed by a latent period of infection of long duration. Spon- 
taneous cure is rare. Therefore, in most cases, the strains may be main- 
tained most easily as latent infections. New infections may be produced 
in normal birds by inoculating them with blood from the birds with 
latent infections. A few drops of blood are taken from a leg vein into 
physiological saline solution (0.85%) and injected by means of a syringe 
and inoculating needle into a normal bird. This may be done intra- 



98 Phylum Protozoa 

peritoneally, intramuscularly, or intravenously. The technique for 
intravenous inoculation has been described by Taliaferro and Taliaferro 
( 1929). It is to be recommended where heavy infections are desired, and 
particularly for such species as normally produce light infections (P. 
rouxi and P. elongatum). 

It is highly desirable to keep all normal and infected birds in screened 
cages in order to avoid the possibility of mixing infections through the 
bites of mosquitoes. A cage which has been found to be satisfactory for 
this purpose is the mosquito-proof, metal cage shown in figure 43. 

HAEMOPROTEUS INFECTIONS IN PIGEONS 

Since the asexual forms of Haemoproteus do not inhabit the blood, 
it is not possible to maintain these infections by blood transfer as 
described above for infections with Plasmodium. The transference of 
these infections has been accomplished by tissue transplants from infected 
bird to normal bird, but it is a difficult method and rarely meets with 
success. We are, therefore, almost entirely dependent upon the insect 
host for transmission of the infection. 

A great many species of birds carry Haemoproteus infections in nature. 
Of the domestic birds, pigeons and doves are satisfactory for laboratory 
hosts. All known vectors of avian Haemoproteus belong to the Hip- 
poboscidae, a family of ectoparasitic flies. The species found commonly 
on pigeons in tropical and subtropical regions, Pseudolynchia maura, 
may be satisfactorily grown in the laboratory and used for the transmis- 
sion of Haemoproteus columbae (Huff, 1932) [See p. 447]. 

References 
For Coccidia see p. 89. 

Bibliography 

Huff, C. G. 1932. Studies on Haemoproteus of Mourning Doves. Amer. J. 

Hyg. 16:618. 
Reichenow, E. 1932. Die Entwicklung von Proteosoma circumflexum in Theo- 

baldia annulata nebst Beobactungen liber das verhalten anderer Vogelplasmodien 

in Miicken. Jenaische Zeitschr Naturwiss. 67 Festschr. p. 434. 
Taliaferro, W. H., and Taliaferro, L. G. 1929. Acquired immunity in avian 

malaria. J. Prev. Med. 3:197. 

Class Ciliata 

CULTURE MEDIA FOR OPALINIDAE* 

THERE seem to be three major desiderata in culturing Opalinids: 
( 1 ) To supply predigested food. ( 2 ) To avoid free oxygen in the cul- 
ture fluid. (3) To avoid contamination of the culture medium. None of 

♦Abstracted from a paper in Science 72:561, 1930, by Maynard M. Metcalf. Johns 
Hopkins University. 



Holophryidae 99 

the several culture methods that have been suggested since the time of 
Putter's first studies provide the first two desiderata mentioned. The 
third can perhaps be secured by frequent transfer of the animals to new 
culture fluid. Supplying predigested food or foods may not prove diffi- 
cult. On the other hand, to keep the culture free of oxygen is not a 
simple problem. It requires a technique not yet developed, so far as I 
know, for culturing any protozoan, except such as will thrive within an 
agar or gelatin medium. Frequent changing of cultured opalinids to 
fresh culture fluid without introduction of considerable oxygen by ex- 
posure to the air involves still greater technical difficulty. It could doubt- 
less be done with the aid of a gas mask in an oxygen-free room. 

Protoopalinae, when kept in Putter's or Locke's solution, either with 
or without bits of the rectal wall of the host, show signs of abnormality 
within a few hours, often within 4 hours or so. These facts, and the 
further fact that in this country Protoopalinas are available for study 
only in a few regions and in the northeastern states not at all, have made 
me hesitate to attempt to develop a culture medium and culture methods. 
On the other hand, given a suitable culture medium and procedure, the 
prompt response by Protoopalina by visible cytological changes under 
unfavorable conditions might render Protoopalina a peculiarly favorable 
test animal for studies of protozoan physiology. 

References 
For Opalina ranarum see p. 69. 

Bibliography 

Konsxjloff, S. 1922, Untersuchungen iiber Opalina. Arch. f. Protist. 44:285. 
Larson and Allen, 1928. Further studies on the reaction of Opalina to various 
laboratory culture media. Univ. Kansas Sci. Bull. 18:8. 

M. E. D. 

Order holotrichida, Family holophryidae 

CULTURE MEDIUM FOR THE CILIATE LACRYMARIA* 

MAST, who has had wide experience in collecting Protozoa, says 
(1911, p. 230) : "Lacrymaria is relatively scarce in nature. It is 
occasionally found in cultures containing decaying aquatic plants but 
never in great numbers. One rarely finds more than two or three speci- 
mens in a drop of solution." No one has succeeded heretofore in culti- 
vating it in the laboratory. 

Malted milk in distilled water was prepared in two sets, one of which 
was boiled and the other not. Both were seeded with Lacrymaria and 

* Abstracted from a paper in Science 63:212, 19:6, by Y. Ibara, Johns Hopkins 
University. 



IO o Phylum Protozoa 

examined from time to time for several weeks. In cultures containing 
1-5 mg. of malted milk to ioo cc. of water the Lacrymaria became very 
abundant and continued to thrive for more than 6 weeks without adding 
anything to the cultures. 

These cultures contained Halteria and another similar organism 
which was not identified and numerous bacteria. The Lacrymaria were 
observed to capture Halteria, but they appeared to feed mostly on the 

other organisms. 

Bibliography 

Mast, S. O. 1911. Habits and reactions of the ciliate Lacrymaria. J. Animal 

Behav. 1:229. 

M.E. D. 

THE CULTURE OF DIDINIUM NASUTUM 

C. Dale Beers, University of North Carolina 

THE food of Didinium nasutum is restricted by the nature of its organs 
of food capture to a limited number of relatively large ciliates. 
Paramecium caudatum and P. aurelia are the forms that are most readily 
and commonly ingested, though large specimens of Colpoda, Colpidium, 
and Frontonia are sometimes eaten. Of these food organisms Para- 
mecium caudatum is recommended as the most satisfactory. It is 
easily cultured [See pp. 112 — 128.] and is wholly adequate to sustain 
Didinium indefinitely. The culture of Didinium therefore resolves itself 
into providing the animals with a non-nutrient fluid medium of suitable 
tonicity and reaction and with an abundance of Paramecia. 

The almost complete dependence of Didinium on Paramecium as a 
source of food means that as a rule Didinium is found in nature only 
where Paramecium is abundant. Pools and streams that contain a con- 
siderable amount of decaying organic matter, including preferably a small 
amount of sewage, are suitable collecting places. Sediment, submerged 
decaying leaves, and plant stems should be collected, as well as water 
from the edges of the pool or stream, for more often Didinium is collected 
in the encysted condition, the cysts lying free in the sediment or cemented 
to submerged objects. To this material an equal volume of vigorous 
Paramecium culture should be added, in order to activate the cysts and to 
induce the rapid multiplication of active specimens. I have always used 
timothy hay infusion for growing the Paramecia for Didinium cultures. 
Cysts will usually activate in hay infusion within a day or two, excysta- 
tion being induced by the environmental change from freshwater to 
infusion. The presence of the Paramecia is not a prerequisite to the 
excystation process; the same result may be obtained with old hay 
infusion which has never contained Paramecia. 

The food supply becomes rapidly depleted in Didinium cultures, since 



Holophryidae 101 

four or five generations are usually produced at room temperature 
(21 C.) in every 24-hour period, and at least two Paramecia are needed 
for a single Didinium to attain its full growth prior to division. At higher 
temperatures the voraciousness and rapidity of reproduction of Didinium 
are astonishing, in that as many as nine generations may be produced 
within 24 hours, and the task of supplying Paramecia to large cultures 
becomes overwhelming. 

When Didinia are needed for class use or for research purposes over a 
period of days or weeks, they are therefore best maintained in small 
stock cultures, made up, for example, in watch glasses. In the prepara- 
tion of these cultures chemically clean watch glasses are filled with the 
desired amount of spring water or pond water (filtered or boiled to re- 
move or kill Entomostraca) . Or they may be filled with 0.01% modified 
Knop solution or with diluted hay infusion from a Paramecium culture. 
Paramecia are then concentrated with the centrifuge and transferred to 
the watch glasses, after which a few specimens of Didinium are added. A 
temperature of about 20 C. is the optimum for the growth of Didinium. 

The Paramecia should react normally soon after being transferred 
to the watch glasses. A medium that is injurious to Paramecium is un- 
suitable for the growth of Didinium, and, conversely, any fluid medium 
in which Paramecium exhibits normal behavior and retains its normal 
cell configuration is usually favorable for Didinium. Occasionally spring 
water and pond water have a slightly unfavorable hydrogen-ion concen- 
tration which needs to be corrected by the addition of about 5 cc. of 
phosphate buffer mixture of pH 6.8 to each 100 cc. of water. The range 
of hydrogen-ion concentration which active specimens of Didinium can 
tolerate is considerable. It varies from pH 5.0 to pH 9.6, but the 
optimum lies between pH 6.8 and pH 7.2. 

Modified Knop solution of suitable concentration (about 0.01%) may 
be prepared conveniently from the usual three stock solutions: namely, 
10% Ca(N0 3 ) 2 , 5% KNO3 and, 5% MgS0 4 . 7 H 2 0. A 1% solution is 
first made up by adding the following amounts of the stock solutions to 
150 cc. distilled water: 10 cc. Ca(N0 3 ) 2 , 7-5 cc. KN0 3 and 7.5 cc. 
MgSO.j. Then a 0.01% solution is prepared by adding 1 cc. of the 1% 
solution to 99 cc. of distilled water. Finally, 5 cc. of NaOH-KH 2 P0 4 
buffer mixture of pH 6.8 is added to each 100 cc. of 0.01% solution. 

This solution is often preferable to spring water or pond water for 
maintaining cultures for experimental purposes, in that its chemical 
composition is known and constant. It has an added advantage in that 
it is unfavorable for the growth of bacteria. 

Hay infusion, either from a flourishing Paramecium culture or from 
an old, declining culture, usually has a favorable hydrogen-ion concentra- 
tion, but it is sometimes too concentrated, and in the preparation of 



NAVY PRO IFTT m 



102 Phylum Protozoa 

stock cultures it should be diluted with one or two volumes of distilled 

water. 

New stock cultures should be prepared every three or four days. 
Excessive bacterial activity and an accumulation of metabolic waste are 
inimical to the growth of Didinium, and may lead to the production of 
structural abnormalities, or to encystment, conjugation, or death. 

Pure lines of Didinium may be maintained with no great difficulty in 
depression slides kept in moist chambers. The procedure is similar to 
that employed with stock cultures, but owing to the high rate of reproduc- 
tion and consequent accumulation of waste, transfers should be made 
daily to fresh slides. 

A seemingly inherent predisposition toward conjugation sometimes 
leads to difficulty in the culture of pure lines, and a high mortality among 
exconjugants sometimes leads to the loss of the stock cultures, if con- 
jugation assumes epidemic proportions. Fortunately, the tendency to 
conjugate may be suppressed in most cases by keeping the cultures at a 
temperature which never exceeds 2 1° C. On the other hand, conjugation 
may often be induced by increasing the temperature from 2i°C. to28°C. 
Some races rarely conjugate; others conjugate frequently, and it is some- 
times advisable to discard the latter. Conjugants are distinctly smaller 
than vegetative individuals, and the death of the exconjugants is due in 
some instances to their inability to ingest large Paramecia. A diet of 
smaller Paramecia may enable them to survive. 

Didinium encysts readily as a result of absence of food, accumulation 
of metabolic waste, or excessive bacterial growth. Of the three factors, 
absence of food is most effective in inducing encystment, and it is the 
factor that is most easily controlled experimentally if it is desirable to 
obtain cysts. Exhaustion of the food supply leads commonly to the 
encystment of the animals within eight to twelve hours. While the cysts 
in my experience do not withstand desiccation, they will remain viable 
for five or six years in water, and it is often convenient to keep Didinium 
from year to year in the encysted condition. To store cysts for future 
use, stock cultures may be made up in vials instead of watch glasses. 
When the Paramecia are all consumed, many of the Didinia will encyst 
on the sides and bottom of the vial, or on pieces of hay, if these are pres- 
ent. Didinium usually encysts against a solid object, the cysts being 
cemented to the object by the gelatinous ectocyst. The vials should be 
only half filled with fluid, so that some air will be present when they are 
finally stoppered and stored away. The cysts may be activated later by 
replacing the fluid in the vials with hay infusion from a vigorous Para- 
mecium culture. 

All Paramecia used in the culture of Didinium should be well-fed speci- 
mens. Much of the difficulty which has been experienced in culturing pure 



Ophryoglenidae 103 

lines of Didinium is the result of feeding the Didinia on underfed Para- 
mecia from an old, declining culture. All attempts to maintain cultures 
of Didinium on starved, emaciated Paramecia have led without exception 
in my experience to excessive conjugation, loss of the ability to encyst, 
reduced division rate, structural abnormalities, and death of the Didinia 
within two or three weeks at most. 

References 

For the culture of Didinium see also p. 63. 
Family Chilodontidae 

For the culture of Chilodon see pp. 55 and 62. 
For the culture of Chilodon cucullulus see p. 104. 

Family ophryoglenidae 

CONTROLLED CULTURES OF FRESHWATER CILIATES 

Alford Hetherlngton, Stanford University 

THE original mass-cultures which provide a varied abundance of 
ciliates for observation and description prove unsatisfactory for the 
needs of modern experimentation. Control of culture media has pro- 
ceeded along two lines, the traditional objective of pure culture (i.e., cul- 
ture in the absence of other life), and culture on a controlled source of 
living food. Pure culture is so far successful only for some of the smaller 
ciliates. 

At first sight, culture of the larger ciliates such as Stentor coeruleus or 
Bursaria truncatella on pure cultures of, for instance, the small ciliate 
Glaucoma pyriformis, would seem to promise a peculiarly elegant con- 
trol of these animals which are so easily manipulated with the unaided 
eye. For very perfect control the food animal could be washed before in- 
troduction in known amounts into a simple inorganic medium containing 
them. 

However it turned out that the conditions of survival of the larger 
free-livi-ng ciliates are much more complex than was anticipated, an out- 
come regretted by those who wish to use them simply as material for 
physiological investigations, but of interest to those who are concerned 
with the biology of micro-organisms as such. 

PURE CULTURE 

Sterilization of Ciliates. A method combining the advantages of mi- 
gration and of washing was described by Hetherington (1934a). The 
practice of bacteriological methods is assumed in that which follows. 



104 Phylum Protozoa 

Glaucoma pyrijormis (35-60/z)* grows as vigorously as any ciliate 
found in freshwaters. [See also p. 54-] It will grow on heavy suspen- 
sions of Bacterium colt, and is unique in that it adjusts promptly to, and 
grows vigorously in, sterile 0.5% Difco powdered yeast extract. Indi- 
viduals are sufficiently large to be counted under the low powers of 
binocular dissecting microscopes. 

Several strains have been described. Lwoff's (1932) and Elliott's 
(1933) grow on peptone, Butterfield's (1929) grows for several transfers 
on peptone, but ultimately demands yeast extract, while a strain isolated 
by Hetherington (1936) shows no growth on peptone. Elliott (1935) 
observed nutritive differences between peptone-growing strains. Glaser 
and Coria (1935) isolated strains from hot springs which will tolerate 
temperatures of 37 C. 

Extensive trials of different media (Hetherington, 1936) for Glaucoma 
Pyrijormis resulted in the following optimum medium: 

Yeast Extract, Difco 0.5% 

Powd. Whole Yeast, Difco B13412 0.2% 

This is made up in Peters' medium (see p. 000), an inorganic physio- 
logical salt solution, and autoclaved at 15 lbs. for 15 minutes. For stock 
cultures, 125 cc. Erlenmeyer flasks are convenient vessels. Division 
rates observed were rather constant, varying from 7.97 to 8.03 at 25 C. 
Glaucoma scintillans (50-65/x), a typical holotrichous ciliate, was first 
grown in pure culture by Chatton in 1929, later by Hetherington (1933). 
[See also p. 56.] Neither investigator achieved dependable growth. Un- 
published recent work by the writer resulted in the following medium: 

Yeast Extract, Difco 0.5% 

Powd. Whole Yeast, Difco B13412 1.0% 

Division rates observed vary from 2 to 2.7 at 25 C; the ciliates 
appear normal and well nourished. While these rates are much lower 
than those resulting from culture on single strains of bacteria, growth in 
pure culture is now satisfactory for many experimental purposes. 

Other freshwater ciliates which have been obtained in pure culture are 
Colpoda cucullus, C. steinii, Chilodon cucullulus, and Loxocephalus 
granulosus. These investigations are reviewed by Hetherington ( 1934b) . 

* This ciliate was called Paramecium in the first paper of Peters (1920). Lwoff 
(1923) first, and Peters (1929) later, called it Colpidium colpoda. It was called Colpidium 
by Butterfield (1929), Colpidium campylum by Hetherington (1933) and by Taylor, et 
o-l- (i933), Colpidium striatum by Elliott (1933), and Saprophilus oviformis by Glaser 
and Coria (1935). Saprophilus oviformis, Glaucoma pyrijormis, and Colpidium striatum 
are probably synonyms (Kahl, personal communication), but Colpidium campylum is a 
common and distinct ciliate which is distinguished by the possession of a conspicuous 
gullet, inconspicuous oral membranes, and by the fact that it will not grow in pure culture 
in Difco yeast extract (or peptone, tryptone, or neopeptone, Difco). 



Ophryoglenidae 105 

Pringsheim (1915) grew Paramecium bursaria with its green commensals 
in an inorganic medium similar to Knop. 

In addition, Paramecium caudatum may be grown in an enormously 
complex medium developed by Glaser and Coria (1935). The writer 
(unpublished work) verified these results using both rabbit and monkey 
kidney ; here again growth is not as vigorous as in the presence of living 
bacteria. Failure, reported previously (Hetherington, 1934c), was due 
to the use of a Seitz bacteriological filter rather than a Berkefeld. The 
filter pads, not the metal of the Seitz filter, contribute a toxic substance 
to the filtrate. 

While it may stretch the definition of pure culture somewhat to include 
a medium which contains fresh rabbit kidney, these investigations are 
illuminating, as will be indicated in the next following section. 

CULTURE ON SINGLE STRAINS OF BACTERIA 

Freshwater ciliates capture and ingest their food, as higher animals 
do. There is no evidence that they can obtain energy or build proto- 
plasm from simpler nutrients than required by higher organisms. 

A division rate of 8.28 may be maintained in the case of Glaucoma 
scintillans on living bacteria in non-nutrient salt solution (25 C). A 
small quantity of Bacillus megatherium, strain D 20 of the Stanford 
University Bacteriology Department Collection, having a diameter of 
about 1 mm., is removed from a 24-hour nutrient plate by means of a 
platinum needle, put in 0.5 cc. of Peters' medium, together with one 
Glaucoma. Transfer is made every 24 hours. No difference can be 
observed between washed and unwashed bacteria. 

Turning to larger ciliates, Colpidium colpoda (nop) grows with a 
division rate of 4.33 (24 C.) on Aerobacter aerogenes, strain A 2 Stan- 
ford Collection, in the following medium (Hetherington, 1934b) : 

Peptone, Dif co o.i % 

Dextrose, Baker's C. P. powd 0.1% 

This is made up in Peters' medium, and autoclaved at 15 lbs. pressure for 
15 minutes. 

Colpidium campylum (50-90/*) has similar nutritive requirements, 
but is more resistant, and will grow faster in a 0.35% concentration of 
nutrient. Like Colpidium colpoda, it has never been grown in pure 
culture. 

Paramecium caudatum and P. aurelia grow luxuriantly in the same 
system (Aer. aerogenes -f- 0.2% peptone-dextrose), either in tiny isola- 
tion volumes or in flasks. This is a more constant medium than the 
Pseudomonas otio/w-powdered lettuce medium reported by Giese and 
Taylor (1935), which is a technique apparently borrowed from the ex- 



io6 Phylum Protozoa 

tensive work of Phelps (1934)- It is not clear why Phelps found other 
particles (lettuce) in addition to living bacteria in nutrient medium 
necessary for the growth of Paramecium, but it is a general rule that a 
few drops in an isolation dish is not an ideal volume for developing 
optimum culture media. 

The writer has added sterile Didinium to a culture of Paramecium 
caudatum on Acr. aerogenes (= "dreigliedrige Kultur") in liter flasks. 
The cycles of growth, perfectly visible to the unaided eye, are rather 
striking, and are duplicated indefinitely with subcultures. Such cultures 
are easy to maintain; a minute or two for transfer once a month suffices. 

In no case has the writer found that a combination of two or more 
bacteria is better than a suitable single strain for the nutrition of ciliate 
Protozoa. 

Colpidium colpoda, Paramecium caudatum or P. aurelia, Pleurotricha 
lanceolata (Penn, 1935), and Urocentrum turbo (Hetherington, 1934b, 
p. 637), will not grow in an inorganic medium (for instance Peters' 
medium or spring water) -f washed living bacteria. The writer there- 
fore cannot agree with Phelps (1934) that some labile substance in 
the bacterial cell is destroyed upon killing (regardless of method used — ■ 
heat, ultraviolet light, HC1, H 2 2 , (NH 4 ) 2 S0 4 , and toluene), and that 
this is the reason pure culture may not be achieved. It would appear 
rather, since even living bacteria do not suffice when their metabolites are 
removed, that labile substances pass into the medium which "condition" 
it in some way, and that these substances must be constantly supplied by 
metabolizing bacteria. The idea that they are reducing bodies was tested 
by Hetherington (1934b, p. 636), with negative results. Before growth 
on dead bacteria may be achieved, unheated liver (Eli Lilly No. 343) 
and fresh, sterile kidney must be added (Glaser and Coria, 1935) . Para- 
mecium may be the simplest type of animal to the taxonomist, but its 
dietary requirements remind us more of mammals! 

Such is the setting for a consideration of the growth requirements of 
the largest ciliates, which typically eat other Protozoa. 

CULTURE ON SINGLE STRAINS OF PROTOZOA 

The large and beautiful Stentor coeruleus (200-500/x) and Bursaria 
truncatella (300-400^) are known to eat a variety of foods: bacteria, 
green and colorless flagellates, diatoms, ciliates, rotifers, and even smaller 
individuals of their own kind. 

Inorganic medium. Of all the physiological media tested, including 
a variety of spring waters, Peters' medium has proved almost ideal for 
freshwater ciliates* : 

♦Preparation is described in the author's 1934a work, p. 316. 



Ophryoglenidae 107 

Ca(HC0 3 )2 0.00055 M. 

MgS0 4 0.00015 M. 

KH0PO4 0.00030 M. 

NaH 2 P0 4 0.00015 M. 

Since ordinary distilled water is frequently very toxic, it is best to use 
twice distilled water for preparing Peters' medium. 

Glaucoma pyrijormis, Colpidium campylum, or Colpidium colpoda, 
carefully washed free of their medium, and added in sufficient amounts 
(an excess), will support growth of Stentor coeruleus in this medium for 
about four weeks (Hetherington, 1932). After this they degenerate. 

Bursaria truncatella behaves similarly, but degenerates much more 
promptly. Addition of Gonium pectorale, a green form, Chilomonas, a 
colorless flagellate, and several kinds of washed bacteria, does not alter 
behavior in the least. Bursaria truncatella, in the presence or absence 
of washed food, dies in physiological medium exactly as described for 
Pleurotricha lanceolata by Penn (1935, p. 126-27). 

Accordingly, the present status of our knowledge concerning these 
ciliates indicates that suitable food -f- pure physiological medium do not 
meet their requirements, that certain metabolites of growing bacteria are 
essential for the continued survival of the larger ciliate Protozoa. Further 
progress with these animals awaits solution of this problem. 

References 

For the culture of Glaucoma ficaria see p. 54. 

For the culture of Colpidium see also p. 63. 

For the culture of Colpidium striatum see p. 54. 

For the culture of Colpidium campylum see also p. 54. 

Bibliography 

Butterfield, C. T. 1 929. A note on the relation between food concentration in 

liquid media and bacterial growth. U. S. Pub. Health Repts. 44:2865. 
Chatton, E., et Chatton, M. 1929. Les conditions de la conjugaison du Glaucoma 

scintillans en cultures lethobacteriennes. Action directe et specifique de certains 

agents zygogenes. C. R. Acad. Sci. 188:1315. 
Elliott, A. M. 1933. Isolation of Colpidium striatum Stokes in bacteria-free 

media. Biol. Bull. 65:45. 
1935- Effects of carbohydrates on growth of Colpidium. Arch. f. Protist. 

84:156. 
Giese, A. C, and Taylor, C. V. 1935. Paramecia for experimental purposes 

in controlled mass cultures on a single strain of bacteria. Ibid. 84:225. 
Glaser, R., and Coria, N. A. 1935. The culture and reactions of purified Protozoa. 

Amer. J. Hyg. 21:111. 
Hetherington, A. 1932. The constant culture of Stentor coeruleus. Arch. 

f. Protist. 76:118. 

1933- The culture of some holotrichous ciliates. Ibid. 80:225. 

1934a. The sterilization of Protozoa. Biol. Bull. 67:315. 

1934b. The role of bacteria in the growth of Colpidium colpoda. Physiol. 

Zool. 7:618. 



108 Phylum Protozoa 

1934c The pure culture of Paramecium. Science 79:413- 

1936. The precise control of growth in a pure culture of a ciliate, 

Glaucoma pyriformis. Biol. Bull. 71:426. 
Lwoff, A. 1923. Sur la nutrition des Infusoires. C. R. Acad. Sci. 176:928. 

1932. Recherches biochimiques sur la nutrition des Protozoaires. Paris, 

Masson et Cie. 
Penn, A. B. K. 1935. Factors which control encystment in Pleurotricha lanceolata. 

Arch. f. Protist. 84:101. 
Peters, R. A. 1920. Nutrition of the Protozoa: the growth of Paramecium in 
sterile culture medium. J. Physiol. 53: ioS - 

1929. Observation upon the oxygen consumption of Colpidium colpoda. 

Ibid. 68:1. 

Phelps, A. 1934. Studies on the nutrition of Paramecium. Arch. j. Protist. 82 : 134. 
Pringsheim, E. G. 1915. Die Kultur von Paramecium bursaria. Biol. Zentralbl. 

35:375- 
Taylor, C. V., Thomas, J. O., and Brown, M. G. 1933 • Studies on Protozoa, 
IV; Lethal effects of X-radiation of a sterile culture medium for Colpidium 
campylum. Physiol. Zool. 6:467. 

CULTIVATION OF COLPIDIUM CAMPYLUM* 

T. M. Sonneborn, Johns Hopkins University 

USE a basic fluid of spring water in which rye grains, in the concentra- 
tion of 1.5 gms. per 100 cc, have been boiled for 10 minutes. The 
fluid is filtered while hot and allowed to stand 1 day exposed to the air. 
Good mass cultures can be obtained within a day or so by placing 200 cc. 
of ripe fluid in a finger bowl and inoculating with 10 to 15 cc. of an old 
culture of C. campylum. Such cultures should be renewed every 2 to 6 
days. 

Isolation cultures under bacteriologically controlled conditions may 
readily be carried by the following method: Distribute the standard 
rye infusion as soon as filtered into test tubes, plug with cotton, auto- 
clave, and store till needed. When ready for use, open tube in a flame 
and inoculate by means of sterile platinum needle with pure cultures of 
the bacterium, Achromobacter candicans, grown on beef-agar slants. 
Using glassware (petri dishes, Columbia dishes, pipettes, etc.) previously 
baked in an oven at 150 C. for an hour, the one-bacterium rye fluid is 
pipetted into Columbia dishes inside of petri dishes and into these dishes 
sterilized Colpidia are introduced by means of a sterile fine pipette. 

Isolation cultures carried with less refined technique, may be main- 
tained by using depression slides containing one drop of one-day-old 
ripened rye infusion. These should be started with one Colpidium to 
each drop of culture, and should be renewed daily by transferring one 
Colpidium to a fresh drop on a fresh slide. A favorable temperature is 

22°-23°C. 

* Condensed by the author from Biol. Bull. 63:187, 1932. 



Ophryoglenidae 109 

THE CULTURE OF COLPIDIUM CAMPYLUM 

C. V. Taylor, J. O. Thomas, and M. G. Brown, Stanford University 

THE holotrichous ciliate, Colpidium campylnni, was obtained orig- 
inally in an enrichment culture from spinach procured in the vicinity 
of Monterey Bay, California, and subcultured in a continuously thriving 
condition in a sterile tap water extract of common commercial yeast. 

This extract is prepared in the following manner: 450 grams of a com- 
mercial yeast is mixed with 500 cc. of water, kept at 50 C. for 24 hours, 
then neutralized with XaOH to a pH of 7.0, filtered, and autoclaved. 

For cultures the filtrate is diluted ten times with tap water, then appor- 
tioned in 5 cc. amounts into test tubes. Sterile technique is used through- 
out. Each tube is inoculated with approximately 1000 Colpidia. The 
resulting growth is vigorous and uniform. 

Under ordinary laboratory conditions pure strains of this holotrichous 
ciliate, washed free of bacteria, have thrived vigorously for about two 
years in this convenient and reproducible medium. The temperature 
variation was around 20 to 22° C. and the light in the laboratory was a 
north light. 

A CULTURE METHOD FOR COLPODA 

M. S. Briscoe, Storer College 

FINGER bowls are satisfactory vessels for this culture method. They 
are first sterilized by exposing them to streaming steam. In this way 
micro-organisms normally present upon the surfaces of laboratory appa- 
ratus are destroyed. Both moist and dry heat were tried but the former 
is more efficient. If there is no Arnold steam sterilizer in the laboratory 
the sterilizing process may be accomplished in some similar apparatus. 
A temperature of ioo° C, is sufficient to destroy the majority of micro- 
organisms that may be present. 

With the completion of the sterilizing process, spaghetti, which may be 
purchased in cans at a grocery store, should be removed from its con- 
tainer. It should be thoroughly washed so as to remove any other sub- 
stances that may be present. When all of these have been eliminated 
place some of the spaghetti in a finger bowl and cover it completely 
with water. Faucet water is satisfactory. Allow it to stand for several 
days until a scum forms. It is then ready for inoculation with the 
organisms. 

The optimum growth temperature for Colpoda is 70 F. At this 
temperature the organisms appear normally healthy and grow very 
rapidly. Increases in temperature do not increase the rate of growth 
and at sufficiently high temperatures growth ceases. The organisms do 
not thrive when exposed to intense light such as the direct rays of the 



II0 Phylum Protozoa 

sun. Cultures which were not exposed to the air, and hence did not 
become moldy, continued to thrive for long periods of time. 



STOCK CULTURES OF COLPODA* 

DURING the course of investigation with Protozoa, a rather con- 
venient and easy method of obtaining and keeping stock cultures 
of Colpoda was found. 

Colpoda, as is well known, usually occur early in soil cultures from 
which they may be obtained, in the active state, in large numbers. Later 
in the life of the culture the animals encyst and it is upon this condition 
that the following method is based. 

From a young soil culture active Colpoda are isolated, transferred to 
Syracuse watch glasses and ordinary hay infusion added. After one or 
two days the culture fluid in the watch glass is allowed to evaporate 
slowly by exposure to the air. During this slow evaporation the animals 
encyst. The dried up culture is left exposed for one or two days, when 
new hay infusion is added. The animals, having divided within the 
cysts, revive and are found in greatly increased numbers. This drying 
process may be repeated until a more or less concentrated culture of 
organisms is obtained. The concentrated culture of organisms is then 
pipetted into a petri dish in which a piece of ordinary filter paper, cut so 
as to exactly cover the bottom of the dish and moistened with hay infusion, 
is placed. The petri dish is then left uncovered to evaporate slowly. The 
filter paper, with the encysted organisms on it, when thoroughly dry, may 
be cut into small pieces and kept indefinitely. 

To start fresh cultures, pieces of the filter paper are put into watch 
glasses or other containers and hay infusion added. In a short time the 
animals revive and new cultures of the original are thus obtained. 

This method of keeping stock cultures seems to be especially adapted 
for schools and colleges where only a limited amount of time is devoted 
to the Protozoa and where no time for the ordinary culture preparation 
work is available. 

M. E. D. 

Reference 
For the culture of Colpoda see also p. 63. 

THE CULTURE OF COLPODA CUCULLUS 

H. Albert Barker and C. V. Taylor, Stanford University 

Colpoda cucullus may be cultured in finger bowls containing 10-20 cc. 
of a dilute hay infusion. These infusions, however, are singularly unsuit- 

* Reprinted with slight changes from an article in Science 53:92, 192 1, by Joseph H. 
Bodine, Iowa State University. 



Ophryoglenidae 1 1 1 

able as media with which to conduct certain types of experimental in- 
vestigations because their composition is unknown and uncontrollable and 
doubtless varies greatly as prepared at different times and places. 

For these reasons we have used the reproducible, non-nutritive medium 
shown in the following table, supplemented by the addition of a pure 
strain of bacteria for food. 

Balanced Physiological Medium 

(after Osterhout, 1906) 

67c total salt (0.937 M.) 
0.937 M. Parts 

NaCl 1 ,000 

MgCl 2 .6H 2 78 

MgS0 4 . 7 H 2 38 

KC1 , 22 

CaClo 10 

This solution is diluted five times to the approximate concentration 
of pond water, or 0.012% total salt (0.0018 M.). One cc. of M/20 
NaH 2 P0 4 is added to each 30 cc. of this diluted solution for buffering, 
and the pH adjusted with suitable quantities of M/20 NaOH. 

Colpoda in this medium are fed upon suspensions of a pure culture 
of the bacterium Pseudomonas fluorescens. This bacterium is eminently 
suitable as a food for Colpoda. When grown in suspensions of it, they 
appear large and well fed, their division rate is equal to or greater than 
that in the most favorable hay infusions, they continue normally active, 
and may be kept indefinitely on this single diet without degeneration. 

Pseudomonas fluoresceins may be isolated from soils by the use of a 
liquid enrichment culture in which 2% asparagine serves as the only 
source of carbon and nitrogen. The bacterium may be isolated and grows 
rapidly on ordinary yeast — or peptone-glucose-agar plates at 37 C. or 
below. If inoculated heavily on such plates, the colonies spread out 
within a few days into a continuous sheet from which masses of bacteria 
may be removed with a sterile platinum loop in order to be fed to the 
Protozoa. 

The density of the bacterial suspension provided for the Protozoa is 
difficult to define except as that density in which optimal growth occurs. 
This most favorable density of bacterial suspension will of course depend 
in part upon the number of Protozoa inoculated into a given volume of 
the medium. An approximate idea of a favorable suspension density may 
be gained from the following illustration: the mass of bacteria which will 
fill a platinum wire loop 1 mm. in diameter is sufficient to make 6 
suspensions of % cc - into each of which 20 Colpoda are to be inoculated. 

No effort is made to avoid air contaminations of the Pseudomonas 
suspensions. It was assumed that the suspensions would not be suffi- 



ii2 Phylum Protozoa 

ciently nutritive to permit the growth of contaminating micro-organisms. 
Justification for this may be seen in the fact that molds were never 
observed to develop during the experiments. 

All experiments were carried out in a constant temperature room at 
72 ° F. with north light. 

The containers for the culture of Colpoda are watch glasses of about 
2 mm. diameter enclosed in petri dishes of suitable size containing in the 
bottom a small amount of water and so used as moist chambers. 

References 

For the culture of Colpoda cucullus see also p. 104. 
For the culture of Colpoda steinii see pp. 56 and 104. 
Family Urocentridae 

For the culture of Urocentrum turbo see p. 106. 

Bibliography 

Osterhout, W. J. V. 1906. Extreme toxicity of sodium chloride and its prevention 
by other salts. /. Biol. Chem. 1:363. 



Family parameciidae 

SOME METHODS FOR THE PURE CULTURE OF PROTOZOA 

William Trager, Rockefeller Institute for Medical Research 

I. SEPARATION OF THE PROTOZOA FROM 
OTHER MICRO-ORGANISMS 

Migration through Pipettes. The technique developed by Glaser and 
Coria (1930), and used by them for the separation of certain Protozoa 
from their contaminating bacteria, depends essentially on the exhibition 
by the Protozoa of a geotropic response which causes them to swim away 
from the bacteria through a column of sterile liquid. Two methods for 
bringing this about are available. In the first, sterile pipettes are used, at 
least 14 inches long with a % inch bore, a tapering point, and a cotton 
plug at the large end. For negatively geotropic Protozoa such a pipette, 
by suction through a rubber tube attached to the large end, is filled with 
sterile tap water to within 2 inches of the top. Then about 2 cc. of a 
heavy culture of the contaminated Protozoa is carefully sucked up into 
the pipette, so as to form a layer beneath the sterile water. The tapering 
end of the pipette is then sealed by heat, care being taken not to permit 
the formation of air bubbles. The pipette, sealed end down, is set upright 
in a test tube rack. In from 5 to 30 minutes, negatively geotropic Proto- 
zoa will be present at the top of the column of liquid. Even motile 



Parameciidae 113 

bacteria, by their own efforts, can not reach the top in so short a time. 
Usually, however, one such washing does not free the Protozoa from 
adherent or ingested bacteria, and in most cases it is necessary to repeat 
the procedure once or twice, each time sucking up beneath the fresh 
column of sterile water in a fresh pipette about 2 inches of fluid from the 
top of the previous pipette. In some cases it is advisable to leave the first 
or second pipette for 18 to 24 hours to give the surface migrants a chance 
to evacuate the remains of ingested micro-organisms and to multiply to 
some extent. This procedure is followed by another rapid washing. 
Finally a drop of the surface fluid is inoculated into a tube of sterile 
medium (see Part II). In this manner the ciliates Trichoda pur a, three 
strains of the thermal ciliate Saprophilus ovijormis from Hot Springs, 
Virginia, a large undetermined ciliate, Paramecium caudatum and P. 
multimicronucleatum, and the flagellates Chilomonas Paramecium, Para- 
polytoma satura, and an undetermined monad from the intestine of the 
fly, Lucilia caesar, were all freed of bacteria, as shown by consistently 
negative findings in stained films and aerobic and anaerobic cultures on 
routine laboratory media at both room and incubator temperatures. The 
upward migration here involved is a genuine geotropic reaction, as it 
occurs in pipettes held in the dark and in those sealed without any air 
space at the top. 

This same pipette method may be used for positively geotropic Proto- 
zoa. The pipette is nearly filled as before with sterile water, the tip is 
sealed and then a small amount of the contaminated culture is layered on 
top of the sterile liquid. After a suitable time the end of the pipette is 
cut off and one or two drops either inoculated into culture medium or 
placed at the top of another water column for a second washing. In this 
way an undetermined free-living monad was freed of bacteria after two 
washings, each consuming about 30 minutes. With either negatively 
or positively geotropic organisms the addition of killed yeast cells (see 
Part II) to that part of the water column toward which the Protozoa 
were to migrate, accelerated their migration. 

Migration through V-Tubes. In the second method, V-shaped tubes 
are used, one arm of the "V" being 12 cm. long with an inside diameter 
of 28 mm., the other 9 cm. long with an inside diameter of 8 mm. The 
tube, after sterilization, is filled with 15 cc. of melted semi-solid medium 
(see Part II) . When this has set to a soft gel the contaminated Protozoa 
are introduced by means of a long fine capillary through the small arm 
into the bottom of the large one. The tube is permitted to stand for a 
length of time dependent of the Protozoa concerned, and then samples 
are taken from the surface of the medium in the large arm. One such 
treatment frequently suffices. A modification of this technique was used 
to separate Spirillum undulans from other bacteria. A loopful of con- 



H4 Phylum Protozoa 

taminated culture of the Spirillum was placed on the surface of sterile 
tap water in the large arm of a V-tube. Five-tenths of a cc. amounts, 
withdrawn one hour later from the surface of the small arm, gave pure 
cultures of the Spirillum upon inoculation into suitable media (see also 
Part III). 

II. CULTURE MEDIA FOR CERTAIN BACTERIA-FREE PROTOZOA 

"Basic Medium." Among the free-living Protozoa freed of bacteria 
by any of the above methods some, such as Trichoda pura, were able to 
grow well in simple media such as peptone water. Even such Protozoa, 
however, grow better in the so-called "basic medium" of Glaser and 
Coria (1930). This has been recently modified ( Glaser and Coria, 1934a) 
to give more nearly uniform results and is now prepared in the following 
way: Stock Solution A consists of 50 cc. of horse serum in 1000 cc. of 
well water (or distilled water). This is autoclaved for 30 minutes at 
15 lbs. and has a pH of 7.0 without adjustment. Stock solution B consists 
of 50 gms. of timothy hay in 1000 cc. of water. The mixture is infused 
over night in the refrigerator, filtered through cotton, and the reaction 
adjusted to pH 7.2-7.4. The stock solutions A and B are stored 
separately in a refrigerator and when needed are combined with well 
water in these proportions: 

Solution A 500 cc. 
Solution B 250 cc. 
Well water 250 cc. 

The medium is adjusted to pH 7.2-7.4, tubed in 8 or 10 cc. amounts and 
autoclaved. This final medium was used only for Protozoa free of other 
micro-organisms. When heavy initial cultures of unpurified Protozoa 
are desired 2 cc. of the final medium is added to 20 cc. of the original 
Protozoa-containing water. To make a solid medium, 1.5 to 2% agar is 
added. For a semi-solid medium similar to Noguchi's Leptospira 
medium, 100 cc. of the melted solid medium is diluted with 900 cc. of 
warm water. 

Raw Potato. Some Protozoa, which grew well in basic medium when 
contaminated with bacteria, did not grow at all in this medium when 
free of other micro-organisms. Thus a flagellate from the intestine of 
Lucilia caesar would not grow in basic medium except in the presence of 
living bacteria. The bacteria-free flagellate would not develop on auto- 
claved potato, but delicate growths of it were obtained in tubes of raw 
potato under tap water. Such tubes are prepared as follows: Raw po- 
tatoes are scrubbed with hot water and partly dried by heat; a part of 
the surface is washed thoroughly with 70% alcohol and flamed until 
charred. Cylinders are then cut out with a sterile No. 5 cork borer and 



Parameciidae 115 

put in sterile petri dishes where they are cut into pieces % inch long. 
Each piece is placed in a tube of sterile water. Such tubes are held at 
room temperature for some time before use, and any showing con- 
tamination are discarded. 

Special Medium for Paramecia. Two species of Paramecium {P. cauda- 
tum and P. multimicronucleatum) probably furnish the best examples of 
free-living Protozoa which, when free of other micro-organisms, require a 
very special complex nutritive medium (Glaser and Coria, 1933, 1934a). 
Various workers have successfully freed Paramecia of contaminating 
bacteria, usually by means of the washing of isolated individuals, but the 
bacteria-free organisms could not be cultured. Such was at first the 
experience of Glaser and Coria ( 1930) who used the migration technique 
to obtain the pure Protozoa. They established a "pure-mixed" culture 
in basic medium of Paramecium caudatum in association with the yeast, 
Saccharomyces cervisiae. Since the yeast is non-motile and heavier than 
water one can readily take advantage of the negative geotropic migration 
of the Paramecia to secure pure Protozoa. In practise, the "pure-mixed" 
culture was centrifuged and washed three times in sterile water. The 
material so obtained was introduced at the base of a pipette in the 
manner previously described and the Protozoa permitted to migrate 
through the column of sterile water. Every day for 5 days material from 
the top of the pipette was removed and plated on dextrose agar. Only in 
cultures made on the first day were any yeast colonies obtained, showing 
that after the first day the yeast cells brought to the surface by the Para- 
mecia had been digested. Upward migration of the Protozoa was com- 
plete by the fourth day at a temperature of 20 o -2 2° C. Pure Paramecia 
obtained in this way were finally cultured in a medium consisting of liver 
extract, killed yeast, and fresh rabbit kidney. Repeated tests (aerobic 
and anaerobic cultures in a variety of media at room and incubator 
temperatures) showed that the cultures so obtained were free of other 
micro-organisms. 

The liver extract consists of a 0.5% solution of Eli Lilly Company's 
liver extract No. 343 in water. This is filtered through paper and 
sterilized by filtration through a Berkefeld N filter. This extract has a 
pH of 6.2 to 6.4 and is placed in 10 cc. amounts in sterile test tubes. 
Liver extract may also be made in the following way: One hundred grams 
of finely ground rabbit, beef, or swine liver are infused over night in the 
refrigerator in 200 cc. of water. The suspension is filtered through cotton, 
heated over a water bath for about one hour, and then strained through 
fine gauze, all the fluid being squeezed out of the coagulum. The liquid 
is further cleared by centrifuging and is then diluted with water to 400 cc. 
It is warmed to 6o° C. and passed successively through sterile Berke- 
feld V and N candles and then is tubed in 10 cc. amounts. Heat steriliza- 



ZI 6 Phylum Protozoa 

tion of the liver extract, either in the autoclave or in the Arnold sterilizer 
does not give a satisfactory medium. 

Dead yeast is prepared from baker's yeast grown for 5 days on dextrose 
agar in a Blake bottle. The growth is washed off in sterile water, 
centrifuged, and again washed and centrifuged. The washed yeasts are 
suspended in 15 cc. of sterile water and this suspension is distributed in 
5 cc. amounts to tubes which are then sealed and placed for 30 minutes 
in a water bath at a temperature of 75 ° to 8o° C. The heated suspen- 
sions are tested for sterility on dextrose agar slants. Killed cultures of 
Staphylococcus pyogenes aureus or albus may be used instead of the 
killed yeast. 

In the final preparation of the medium, pieces of kidney weighing 
0.2 to 0.5 gms. are aseptically removed from a freshly killed rabbit and 
placed in tubes of liver extract. To each tube is added 0.1 cc. of heat- 
killed yeast suspension and also some Paramecia from the surface of a 
migration pipette. An inoculum of about 870 Paramecia was usually 
used. In such tubes a luxuriant growth of Paramecium caudatum is pres- 
ent by the tenth day and successful subcultures are regularly obtained 
between the seventh and fifteenth day of incubation at room temperatures. 
Table I shows that the liver extract, killed yeast, and fresh kidney are 
all essential to the growth of Paramecium caudatum in the absence of 
other micro-organisms. Very similar results have been obtained with the 
large Paramecium multimicronucleatum. By the ordinary migration 
technique this ciliate was easily freed of all but one contaminant, a small 
motile bacillus. Further migrations were now performed in sterile water 
containing a few drops of living yeast cell suspension. The bacillus 
multiplied very slowly in the water, and the Paramecia fed on the yeast 
cells. These eventually settled out while the Protozoa swam to the top 
of the long water column. Such daily migrations were repeated over a 
period of 6 days and finally a migration in sterile water without yeast 
cells yielded Protozoa free of other micro-organisms. Such Paramecium 
multimicronucleatum grew in the same medium which was used for 
P. caudatum. 

Necessity for a Change of Medium. While both species of Paramecia 
in their special culture medium, and the flagellate Parapolytoma satura on 
blood agar, seem to grow indefinitely, the other purified Protozoa, when 
grown continuously in "basic medium" showed a gradually lessening 
developmental rate and finally died out unless transferred to some other 
culture medium. Thus after several months in "basic medium," with 
transfer intervals of 1 to 2 weeks, when the Protozoa had begun to grow 
less luxuriantly, they were subcultured to potato or carrot water or to 
basic medium containing sterile rabbit kidney or yeast extract. Such 
transfers again gave excellent growth, but this again weakened after 



Parameciidae 



117 



table I — Effect of Liver Extract, Kidney and Yeast on the Growth of 

Paramecium. 



Medium 



Water+kidney*** 

(4 it 

tt tt 

Water+kidney+dead yeast 

tt t* tt tt 

tt tt tt tt 

Water+kidney+ living yeast 

11 tt tt tt 

tt it tt tt 

Lilly's extract+kidney-f dead yeast. 



Lilly's extract+dead yeast 

H tt tt tt 

tt tt it tt 

Liver extract+kidney-f dead yeast. 



tt tt 



Test 

No. 




Degree of growth in daj 


s 














2 


4 


6 


8 


10 


1 


4-*** 


+ 


+ 


+ 


— 


2 


+ 


+ 


+ 


+ 


— 


3 


+ 


+ 


+ 


+ 


— 


4 


+ 


+ 


+ 


+ 


— 


5 


+ 


+ 


+ 


+ 


— 


6 


+ 


+ 


+ 


+ 


— 


7 


+ 


++ 


+++ 


++++ 


++++ 


8 


+ 


++ 


+++ 


++++ 


+++ + 


9 


+ 


++ 


+++ 


++++ 


++++ 


10 


+ 


++ 


+++ 


++++ 


++++ 


11 


+ 


+ + 


+++ 


++++ 


++++ 


12 


+ 


+ + 


+++ 


++++ 


+++ + 


13 


+ 


+ 


+ 


+ 


+ 


14 


+ 


+ 


+ 


+ 


+ 


15 


+ 


+ 


+ 


+ 


+ 


16 


+ 


+ 


+ 


+ 


+ 


17 


+ 


+ 


+ 


+ 


+ 


18 


+ 


+ 


+ 


+ 


+ 


19 


+ 


+ + 


+++ 


++++ 


++++ 


20 


+ 


++ 


+++ 


++++ 


++++ 


21 


+ 


+ + 


+++ 


++++ 


++++ 



Transplant 



Negative 



Positive 



Negative 



Positive 



*** Water=sterile tap water; kidney = fresh rabbit kidney; Lilly's extract = extract prepared from Eli- 
Lilly and Company's liver extract No. 343; liver = rabbit liver extract prepared in this laboratory; — = 
dead; ±.=doubtful growth; +=weak growth: ++=fair growth; +-r-+=good growth; -f-- r -- r -+=lux 
uriant growth. 

(From Glaser and Coria, 1933) 

several months in the new medium, and the Protozoa were then returned 
to "basic medium" and the whole procedure repeated. Potato or carrot 
water is prepared by autoclaving 2 gram pieces of the vegetable in tubes 
with 10 cc. of water. For use, 2 cc. of the liquid so obtained is diluted 
with 10 cc. of sterile water. Yeast extract is prepared in the following 
way. Pure cultures of yeast are grown on dextrose agar in Blake bottles 
for 5 to 7 days at room temperature. The growth is washed and centri- 
fuged three times with 50 cc. of sterile water and 2 cc. of the sedimented 
cells are then ground for 5 hours in a sterile mechanical grinder consisting 
of two ground glass tubes one of which, the pestle, fits snugly within the 
other and is turned by an air turbine motor (see Glaser and Coria 1934a 
for details). Thirty cc. of sterile water are then added and the mixture 
stirred for another 20 minutes. This ground yeast suspension is centri- 
fuged 1 5 minutes to remove the larger particles and is then sterilized com- 
pletely (i.e. freed of intact yeast cells) by filtration through a sterile 
Berkefeld V filter. Small amounts of such a yeast extract when added 
to any of the media greatly stimulated the growth of the Protozoa. 
Extracts filtered through Berkefeld N filters or prepared from yeast killed 
chemically or by heat did not stimulate. It should be noted that even 
the best yeast extracts could not be substituted for the intact heat killed 
yeast cells in the special medium devised for the Paramecia. 



n8 Phylum Protozoa 

III. PARTIAL PURIFICATION OF CULTURES OF 
BALANTIDIUM COLI 

It has long been realized that bacteria-free cultures of the intestinal 
protozoan parasites of vertebrates would be extremely useful for the 
study of the biology and pathogenicity of these organisms. Yet up to the 
present time no such protozoan has been grown in culture free of bacteria 
and in most cases it has been impossible even to free the Protozoa of the 
numerous bacteria which naturally accompany them. Cleveland (1928), 
however, succeeded in freeing the coprozoic organism, Tritrichomonas 
fecalis of man, from bacteria and was able to grow it on heat-killed 
bacteria. Glaser and Coria (1934) have recently applied one of their 
migration techniques to Balantidium coli from swine and have been able 
to free the organism of all bacteria except Bacillus coli. Largely as a 
result of this partial purification, they have been able to maintain this 
parasitic ciliate in better condition and for a much longer time without 
subculturing than has heretofore been possible. 

They used a liquid medium consisting of 9 cc. of sterile Ringer's 
(Schumaker, 193 1) solution, 0.5 cc. of sterile horse serum, and a sprinkle 
of rice starch in each tube. Much better results were obtained with a 
semi-solid medium. This is prepared by adding 25 cc. of 2% standard 
nutrient agar, 1 gram of sterile rice starch and 12.5 cc. of sterile horse 
serum to 250 cc. of sterile Ringer's solution warmed to 50 C. The pH 
of the mixture is adjusted to 7.2 to 7.4 and 15 cc. amounts are transferred 
aseptically to sterile tubes. 

"V" tubes containing 15 cc. of semi-solid medium were inoculated 
(after being warmed to 37 C.) on the surface of one arm with material 
scraped from crypts in the mucosa near the ilio-cecal valve of a pig. 
The Balantidia migrated downwards within 24 to 48 hours at 37 C. and 
could then be recovered in a sterile pipette inserted through the uninoc- 
ulated arm of the "V" tube. The semi-solid nature of the medium checked 
the spread of bacteria, which in a liquid medium would rapidly grow over 
both arms of the "V" tube. The Balantidia were able to push their way 
through the gelatinous semi-solid medium and it was found that in such 
a medium a much higher percentage of positive initial cultures could be 
obtained than in a liquid medium. Balantidia removed after the final 
migration were again placed on the surface of one arm of a fresh "V" 
tube and again permitted to migrate for 24 to 48 hours. This procedure 
was repeated five times, after which the ciliates were further cultured in 
ordinary tubes of semi-solid medium. Strains which would not originally 
grow in the liquid medium could be adapted to this after varying lengths 
of culture in the semi-solid medium. 

These methods failed to free Balantidium of Bacillus coli but they did 
yield cultures which might be regularly transplanted at 8-day intervals 



Parameciidae 119 

and one strain (in liquid medium) which was transferred every 20 days 
thrived for over 2% years. 

Bibliography 

Glaser, R. W., and Coria, N. A. 1930. Methods for the pure culture of certain 
Protozoa. J. Exper. Med. 51:787. 

■ — ■ ■ 1933. The culture of Paramecium caudatum free from living microorgan- 
isms. J. Paras. 20:33. 

1934a. The culture and reactions of purified Protozoa. Amer. J. Hyg. 21:111. 

— — ■ — ■ 1934b. The partial purification of Balantidium coli from swine. J. Paras. 



21:190. 
Cleveland, L. R. 1928. The suitability of various bacteria, molds, yeasts and 

spirochaetes as food for the flagellate Tritrichomonas fecalis of man, etc. Amer. 

J. Hyg. 8:990. 
Schumaker, E. i 93 i. The cultivation of Balantidium coli. Ibid. 13:281. 

THYROID CULTURES OF PARAMECIA* 

AT VARIOUS times the writer has had occasion to make some thyroid 
l\ cultures of Paramecium caudatum. It was noted that these animals 
found this habitat more favorable to existence and reproduction than the 
ordinary hay infusions. 

Such a Paramecium thyroid culture may easily be made by mixing 
about 2 grams of Armour's Desiccated Sheep Thyroids (U.S.P.) with 
2,500 cc. spring water. This mixture should be slightly stirred and al- 
lowed to stand exposed to the air for half an hour. Several pipettes full 
of fluid containing Paramecia are then introduced. If the culture jar 
is covered with a top and carefully sealed with vaseline an excellent, 
clear culture will be obtained. After several days it may be noticed that 
the animals are evenly distributed throughout the liquid, and are not 
congested about the top of the jar as in ordinary cultures. The cultures 
usually need but little attention. However, it is sometimes found desir- 
able to add a little fresh water every week or ten days. 

M. E. D. 

A COMBINED CULTURE METHOD AND INDICATOR FOR 

PARAMECIUM** 

BRAGG AND HULPIEU (1925) describe the effect of a stain obtained 
from red cabbage leaves as an indicator of the acidity of the food 
vacuoles of Paramecium. I have been unable to secure similar satis- 
factory results with the races I am using, but have found that a dilute 
infusion of red cabbage leaves (about 30 grams to 1 liter of water) is an 
excellent medium in which the animals reproduce rapidly; at the same 

* Reprinted with slight changes from an article in Science 58:205, 1923, by William L. 
Straus, Jr. 

**Reprinted with slight changes from an article in Science 62:351, 1925, by Robert T. 
Hance, University of Pittsburgh. 



I2 o Phylum Protozoa 

time the color of the infusion indicates the chemical condition of the cul- 
ture. When fresh, the cabbage leaf culture medium is light reddish 
purple in color, but about 24 hours after being seeded with Paramecium, 
it turns red, indicating the formation of acid. In four or five days to two 
weeks, as the Paramecia increase in number, the medium gradually be- 
comes alkaline, as is shown by its change of color to green. 

The culture, as far as quantity of Paramecia is concerned, is at its 
height when it becomes a brilliant green and has lost its early turbidity. 
The behavior of the cultures may be varied considerably by adding a trace 
of sodium bicarbonate or a weak acid. In from one to two months, the 
culture becomes the color of an old hay infusion, fails to react to acids or 
alkalies and the Paramecia have either wholly or almost wholly died off. 

Bibliography 
Bragg, A. N., and Htjlpieu, H. 1925. A method of demonstrating acidity of 

food vacuoles in Paramecium. Science 61:392. 

M. E. D. 

PARAMECIUM* 
William LeRay and Norma Ford, University of Toronto 

PARAMECIUM is a form which is easily reared. From the bottom of 
a permanent pond the foul-smelling debris is taken and kept in a 
bowl barely covered with water and at a temperature of approximately 
73 F. As the debris fouls, the Paramecium become abundant. From 
time to time (about once a week) a half-inch cube of fish is added to 
the bowl to maintain a supply of food. Such a culture as this will carry on 
for months. It is advisable to select a large race and to rear in sepa- 
rate containers. Should some small forms be present in the new bowls 
they are usually unsuccessful in the competition with the large ones. 

A CULTURE MEDIUM FOR PARAMECIUM** 

THIS medium has proven very satisfactory for culturing various 
species of Paramecium in pure line cultures. The main result of the 
use of the medium is that the organisms do not exhibit a lowering of their 
normal metabolism after continuous culturing. 

* Editor's Note: For the technique used by Prof. L. L. Woodruff of Yale University in 
maintaining his famous pedigreed cultures of Paramecia the reader is referred to the 
following articles: 

Woodruff, L. L. 1908. The life cycle of Paramecium when subjected to a varied 
environment. Amer. Nat. 42:520. 

1911. Two thousand generations of Paramecium. Archiv. f. Protist. 21:263. 

1932. Paramecium aurelia in pedigreed culture for twenty-five years. Trans. Amer. 

Micr. Soc. 51:196. 

Woodruff, L. L., and Erdmann, Rhoda. 1914- A normal periodic reorganization 
process without cell fusion in Paramecium. /. Exper. Zool. 17:425- 

♦♦Reprinted, with slight changes, from Science 75:364. 1932, by Lauren E. 
Rosenberg, University of California. 



Parameciidae 121 

The basic part of the medium is the usual hay infusion of 10 grams of 
chopped timothy hay boiled for 15 minutes in a liter of well water. This 
infusion is filtered and sterilized in the Arnold sterilizer at ioo° C. one 
hour a day for 3 days. It is diluted with 9 volumes of sterile well water 
just before using. Two portions of this infusion are placed in sterile 
liter flasks with sterile cotton stoppers. One liter is inoculated with 
Bacillus subtilus and the second with B. coli communis. A third portion 
of the medium is made up as follows: Approximately 30 grains of wheat 
are boiled in a small amount of water for 10 minutes. The wheat grains 
only are then placed in a third liter flask of sterile well water. The three 
portions are incubated at 37 C. for 24 hours, and then combined in 
one large sterile flask. The medium is now ready for use. The culture 
may be used in almost any size of container, but that used has been the 
300 cc. Erlenmeyer flask. These flasks are fitted with cotton stoppers 
and sterilized. Each flask is filled about % full of the medium, and 
different species are transferred to the cultures with sterile pipettes. 

The original basic infusion may be made up, sterilized, and stored in a 
refrigerator until ready for use. Likewise, the medium, made of the three 
portions, may be stored in a refrigerator for later use. 

Sterile precautions are maintained throughout the procedure, but after 
Paramecium has been transplanted, such strict precautions are no longer 
necessary. The essential part of the process is to provide a medium 
rich with a suitable food in which Paramecium will continue to grow 
normally. With these sterile precautions, other ciliates and flagellates 
are eliminated. A single organism placed in such a medium will produce 
a flourishing culture in 7 to 10 days. One should transplant every 2 to 
3 weeks. Paramecium multimicronucleatum, P. bursaria, and P. aurelia 
have thrived in this medium. 

M. E. D. 

Reference 

For the culture of Paramecium see also pp. 62, 63, 72, 134, 136, and 177. 

CULTIVATION OF PARAMECIUM AURELIA AND 
P. MULTIMICRONUCLEATUM 

T. M. Sonneborn, Johns Hopkins University 

OF THE many media tried, the most successful is an infusion of 
1.5 gms. of desiccated (but not burned), powdered lettuce boiled 
for 3 minutes in a liter of double distilled water. This infusion is filtered 
while hot, dispensed into pyrex test tubes or flasks, containing an excess 
of pure CaC03 (which adjusts the pH to about 7.2 ), plugged with cotton, 
and autoclaved. When ready for use, it is filtered to eliminate the 
CaCOs and is inoculated with the bacterium, Flavobactcrium brunneum, 



122 Phylum Protozoa 

grown on beef-agar slants, and the alga Stichococcus bacillaris, grown on 
0.05% Benecke's agar slants. The bacterial slants may be used when 
i to 5 days old; the algal slants when 1 8 to 24 days old. The quantities 
inoculated into the culture fluid are one i-mm. loop level full of bac- 
teria and three 2-mm. loops of algae to 20 cc. culture fluid. 

Isolation cultures may be carried on depression slides by using two 
drops of this fluid per depression. Such cultures must be renewed daily 
by transferring one Paramecium to freshly inoculated fluid on a fresh 
slide. The optimum temperature is 2 7°-28° C. 

Mass cultures may be carried in cotton-stoppered flasks by using the 
same fluid. Such cultures must be made frequently to keep the Para- 
mecia in good condition, but the organisms will live for three months or 
more in a greatly depressed condition without renewal of fluid. 

References 

For the culture of Paramecium aurelia see also pp. 105 and 120. 

For the culture of Paramecium multimicronucleatum see also pp. 113, 115 and 128. 

For the culture of Paramecium bursaria see pp. 55 and 105. 

PARAMECIUM MULTIMICRONUCLEATUM; MASS- 

CULTURING, MAINTAINING AND REHABILITATING 

MASS-CULTURES, AND SECURING CONCENTRATIONS 

Edgar P. Jones, University of Akron and University of Pittsburgh 

MASS-CULTURING 

VARIETY in composition is one of the outstanding characteristics of 
infusions in which Paramecia are to be mass-cultured. There ap- 
pears to be no single method of successful culturing. In general, small 
quantities of materials, usually organic, must be introduced into water to 
induce bacterial multiplication. These bacteria are the chief food supply 
of the Paramecia, at least in the earlier stages of the culture. Liebig's 
beef extract (Woodruff and Baitsell, 191 1), mangle beet water (Glaser 
and Coria, 1930), sewage (Butterfield, Purdy, and Theriault, 1931), let- 
tuce leaves (Dimitrowa, 1930), wheat (Turtox Leaflet No. 4), bananas 
(Turtox Leaflet No. 4), timothy hay (Petersen, 1929), Horlick's malted 
milk (Jennings and Lashley, 1914), gelatin or curd placed under earth, 
meat, pond lily leaves, red cabbage leaves, and soil are some of the 
materials which are reported to have been used. Such diverse organic 
materials may be boiled in water, or they may be allowed to macerate. 
When the infusion is ready, it may be autoclaved, especially if portions 
are to be preserved for future use; or it may be used without sterilization. 
Slight variations of the above techniques require the introduction of 
algae (Raffel, 1930), yeast (Lund, 1918), or bacteria (Phelps, 1934) 



Parameciidae 123 

into infusions such as the above. Certain investigators adjust osmotic 
pressures, pH, or other ionic concentrations by adding salts or other 
compounds. Such techniques are not essential to the production of ex- 
cellent mass-cultures of Paramecia, although they will enhance the uni- 
formity of the medium. 

The present discussion is limited to cultures of P. multimicronucleatum 
grown in infusions prepared by boiling hay or hay-flour combinations in 
water. The following is a formula which I have employed frequently 
and successfully: 

1 gram hay 
0.1 grams white flour 
700 cc. distilled water 

Stir the flour through the hay. Bring the water to a boil. Add the hay-flour 
mixture. Boil 10 minutes. Cool. Add distilled water to replace that evaporated. 
Seed with 200 Paramecia on the second day. 

The above formula may be considerably altered, especially in routine 
work, without materially interfering with the production of a satisfactory 
population. Permissible variations include the use of hay up to 4 grams 
if flour be omitted (Jones, 1930) ; the use of tap or pond water if non- 
toxic; and variation in the date of seeding and the number of seed Para- 
mecia introduced. 

Additions of hay or flour in excess of the amounts stated will produce 
a hydrogen-ion concentration which will destroy the Paramecia on the 
fourth or fifth day (Jones, 1930) (pH 4.83 or less) . The "seeding" must, 
under such circumstances be delayed until the pH exceeds 5.0. In the 
presence of such excesses of food, certain other unidentified split products 
which result may prove to be toxic, even if the pH be satisfactory. In- 
troduction of excesses of food material must therefore be avoided, either 
when originally preparing the infusion, or later, if feeding techniques be 
employed. 

Cultures should not be covered if populations of maximum concentra- 
tion are desired. Non-evaporating cultures made as described above will 
produce populations of approximately 300 Paramecia per cc. of solution 
as a maximum, whereas evaporating cultures may eventually yield from 
2000 to 4000 per cc. 

MAINTAINING AND REHABILITATING MASS-CULTURES 

Mass-cultures, especially of the non-evaporating type (in which the 
cover is placed on the jar, but not screwed down) , may be revived by feed- 
ing when they approach the point in the cycle of culture conditions at 
which the Paramecia would normally disappear. When a population 
of Paramecia is present in a culture it may frequently be increased by 
the same means. At various times I have with success employed flour, 



124 Phylum Protozoa 

fresh hay infusion, egg, chocolate, peptone solution, or milk as supple- 
mental foods (Jones, 1933). Any simple method of introducing such 
substances is usually satisfactory if the materials are in a finely divided 
state or a fluid condition. Flour is introduced by sprinkling approxi- 
mately y 2 gram upon the surface of 700 cc. of infusion in as unlumped a 
condition as possible, and stirring. A solution of egg may be prepared 
by breaking an egg into 100 cc. of distilled water in a flask, and shaking 
with glass beads. Sterile technique may be employed, but I have not 
observed that it materially alters the end result. 

Other investigators report temporary revival of declining cultures 
secured by introducing sugar (McClendon, 1909), bread (Bauer, 1926), 
or fresh hay infusion (Kudo, 193 1) ; and the culture has been maintained 
for a year, in one instance (Kudo, 1934) by occasionally introducing a 
few grains of wheat and a few pieces of timothy hay. 

The degree of success which may be had in maintaining mass-cultures 
by feeding is indicated by the following preliminary report of the first 
which I so tested. This was a 700 cc. culture, loosely covered, made in 
accordance with the formula furnished above. Controls were initiated 
in similar fashion. 

In the controls, which developed and declined in the usual mass- 
culture fashion, Paramecia could be found for a period of approximately 
six months. Dense populations were not present after the initial three 
months. The fed culture maintained Paramecia for more than twenty 
months. It produced a dense population when fed when fourteen months 
old, although the Paramecium population had dropped almost to zero 
while the culture had gone unfed over a summer's vacation which had 
just preceded. The Paramecia eventually disappeared for no known 
reason, other than that the culture had not been fed for some time. 

To further test the potential longevity of Paramecium cultures, four- 
teen cultures of a capacity of one gallon were prepared. These received 
infusion of approximately the same composition, and seed in the same 
ration per volume as described above. They were seeded November 12, 
1932. 

It became necessary to feed flour to all of these gallon cultures before 
they were six months old, because, in every culture the Paramecium 
population had declined almost to extinction. Six cultures now survive 
(June, 1935). These cultures have been fed repeatedly, usually only 
when the Paramecium populations were quite low, whereupon they have 
characteristically produced larger populations. Flour has been most 
frequently fed, but milk, glucose, albumin, and ethyl alcohol have been 
used at times. An epidemic of mold which destroyed the Paramecia 
appeared in practically every culture to which ethyl alcohol was fed. 

At the present time, the six of the series which now retain populations 



Parameciidae 



125 



27 



32 



37 



42 



PLATE DS~5b 

47 52 57 62 



67 



72 



77 



760 

700 
660 



400 




250 
200 
150 
100- 



50- 
35 



400 



200 



VOLUME OF CULT URL 
IN CCS 




s 



,H 



n>OD ADDED 'E 9 f IN CC$ ? 



TTTTT 



POPULATION PER C.C 





AVERAGE LENGTH ° 
IN MICRA » 




o u o 



> o" °o° °° 



AVERAGE WIDTH 
IN MICRA 



TURBIDITY TEN MINUTE5 
AFTER STIRRING 



r^VV2i 





^^v 




TURBIDTTY BEFORE STIRRING^ 

Tcoj 
CLEAR JK, CLEAR, 



AVERAGE VOLUME OF ONE 
ANIMAL IN (itifyV- 



^ 'VOLUME OF RR$T0PLA5M 



27 32 37 42 47 52 57 62 

A6E IN MY5 



67 72 



400 



200 



a 



TURBIDITY 
5 

I? 

1$ 

20 



77 



Fig. 44. — Record of the decline of a Paramecium population as well as its later rehabilita- 
tion after feeding egg. The abscissa represents time in days, beginning with the 27th 
day. The Paramecium population was near its maximum. The ordinate presents the 
following data: volume of infusion; pH; dates when egg was added, with the approximate 
amounts; concentration of Paramecia/cc; average lengths and widths of 10 living animals, 
with maxima and minima indicated; turbidity of the culture before stirring (To), and 
10 minutes after stirring (Tio) ; the average volume of single animals in thousands of 
cubic micra, and the average volume of Paramecium protoplasm maintained per cc. of 
infusion in millions of cubic micra. 



126 Phylum Protozoa 

of Paramecia are 30 months of age. To this group I have recently fed 
flour at approximately weekly intervals to determine whether dense 
populations can still be produced. Four of the cultures now contain 
an average concentration of 300 to 400 per cc. Such concentrations are 
high for large cultures which are not evaporating. To the best of my 
knowledge, none of these cultures has at any time contained a higher con- 
centration. 

Reductions in length and number of Paramecia which are character- 
istic of the declining culture are graphed in Fig. 44. The corresponding 
increases which resulted when egg was fed are likewise indicated. 

SECURING CONCENTRATIONS 

These methods have been employed for concentrating either ro- 
tifers or paramecia (Jones, 1932). There is reason to believe that a 
considerable variety of protozoans and small metazoans will respond in 
like fashion. 

Concentrations not needed for two days. Distribute cultures which 
contain Paramecia among containers having the approximate dimensions 
of quart fruit jars, filling each container half full. Add to each enough 
freshly prepared, cooled infusion, made according to the formula given 
above (variation is permissible), to fill it completely. Do not cover. 
Within 40 to 60 hours the Paramecia will congregate on the sides of 
the container at or immediately below the surface. Remove the con- 
centration with a pipette having a finely drawn tip and a bulb. 

Concentrations for immediate use. Such concentrations may usually 
be picked up directly from the bottom of an older culture, if a longer 
pipette is employed. The Paramecia secured by such methods are usu- 
ally smaller than those concentrated by the first method. 

Debris-free concentrations. Paramecia may be freed from the culture 
debris by introducing into concentration tubes the animals and infusion 
taken by either of the preceding methods. For this purpose I have em- 
ployed glass tubes which were 30 cm. long and which had an internal 
diameter of 8 mm. The organisms, following introduction, will settle 
to the bottom, after which they will systematically migrate to the sur- 
face, from which position they may be removed with a pipette. 

Bibliography 

Bauer, Frederick. 1926. Science 64:362. 

Butterfleld, C. T., Purdy, W. C, and Theriault, E. J. 1931. Pub. Health 

Repts. 46:393- 
Dimitrowa, Ariadne. 1930. Arch. f. Protist. 72:554. 
Glaser, R. W., and Coria, N. A. 1930. /. Exper. Med. 51 : 7S7. 
Jennings, H. S., Lashley, K. S. 1914. J. Exper. Zool. 14:393. 
Jones, Edgar P. 1930. Biol. Bull. 59:275. 
1932. Science 75:52. 



Parameciidae 127 

1933. Univ. of Pittsburgh Ball. 29:141. 
Kudo, R. R. 1931. Handbook of Protozoology. Springfield, 111. C. C. Thomas. 

1934- Turtox News 12:127. 

Lund, Barbara L. 1918. Biol. Bull. 35:211. 

McClendon, J. F. 1909. J. Exper. Zool. 6:265. 

Petersen, W. A. 1929. Physiol. Zool. 2:221. 

Phelps, Austin. 1934. Arch. f. Protist. 82:134. 

Raffel, Daniel. 1930. Biol. Bull. 58:293. 

Turtox Service Leaflet No. 4, General Biological Supply House, Chicago, 111. 

Woodruff, L. L., and Baitsell, G. A. 1911. /. Exper. Zool. 11:135. 



CULTURING PARAMECIUM CAUDATUM IN OAT STRAW 

INFUSION 

George A. Smith, Eugenics Record Office 

AN EXCELLENT medium for culturing Paramecium caudatum for 
JL\ a rapid population growth is an oat straw infusion. After having 
experimented with various types and concentrations of culture media 
made from timothy hay, oat straw, barley straw, oak leaves, elm leaves, 
clover, alfalfa, etc., oat straw was found to be the most favorable medium 
for rapid growth of my strain of Paramecium. 

Cut 15 grams of oat straw into short lengths (1-3 cm. long) and place 
in a quart glass container. Pour 900 cc. of boiling distilled water over 
the straw. Plug the container with cotton and allow the mixture to cool. 
Adjust the pH to 7.8 with NaOH. The colorimetric method is sufficiently 
accurate. Keep the mixture at approximately 25 C. temperature for 
48 hours. Shake the culture medium until it is thoroughly mixed; again 
adjust the pH to 7.8 and the infusion is ready to use. Add approximately 
250 Paramecia and in a few days a mass-culture should have developed. 

For best results make new culture medium every 48 hours and make 
new inoculations as often, because usually after 72 hours have elapsed 
the culture medium begins to deteriorate and is not at its best for op- 
timum growth. 

By following the above procedure a number of times a colony of 
rapidly dividing Paramecium caudatum can be developed, each animal 
dividing at an average rate of once every 8 hours. This is considered 
optimum growth under these conditions. 

For culturing animals for classroom use put two or three dozen grains 
of oats in 1,000 cc. of water and allow the mixture to stand for three 
days before inoculating with Paramecia. Within a week a mass-culture 
of the animals usually develops. It is best to keep the culture covered 
when it is not in use. 

Reference 

For the culture of Paramecium caudatum see also pp. 56, 105, 113, and 119. 



128 Phylum Protozoa 

A CULTURE METHOD FOR PARAMECIUM MULTIMI- 
CRONUCLEATUM AND OXYTRICHA FALLAX 

A. C. Geese, Stanford University 

BOTH of these may be grown in 0.1% lettuce infusion. The lettuce 
for this infusion is obtained by drying lettuce leaves in an oven and 
pulverizing. The proper weight of lettuce is boiled in 0.005 M. KH 2 P0 4 
solution for 3 minutes, after which the infusion is titrated to a pH of 7.0 
with NaOH. The particles of lettuce may be left in or removed. The 
culture lasts longer if the particles remain. Usually 15 cc. of such an 
infusion was seeded with 20 to 100 Paramecia or Oxytrichae. It is quite 
satisfactory for rough work to use tap water instead of the buffer solu- 
tion. 

Order heterotrichida 
Family plagiotomidae 

THE CULTIVATION OF NYCTOTHERUS OVALIS AND 
ENDAMOEBA BLATTAE* 

Nyctotherus ovalis from the hindgut of the cockroach, Blattella 
germanica, may easily be cultured in a modified Smith and Barret ( 1928) 
medium. This medium was used by the discoverers for Endamoeba 
(Entamoeba) thomsoni, and according to Lucas (1928) it is suitable for 
the cultivation of neither Endamoeba blattae nor N. ovalis. The medium 
used by Smith and Barret consists of 19 parts of 0.5% NaCl to 1 part of 
inactivated human blood serum. By substituting non-inactivated rabbit 
serum for the human serum a medium is produced in which N. ovalis lives 
and multiplies freely. Dividing forms are common, and occasionally 
precystic and cystic forms are met with. Three cultures have been main- 
tained for 40 days and at the last examination the organisms were as 
normal in appearance as those found in their native habitat. Sub- 
culturing is done at weekly intervals, and the cultures are maintained at 
room temperature. 

The cultivation of E. blattae has been less successful than that of 
N. ovalis. Two cultures out of twelve attempts were maintained for 29 
days. At the end of this time the organisms were few in number but 
entirely normal in appearance and movement. One 2- and one 8-nucleate 
form were seen, the latter with nuclei of different sizes and evidently pre- 
cystic. The next examination was negative. This gradual dwindling in 
number does not necessarily indicate an unfavorable environment, but 

* Reprinted with slight changes from an article in Science 76:237, 1932, by Harry E. 
Balch, University of California. 



Bursariidae 129 

rather that division is not frequent enough to permit weekly subcultur- 
ing without gradually diminishing the number of organisms to the point 
of extinction. Longer intervals between subcultures result in an over- 
growth of bacteria and the small flagellate Monocercomonas orthopter- 

orum. 

References 

For the culture of Blepharisma see p. 60. 

For the culture of Spirostomum ambiguum and 5. teres see p. 60. 

For the culture of Nyctotherus cordijormis see p. 69. 

Bibliography 
Lucas, C. L. T. 1928. A study of excystation in Nyctotherus ovalis with notes 

on other intestinal Protozoa of the cockroach. J. Paras. 14:272. 
Smith, N. M., and Barret, H. P. 1928. The cultivation of a parasitic Anioeba 
from the cockroach. Ibid. 14:161. 

M. E. D. 



Family Bursariidae 



A METHOD FOR CULTURING BURSARIA TRUNCATELLA* 

Amos B. K. Penn, Tsing Hua University, Peiping, China 

ADD 1 gram of timothy hay, 1 gram of rye, and 5 grams of fresh cab- 
l\ bage to 600 cc. of spring water. Boil slowly for 5 minutes. Let 
stand uncovered for 2 days to allow development of bacteria, then re- 
move the cabbage, add 500 cc. spring water, transfer 250 cc. of the solu- 
tion with corresponding amounts of hay and rye to each of several % 
liter jars and inoculate with Paramecium, Colpidium, and Chilomonas. 
After 2 or 3 days, i.e., when these organisms have become abundant, 
inoculate with Bursaria. Cover the jars and keep them in indirect sun- 
light at room temperature. 

If a film of gummy substance has developed on the surface of the in- 
fusion, break it. If no gummy substance is present, add some from an 
old culture. 

A culture thus prepared reaches a flourishing condition (iohh individ- 
uals per cc.) in 2 or 3 days, and continues in this condition for 3 or 
4 days. If the infusion is more dilute, the cultures flourish longer, but 
the Bursaria does not become so abundant. 

References 
For the culture of Bursaria truncatella see also p. 103. 
For the culture of Balantidium coli see p. 118. 
Family Stentoridae 

For the culture of Stentor see pp. 64 and 134. 
For the culture of Stentor coeruleus see pp. 60 and 103. 
Order Oligotrichida 

* Reprinted from Anat. Rec. 54:99, 1932, at suggestion of the author. 



!3o Phylum Protozoa 

Family Halteriidae 
For the culture of Halteria see pp. ioo and 177. 

Order hypotrichida 
Family oxytrichidae 

UROLEPTUS MOBILIS* 

THIS organism, appearing in considerable numbers in an old hay 
infusion that had been standing for several months, was successfully 
cultivated and abundant material for study of all the important phases 
of the life history was secured. 

After attempts to cultivate Uroleptus on fresh hay infusion failed, this 
medium was discarded and boiled flour water, 24 hours old, was sub- 
stituted. To make this, 150 mg. of white flour is boiled for 10 minutes 
in 100 cc. of spring water and allowed to stand exposed to the air for 
24 hours. 

With this medium it was found that the organisms would live and 
would divide about once in three days. Later, a more satisfactory me- 
dium was obtained by mixing 2 parts of the flour water, 2 parts of spring 
water, and 1 part of old hay infusion. This improved medium was used 
for nearly 3 months, the individuals dividing approximately once a 
day. Finally a still better medium was obtained by boiling 100 mg. of 
chopped hay with 130 mg. of flour in 100 cc. of spring water for 10 
minutes and diluting this, when 24 hours old, with an equal part of fresh 
spring water. With this medium made fresh every day, the organisms 
divide from one to three times per day. 

As in previous culture work, a single individual is transferred to about 
200 mg. of the culture medium contained in a flat, 40-mm. square, ground 
glass, hollowed dish, 8 mm. in thickness. On the following day the 
number of individuals is counted and a single individual from these is 
then isolated and transferred to fresh culture medium made the day be- 
fore. After an individual is transferred to fresh medium, the remaining 
individuals are placed in a Syracuse dish containing about 10 cc. of the 
fresh culture medium. Here they multiply in large numbers, consti- 
tuting the "stock" material, the source of dividing and conjugating 
forms. 

M. E. D. 
References 

For the culture of Oxytricha see p. 64. 

For the culture of Oxytricha jallax see p. 128. 

For the culture of Stylonychia see p. 64. 

♦Abstracted from a paper in /. Exper. Zool. 27:293, 1919, by Gary N. Calkins, 
Columbia University. 



Oxytrichidae 131 

METHODS FOR CULTURING PLEUROTRICHA* 

Amos B. K. Penn, Tsing Hua University, Peiping, China 

Pleurotricha may be cultured in a hay-rye infusion with Colpidium as 
food or in a physiological medium with Chlorogonium as food. For 
experimental work, the latter method is preferred. 

A 0.2% hay and 0.2% rye infusion is prepared by boiling in a beaker 
for 8 minutes 1 gm. of hay and 1 gm. of rye in 600 cc. of spring water. 
After it has been boiled and cooled, there are about 500 cc. of solution. 
Then half of the rye grains are removed, leaving the other half with all 
the hay in the solution. This is then transferred to a battery jar of 1 liter 
capacity and left for two days, in order to allow bacteria to grow. When 
the infusion is 2 days old and contains many bacteria, it is inoculated 
with Colpidium (or Chilomonas). After 24 hours, there are numerous 
Colpidia present in the infusion. This is then inoculated with Pleuro- 
tricha. From time to time rich cultures of Colpidia raised separately 
[See also p. 51.] are added to the jar as additional food supply. 

When doing physiological work where bacteria and organic matter 
are to be avoided, a physiological medium consisting of all inorganic 
salts may be prepared according to the formula given below: 

CaCb 0.0008 N 

NaN03 0.0003 N 

MgSO.i 0.0002 N 

K0HPO4 0.0001 N 

KH2PO4 0.0001 N 

NH4NO3 0.0008 N 

In culturing Pleurotricha, Boveri dishes of 50 cm. capacity provided 
with covers may be used. Place 20 cm. of this medium in each Boveri 
dish. Add to each dish one pipette of concentrated culture of Chloro- 
gonium, cultivated separately with the same medium. Then transfer 
one or several pleurotrichs into each dish. Cover and place the cultures 
in the bright part of the room. Pleurotrichs so cultivated are large and 
uniform, morphologically and physiologically. They divide four times 
a day. With this high rate of fission, a single individual may give rise 
to several hundred individuals in a few days. 

References 

For the culture of Pleurotricha lanceolata see p. 107. 
Family Euplotidae 

For the culture of Euplotes see p. 63. 

For the culture of Euplotes patella see p. 60. 

For the culture of other hypotrichs see p. 136. 

* See also Arch. j. Protist. Vol. 84, 1934, and Science 80:316, 1934. 



132 Phylum Protozoa 

STYLONETHES STERKII* 

BOTH protective and reproductive cysts of 5. sterkii remain viable 
when dried. This discovery made possible a transfer of the new 
hypotrich from Plymouth, England, to Stanford University, where the 
strain was continued. 

The following wheat infusion method of culturing was used exclusively. 
Twenty grains of wheat were cracked and then boiled in 15 cc. of glass- 
distilled water for from 3 to 5 minutes. The fluid containing numerous 
starch grains was used immediately after cooling and was transferred 
to the culture by means of a pipette having a bore of 1.5 mm. 

Experiments have shown that thriving cultures are most easily main- 
tained when complete evaporation of the medium takes place at inter- 
vals. Consequently, the organisms were grown in watch glasses holding 
conveniently about 4 cc. of fluid. These were ordinarily enclosed in 
petri dishes to prevent evaporation and to facilitate handling, but when 
mass encystment and complete evaporation of the medium was desired, 
the cover was removed. Or, the cover may be partly removed so that it 
protects the watch glass from dust but leaves a wide open gap between 
the two dishes. Within 8-10 hours, at a temperature varying from 15- 
22 C, the cultures were completely dried out. 

New cultures were started daily when free-swimming individuals 
were wanted for study. By means of a mouth pipette 20 organisms 
from a thriving culture were transferred to a watch glass containing 
4 cc. of tap water and a drop of fresh wheat infusion. A drop of infusion 
was added daily to old cultures until protective cysts began to appear 
(2-4 days). Then they were allowed to dry out. Thus it may be 
arranged that there are always on hand about as many new cultures as 
old ones, and a reserve supply of dry cysts. If the study of active or- 
ganisms is to be suspended for a few days or weeks, it is safe to rely upon 
the stock of dry cysts to begin new cultures, as was proved by the trans- 
fer of cysts in watch glasses from England to California. These cysts 
were about 3 weeks old when they arrived. Excystment occurred in 3 to 
4 hours after distilled water or tap water had been added. 

M. E. D. 

Order peritrichida, Family vorticellidae 

A METHOD FOR INDUCING CONJUGATION WITHIN 
VORTICELLA CULTURES 

Harold E. Finley, West Virginia State College 

Materials: Columbia culture dishes, depression slides, culture tubes, 

* Abstracted from a paper in /. Mar. Biol. Assoc. 19:707, 1934, by Laura Garnjobst, 
Stanford University. 



V orticellidae 133 

moist chambers, platinum loop, non-absorbent cotton, alfalfa hay, wheat 
kernels, glass-distilled water, spring water, La Motte buffer mixtures of 
known pH, agar-slant cultures of the bacterium Achromobacter liquefa- 
ciens. From the materials listed above prepare the following: 

Standard liquid nutrient: 2 grams alfalfa hay, 3 grams wheat kernels, 
100 cc. glass-distilled water. Boil 5 minutes, pour off the liquid, filter it, 
restore to the original volume (100 cc.) by adding glass-distilled water, 
sterilize under 15 pounds' steam pressure for 10 minutes. 

Activating liquid, solution 1 : 5 cc. sterile standard liquid nutrient, 10 
cc. filtered sterile spring water, 1 loopful Achromobacter liquejaciens. 
Approximate pH value 6.2. 

Activating liquid, solution II: Dilute 2 parts of a freshly prepared 
activating liquid solution I to 50 parts by adding glass-distilled water; 
to 5 parts of a La Motte buffer mixture of known pH (best results ob- 
tained when buffers are in the range pH 6.2 to 6.8 inclusive) add 3 parts 
of the dilute activating liquid. Thus activating liquids may be pre- 
pared in the pH range of the buffers. 

Method: Prepare a cyst culture by obtaining 50 or more organisms 
in a Columbia culture dish ; then the glass cover for the dish should be 
sealed in place with petrolatum and the vessel set aside in a moist cham- 
ber until starvation and lack of oxygen induces encystment. Activate 
the cysts by removing the old culture fluid from the dish containing the 
cysts; wash cysts in three changes of distilled water and cover them by 
adding either solution I or solution II. At room temperatures of 20 
to 24 C, excystment begins within 30 to 55 minutes after activation. 
Conjugation begins approximately 14 hours after activation, reaches 
its maximum intensity at the end of 24 hours and begins to decline at 
the end of 36 hours. The duration of conjugation epidemics may be pro- 
longed for a variable period of 12 to 36 hours by removing all except a 
few drops of the liquid from the culture dish and adding fresh activating 
liquid; best results are obtained when this change is made at the time 
when the conjugation epidemic begins to subside. The method is 
invariably successful for Vorticella microstoma, V. convallaria, and V. 
nebulijera var. similis. The excystment technique is a modification 
of the one described by Barker and Taylor (1933). 

References 
For the culture of Vorticella see also pp. 60, 134, and 136. 

Bibliography 

Barker, H. A., and Taylor, C. V. 1933. Studies on the excystment of Colpoda 

cucullus. Physiol. Zool. 6:127. 
La Motte. 1933. The A. B. C. of pH control. Baltimore. 



!34 Phylum Protozoa 

Miscellaneous Classes and Microbiology 

A NOVEL METHOD OF OBTAINING PROTOZOA 

W. H. Davis, Massachusetts State College 

FOR years, when I taught zoology, I placed small, green grass culms 
(in April) in covered jars with wet cotton in the bottom. These 
remained in an upright position against the glass surface. When Amoeba, 
Arcella, Stentor, Paramecium, Vorticella, etc., were desired, I scraped 
dead plant tissue from the surface of the stems or mashed the rotten 
leaves and incubated 24 hours in a 1% aqueous solution of citric acid. 

For the region where this method was developed it did not fail for five 
consecutive years to produce the desired results. 

PERMANENT CULTURES* 

Very frequently instructors are required to keep protozoan cultures 
over long periods of time. The following method has been used with 
great success for such cultures as Paramecia, the smaller forms of 
Amoeba, and certain forms of flagellates. 

A large number of hay infusions are started in ordinary drinking- 
water tumblers, using pond water from different localities. They are 
then placed in various positions about the room and examined from time 
to time until the proper culture has been found. When a desired culture 
is found it should be fed five or six scrapings of dried whole wheat bread. 
These scrapings are made by taking a scalpel and scraping a crust of 
bread, care being taken to feed only what the culture will utilize. The 
glasses are then covered and the process repeated every two weeks or 
so. Whole wheat bread is far superior to ordinary wheat bread. 

Using the above method I have kept ordinary classroom cultures alive 
for a period of a year. It is also excellent for maintaining such cultures 
as rotifers and small crustaceans. 

FOOD ORGANISMS FOR MARINE AND HALOBIONT 

ANIMALS 

R. M. Bond, Santa Barbara School, Carpinteria, California 

Dunaliella salina is a large green, yellow, or orange flagellate of world- 
wide distribution in natural and artificial brines of various compositions. 
It is most easily obtained from salt-works recovering salt from seawater 
by solar evaporation. It may sometimes be raised from crude sea-salt, 
and I once recovered it from seawater from Monterey Bay. 

Pure cultures, free of all other organisms, may be grown in any of the 

* Reprinted from Science 64:362, 1926, by Frederick Bauer, Rhode Island State 
College. 



Miscellaneous Classes 135 

ordinary inorganic or organic culture media made up in seawater. Miquel 
seawater is very satisfactory. [See p. 33.] 

D. salina tends to die out in competition with other organisms, except 
in inorganic media containing 10-20% NaCl. It should, therefore, be 
kept in a stock culture of Miquel seawater plus 10% NaCl, or in some 
other equally concentrated medium, and subcultured (if necessary) in 
a more dilute medium before use. 

The cultures should be kept in strong light, though direct sunlight 
should be avoided in young cultures. The organism can grow through- 
out a wide temperature range, but 30 C. or just below seems to be 
optimal. 

Dunaliella viridis is always green, and is much smaller than D. salina. 
It seems to do best in a medium of 5-10% salinity. Otherwise, the 
statements made about D. salina hold equally true for this organism. 

Platymonas subcordaeformis is a small, green 4-flagellated alga. It 
is found in saline waters (up to 8-10% salinity) in warm-temperate re- 
gions, and is probably of wider range than has been reported. It is 
frequently found, often in very rich cultures, in spray-pools above tide 
line on rocks frequented by sea birds. It may sometimes be recovered 
from seawater. 

It grows rapidly and well in seawater (even considerably diluted sea- 
water) to which Miquel's solutions have been added, so that no concen- 
trated stock culture is required. 

Light and temperature requirements are as for Dunaliella salina. 

Reference 
For the culture of Ankistrodesmus see p. 227. 

WHEAT-GRAIN INFUSION 

John W. Nuttycombe, University 0} Georgia 

THE culture medium here described has been used constantly 
for nine years and has proven extremely satisfactory for culturing a 
wide variety of aquatic invertebrates. Its chief advantages lie in the 
ease of preparation, wide range of use and the relatively long period 
of time required for the culture to reach its maximum. 

In practice we add 200 or 300 grains of seed wheat to about 250 cc. 
of spring water in a flask. This is heated over a burner until the 
water comes to a sharp boil and is then allowed to cool. If it is desired 
that the cultures reach a maximum more quickly the boiling is con- 
tinued for several minutes so as to make the contents of the wheat grains 
more quickly available. 

We usually boil spring water to kill the free organisms in it, allow it to 



136 Phylum Protozoa 

cool to room temperature, distribute it in dishes* holding 200 cc. each, 
and place 3 or 4 grains of the prepared wheat in each dish of water. 
The dishes are now stacked and allowed to stand 2 or 3 days, during 
which time enough air is dissolved to make the medium ready for in- 
oculation. If it is desired to immediately inoculate the dishes the water 
may be artificially aerated. 

In general this first inoculation is made with material (bacterial glea, 
etc.) from successful cultures and after considerable growth has set up 
around the wheat grains (in 3 or 4 days) we subculture into the dish 
the particular organism desired from the best of the previous cultures. 

Initial cultures of an organism are made in the same way except that 
inoculations are made from the medium in which the organism was 
collected. 

We have made no attempt to control the pH of our cultures within 
any narrow limits but the range has been between 7.05 and 7.40. Our 
spring water generally has a pH value of about 7.05, a two-weeks cul- 
ture about 7.40 and a two-months culture about 7.12. There is some 
slight seasonal variation in the water which we use. 

We have cultured very successfully in this medium the following groups 
of fresh-water inverteb r ates: 

Protozoa — Several species of Amoeba, Actinophrys, various Difflugia, 
Chilomonas, Peranema and many other small flagellates, Paramecium, 
Vorticella, several hypotrichs, and many other Infusoria. 

Plathelminthes — Catenula, some 15 species of Stenostomum, Micros- 
tomum, and small triclads. 

Nemathelminthes — Several species of freshwater nematodes. 

Rotifera — Some 20 species. 

Annelida — Several species of oligochaetes. 

Bryozoa — Plumatella. 

Arthropoda — Copepoda, Cladocera, Ostracoda, Hydracarina, mos- 
quito and midge larvae. 

We especially recommend this method as a means of maintaining 
constant supplies of Amoeba for class use. We have for 5 years main- 
tained cultures in the original dishes by simply pouring out the water 
from each dish every two months and adding fresh water (boiled, cooled, 
aerated) and 3 or 4 grains of boiled wheat. A sufficient number of 
Amoebae stick to the bottom of the dish when the water is poured off 
to seed the culture. 

* The ice-box dishes (Hazel Atlas Glass Company) which we use for most of our 
general culture work may be purchased for 10 cents each at any io-cent store. They have 
a capacity of 400 cc; they may be stacked; and they may be obtained in either clear or 
green glass. These dishes are, for general purposes, quite as satisfactory as the much more 
expensive pyrex dishes sold for such purposes. 



Phylum II 

Porifera, Class Noncalcarea 



NOTES ON THE CULTIVATION AND GROWTH OF SPONGES 
FROM REDUCTION BODIES, DISSOCIATED CELLS, AND 

LARVAE 

H. V. Wilson, University of North Carolina 

PRODUCTION OF, AND GROWTH OF SPONGES FROM, REDUCTION BODIES 

SPONGES {Stylotella heliophila) are placed in outside aquaria, con- 
crete or wooden tubs, covered with glass and not in direct sunlight. 
The sponges should be clean, raised from the bottom on bricks; half a 
dozen to an aquarium 60 cm. in diameter and 30 cm. deep. The aqua- 
rium (tub) is emptied, filled, and flushed for some minutes three times 
in every twenty-four hours. Reduction begins in a day or two. In the 
course of two or three weeks gradual death of the tissues coupled with 
reduction leads to the formation of many small living masses of varying 
shape lodged on and through the skeletal network of the sponge. In the 
most striking cases these masses are numerous, more or less spheroidal 
and small, 1 to 1% mm. in diameter. Such a dead and macerated sponge 
body with its contained nodules of brightly colored living tissue suggests 
a Spongilla full of gemmules. The histological structure of nodules varies 
with their age, but is very simple, although many details in the process 
of reduction are unknown. 

The reduction bodies have regenerative power. If enclosed in bolting 
cloth bags and hung in a live box they quickly transform into sponges. 
Probably with a very excellent water supply the transformation could 
be induced in the laboratory. 

Similar bodies have been produced in the Calcarea (Otto Maas) and 
in freshwater sponges (K. Muller). (See Wilson, 1907a.) 

GROWTH OF SPONGES FROM DISSOCIATED CELLS 

A sponge {Microciona prolijera) is cut up into small bits about 3 
mm. in diameter, which are allowed to fall on a piece of fine bolting cloth 
supported on the edge of a stender dish and semi-immersed. The cloth 
is then folded like a bag around the bits of sponge, is partially immersed 

137 



138 Phylum Porijera 

in a small dish of filtered seawater, and while it is kept closed with the 
fingers of one hand is repeatedly squeezed between the arms of a small 
forceps. The pressure and the elastic recoil of the skeleton break up 
the tissue into its constituent cells and these pass out through the pores 
of the cloth into the surrounding water. The cells fall to the bottom 
and may be sown with a pipette on any desirable substratum (slide, 
cover glass, or oyster shell) immersed in a culture dish of seawater. Or 
the cells, as pressed out, may be allowed to fall at once on the definitive 
substratum. The cells attach in the course of an hour or so and the 
slide or other body may be removed to a fresh dish of water or to a 
running water aquarium, where it should be raised well above the bottom 
and protected from the force of the current. Attachment will usually 
take place by means of coarse reticula which remain permanently at- 
tached. Reticular pieces if partially freed from the slide will curl up and 
form balls, the size of which is under control. Such balls may be trans- 
ferred to slides in other dishes or in running water aquaria. A convenient 
vessel to use is a porcelain-lined bucket, in which the slides rest on in- 
verted bottles and are so brought near the surface of the water, the 
current entering at the bottom. By changing the water two or three times 
a day, such an arrangement serves in place of a running water aquarium. 
The balls attach and sponges of desired size may be obtained. The at- 
tached reticula or balls metamorphose and in the course of a few days 
will have transformed into incrusting sponges with functional canal 
systems. Such cultures are easily kept for long periods of time in small 
wire gauze cages hung in live boxes. Lobular outgrowths and even 
embryos have developed in sponges treated in this way. 

Modifications in this method of growing sponges have been introduced 
by J. S. Huxley, K. Miiller, P. Galtsoff, M. E. Faure-Fremiet, M. E. de 
Laubenfels, J. T. Penney, P. Brien, and others. (See Wilson, 1907 and 
1911.) 

GROWTH OF SPONGES FROM CILIATED LARVAE 

Mycale (Esperella) fibrexilis has embryos during July-August at 
Woods Hole, Mass. Larvae are liberated, sometimes at once, on placing 
the sponge in an ordinary 2 gallon glass aquarium jar. Larvae are 
picked out with a pipette and transferred to culture dishes where they 
may be kept by changing the water several times a day. They attach 
in a day or two. They may be made to attach to cover glasses or if they 
are to be used as section material it is convenient to coat the dish with a 
thin layer of paraffin and let them attach to this. Little pieces of paraffin 
with the attached and metamorphosing larvae may then be cut out, fixed 
(paraffin and sponge), and hardened, the sponge often detaching itself 
from the paraffin. 



Noncalcarea 139 

Other halichondrine sponges (Lissodendoryx, Microciona) breed dur- 
ing the summer (July- August) at Beaufort, N. C; Stylotella during 
October in the same locality. Larvae may be reared in the same way 
or may be placed in wire gauze cages after attachment and hung in a 
live box. 

In sponges in general, fertilization is internal and the egg develops in 
the body of the parent to the stage of the ciliated larva. Sponge eggs 
and embryos are commonly abundant in a breeding sponge and may be 
seen scattered through the interior with the naked eye. In some marine 
sponges asexual masses analogous to spongillid gemmules develop likewise 
in the body of the parent into ciliated larvae. In order to obtain larvae 
all that is necessary is to place a breeding sponge in an aquarium jar. 
(See Wilson, 1894.) 

GROWTH OF SPONGES FROM FUSION LARVAE 

The ciliated larvae of Lissodendoryx carolinensis may easily be made 
to fuse with one another after they have begun to creep over the bottom 
of the culture dish and are thus approaching the phase in which they 
attach. It is only necessary to bring them in contact, coaxing them 
together with needle and pipette in a deep, round watch glass. The 
compound larva so produced has a feeble locomotory power. Using 
pairs that are nearly motionless, fusion masses of desired shapes may be 
produced on cover glasses. Or small excavations may be made in 
paraffin-coated dishes, and the larvae driven into such holes in large 
numbers. In this way, cake-like masses may be produced measuring 3-4 
mm. in diameter. The smaller compound masses metamorphose without 
difficulty. The larger in the actual experiments died, sometimes after a 
partial metamorphosis. (See Wilson, 1907.) 

Bibliography 

Wilson, H. V. 1894. Observations on the egg and gemmule development of marine 

sponges. /. Morph. 9:277. 
1907. On some phenomena of coalescence and regeneration in sponges. 

J. Exper. Zool. 5:245. 
1907a. A new method by which sponges may be artificially reared. Science 



25:912. 

191 1. Development of sponges from dissociated tissue cells. Bull. U. S. Bur, 

Fish. 30:1. 



Phylum III 

Coelenterata, Class Hydrozoa 



HYDRAS 

Libbie H. Hyman, New York City 

Collection. Hydras most commonly occur attached to the submerged 
vegetation, fallen leaves, or other objects in pools, ponds, lakes, and 
the slow portions of rivers. They are collected from such habitats by 
gathering a quantity of vegetation and placing it in jars with a relatively 
small amount of water. As the vegetation begins to decay, the hydras 
usually come to the top of the jar or the surface film and may be picked 
out and transferred to suitable containers. One species, Hydra littoralis, 
occurs in enormous numbers on the under surface of stones in streams, 
spillways, and along shores subject to wave action. When the stones 
are turned over, this species appears like an orange jelly on the stone. 
They may be obtained in large numbers by squirting the animals from 
the stone into a pan by means of a squirter made from an atomizer bulb 
and a glass tube. Unfortunately this species is not very suitable for 
laboratory cultivation, but is useful when large numbers are needed 
for a short time, or when large quantities of hydras are wanted for pres- 
ervation. Hydras are sometimes found in large numbers attached to 
the surface film and in such situations are easily gathered. They do not 
in general live in stagnant water but require rather clean water with 
adequate oxygen supply. 

Laboratory cultivation. The most suitable hydra for laboratory cul- 
tivation is the brown hydra, Pelmato hydra oligactis, but any of the 
species which live in standing water may be grown successfully in the 
laboratory. The species which inhabits moving water, Hydra littoralis, 
mentioned above, may be grown in the laboratory if the water con- 
tains a supply of algae or other vegetation to keep up a high oxygen 
content. The author has not tried bubbling air through a culture of 
this species but such a procedure would probably be successful. The 
green hydra, Chlorohydra viridissima, is one of the most hardy hydras, 
very common everywhere, and easy to maintain in the laboratory except 
for one difficulty. It is difficult to find a food crustacean which is small 

140 



Hydrozoa 141 

enough to be ingested by the green hydra. Cultures of the green hydra 
should be exposed to the light, but for other species a moderate light 
is best. 

The first prerequisite of laboratory cultivation of hydras is suitable 
water. Hydras should never be put into freshly drawn tap water and 
in general only natural pond or river water should be used for their 
culture. If tap water must be used, it must stand for several weeks 
with growing algae or other aquatic plants in it so that it may become 
conditioned. The suitability of water for hydras should be tested by 
placing a few specimens in it. If these expand fully, with tentacles 
extended to their maximum extent, the water is suitable. In unsuitable 
water, the column remains contracted and the tentacles fail to expand. 
The presence of plants is not necessary in a hydra culture except in the 
case of Hydra Uttoralis, as stated above. 

When a suitable water has been found, the hydras are placed in it, and 
fed daily. At intervals, the accumulation of bottom debris should be 
removed and small amounts of suitable water may be added from time 
to time. In general it is desirable that the cultures be covered. 

In general hydra cultures succeed better if the temperatures are not 
too high; 20 C. is a very suitable temperature. The brown hydra, 
P. oligactis, is more susceptible to rise of temperature than any other 
species and usually dies at 25 C. 

The great difficulty in the continuous culture of hydra is the occur- 
rence of the phenomenon of depression. In spite of every care, hydra 
cultures will pass into this state at intervals and, unless prompt measures 
are taken, will die out. In depression, column and tentacles fail to 
expand, the animal ceases to feed, shortens to a stumpy appearance, and 
finally gradually disintegrates from the tips of the tentacles aborally. 
Depression is caused by over-feeding, fouling of the water, too high 
temperatures, and general aging of the culture with accumulation of 
waste products. The most successful method of reviving the animals 
from the depressed state is to transfer them to a fresh jar of suitable 
water. Lowering the temperature is also of assistance. 

Sex organs when wanted for class display may be induced in most 
species of hydra, notably in the brown hydra, by placing the culture in a 
refrigerator for two or three weeks at a temperature between 10 and 15 C. 
Such cultures should be fed regularly. In nature most species are found 
with sex organs in the late fall but the green hydra is said to be sexual 
in the summer. 

Food. Naturally for the continued maintenance of hydra in the 
laboratory it is necessary to have a food source that is easily cultured. 
The most suitable animal for this purpose is Daphnia because of its 
slow movements, weak resistance, and habit of moving about continu- 



142 Phylum Coelenterata 

ously.* Some other cladocerans such as Simocephalus are easily cul- 
tivated also but are too strong and powerful for the hydras or else tend 
to stay on the bottom out of reach. Hydras will not eat ostracods. Oli- 
gochaete worms are eagerly accepted but their habits are such that 
the hydras would seldom have a chance to catch them. The very tiny 
newly hatched Daphnia in a Daphnia culture may be used as food for 
Chlorohydra viridissima; some small forms such as Ceriodaphnia or the 
bosminids are suitable and may be grown like Daphnia. [See pp. 207- 
220.] In case of necessity hydras may be fed on oligochaetes such as 
naids, tubificids, or enchytraeids, and often these may be purchased from 
pet shops. It is necessary to cut these up into pieces and to place the 
pieces in contact with the tentacles with a dropper or forceps ; otherwise 
they fall to the bottom where they are out of reach of the hydras. This 
method of feeding is very time-consuming, but may be used when it is 
desired to save valuable experimental material. Experimental material 
which does not feed well of its own initiative, may often be fed success- 
fully by placing a crushed daphnid, Cyclops, or bit of oligochaete in con- 
tact with the tentacles. 

Tubificids are easily maintained in the laboratory for food by placing 
them in containers having two or three inches of pond mud on the 
bottom. For food, almost any kind of organic material such as boiled 
lettuce leaves, boiled wheat grains, pieces of bread, or bits of animal 
flesh, may be added from time to time. 

THE CULTURE OF SOME MISCELLANEOUS SMALL 

INVERTEBRATES 

Paul Brandwein, Washington Square College, New York University 
Hydra. These animals are maintained in great numbers in balanced 
aquaria when they are fed constantly with any of the Entomostraca men- 
tioned below. 

Plathelminthes. Stenostomum may be cultured using the method used 
for Paramecium. [See p. 63.] Planaria are usually cultured in enameled 
pans containing clear pond or spring water; they are fed with boiled 
egg yolk or fresh liver, care being taken to remove the excess food at 
the end of a few hours before putrefaction occurs. The Planaria are cut 
transversely when they reach the size of about 8-12 mm. with a 00 
cover glass; regeneration occurred rapidly.** 

* Editor's Note: William LeRay and Norma Ford, of the University of Toronto, call 
attention to the fact that in contrast to the grey hydra (H. vulgaris americana) the brown 
species (Hydra i = Pelmatohydra] oligactis) stings its prey only when it needs it for food. 
Thus in a culture of the brown form the uneaten Daphnia continue to live. The grey 
hydra on the other hand stings to death any Daphnia which it happens to touch and on the 
bottom of its culture bowl there will be found a ring of the dead crustaceans. M.E.D. 

** This method for Planaria does not differ significantly from the procedure recommended 
by the commercial houses which supply this organism. 



Scyphozoa 143 

Rotijera. Philodina is easily cultured by either the method of 
Stylonychia [See p. 64.] or of Paramecium. [See p. 63.] 

Entomostraca. Cyclops, Canthocamptus, Diaptomus, Cypris, and 
Daphnia have been cultured with moderate success by the following 
method. Two grams of egg yolk, ground into a paste, are added to a 
gallon jar filled with green pond or aquarium water. This is allowed 
to stand for about three days and then inoculated with small Protozoa 
(any species not larger than Colpoda). Next the organism to be cul- 
tured is introduced — for Cyclops, Canthocamptus, and Diaptomus a few 
males and egg-bearing females will suffice, but for Daphnia and Cypris, 
as many individuals as possible are added. Within a month successful 
cultures will show organisms in abundance, a condition which will last 
some 6 weeks.* 

Annelida. Microdrilli, such as Nais, Aelosoma, and Dero have re- 
sponded splendidly to culture in 30 cc. of Solution A (see footnote on 
p. 73) added to rice-agar as in the case of Amoeba [see p. 72]. The 
medium in this case, however, should stand for three days before in- 
oculating with the annelids (about 5 will suffice). The number of these 
organisms can be increased by using larger vessels, i.e., allowing for more 
fluid and increasing the surface area of the agar. 



Class Scyphozoa 



REARING THE SCYPHISTOMA OF AURELIA IN THE 

LABORATORY** 

F. G. Gilchrist, Pomona College 

THE scyphistoma of Aurelia proves to be a very hardy marine form; 
it may be maintained alive and in fairly active state of budding by 
keeping it in shallow dishes of sea water (it is well to have the sea- 
water slightly hypotonic) and feeding with ground shrimp or particles 
of meat. Of course the water should be changed after feeding. The 
scyphistoma does best with a mixed diet. Scyphistomas have been 
reared at marine laboratories, using plankton tow (Delap, 1905, 1907) 
or sea urchin ovaries (Herouard, 1909). 

Bibliography 

Delap. 1905. Rep. of Sea and Inland Fisheries of Ireland for 1902 and 1903. 

■ 1907. Ibid, for 1905. 

Herouard. 1909. C. R. Acad. Sci. Paris, p. 148. 

♦For Daphnia, the cultures require a temperature of 17-19 C. This was obtained by 
circulating cold water through a coil of glass tubing (6-8 mm.), set within the gallon jar. 

** Editor's Note: For a more complete description of the methods of rearing Aurelia 
aurita and other medusae see Hagmeier, A.: Die Zuchtung verschiedener wirbelloser Mee- 
restiere. In Abderhalden, 192 7-1933: Handbuch der Biologischen Arbeitsmethoden Abt. 
9, Teil 5:553-562. P. S. G. 



144 Phylum Coelenterata 

Class Anthozoa 

SAGARTIA LUCIAE 

Donald W. Davis, The College of William and Mary 

Sagartia luciae may be collected at any season from tide pools, piles, 
rocks, and seaweed within its range — Atlantic coast, Massachusetts to 
Virginia; also Oakland Harbor, San Francisco Bay. It lives indefinitely 
in the laboratory with slight care and reproduces freely asexually, but has 
not been known to reproduce sexually under laboratory conditions. 
Specimens should be placed in seawater of a depth of one or two inches 
and exposed to diffuse sunlight. The glass container may well be cov- 
ered lightly to reduce evaporation and to exclude dust. For the first few 
days after bringing specimens into the laboratory care should be exer- 
cised that the water does not become foul through decomposition of frag- 
ments of the specimens, of undigested food that they may eject, or of 
other organisms that do not survive the change. It is, therefore, ad- 
visable to change the water occasionally during the first few days. If 
economy of the water supply is required, it should be filtered and may 
then be used over and over. 

These anemones thrive in tide pools of a rocky coast and gentleness 
is not essential. Before long a growth of algae appears on the dish and 
takes care of the oxygen supply. Probably it provides directly or in- 
directly for the food requirements of the anemones as well, for specimens 
thrive without special provision for feeding. They will take minute 
fragments of fish, crab, or beef. If so fed, only firm fragments, not 
juicy materials, should be used and the greatest care must be taken that 
fragments, unin jested or voided after a few minutes or hours, be not left 
to foul the water. 

One other precaution is of much importance. If specimens are left 
undisturbed, a zoogloea-like coating, probably consisting of slime with 
imbedded organisms, covers the column and eventually the whole con- 
tracted specimen. At intervals of two or three days each such coat 
should be removed from the dish after being separated from the anem- 
one by means of a gentle stream of water directed at its attachment to 
the glass. Occasionally rain water should be added to compensate ap- 
proximately for evaporation. 

Bibliography 

Davis, Donald W. 1919. Asexual multiplication and regeneration in Sagartia luciae 
Verrill. J. Exper. Zool. 28:161. 



Anthozoa 145 

REARING CORAL COLONIES FROM CORAL PLANULAE 

Thomas Wayland Vaughan, Scripps Institution of Oceanography 

DURING my field work on the corals of Florida and the Bahamas 
from 1908 to 19 1 5, an endeavor was made to rear coral colonies 
from planulae in order to ascertain the growth rates of young colonies 
of the different species. Although the technique used in collecting and 
rearing planulae and young colonies has been described in a number of 
publications, all that is essential is contained in those at the end of this 
note. 

The colonies from which it was hoped to obtain planulae were brought 
from the sea into the laboratory and placed in glass vessels which were 
deep enough for the specimens to be covered with seawater, but not so 
deep as to interfere with noticing any planulae that were extruded. Since 
the water on the parent colonies kept in the laboratory had to be pure, 
it was necessary to change the water on specimens kept for several days. 
This was easily done by siphoning the stale water from around the speci- 
mens and pouring fresh water into the vessels. 

The planulae which were extruded were removed by pipettes and 
transferred to vessels that contained objects on which it was expected 
that they would settle. 

In my work on the corals of Tortugas, planulae were obtained from 
five species, as follows: Astrangia solitaria, Favia fragum, Agaricia pur- 
purea, Pontes clavaria, Pontes astreoides. The species with which 
most success was obtained were Favia fragum and Pontes astreoides. 
The duration of the free-swimming larval stage is variable both for the 
same species and for different species. The duration for Favia fragum 
was 6 to 23 days; Agaricia purpucea, 11 to 17 days; Pontes clavaria, 
12 to 20 days; P. astreoides, 7 or 8 to 22 days. 

An effort was made to have the planulae settle on tiles (terra-cotta 
discs) having a central perforation by which they might be fitted over 
the heads of iron stakes. The tiles had a diameter of 8 inches and were 
placed in jars, the inside diameter of which was about 8.25 inches and 
the depth about 8.5 inches. After the bottom of a jar had been covered 
with clean sand, a tile was placed in it and the central perforation and 
the space between the periphery of the tile and the sides of the jar were 
filled with sand to the level of the upper surface of the tile. As the 
planulae tend to settle in depressions, it was necessary to fill these 
spaces. After this preparation, fresh seawater was gently poured in 
through a funnel until the jar was nearly full. The extruded planulae 
were pipetted from the vessels containing the parent colonies and placed 
in the culture jars. 

The water in the culture jars must be fresh and pure. It may be 



146 Phylum Coelenterata 

changed by one of several devices. In order not to draw off the planulae, 
which are very small, a bag of fine-mesh bolting cloth must be affixed 
to any tube used in withdrawing the stale water. One method was to 
siphon off the stale water with a rubber tube, the end of the tube inserted 
into the culture jar having been drawn over one end of a glass tube, the 
other end of which was enveloped in a bolting cloth bag. The table on 
which the culture jars stood was provided with a gutter into which the 
water drawn off was discharged, ultimately flowing outside the building 
through a pipe through the floor. After a jar had been emptied to within 
an inch of the tile, it was refilled with fresh seawater. This method 
caused a change in the level of the water, and by the pouring stirred up 
the unattached planulae. 

A second method, which was the one usually employed, was to with- 
draw the old water by a glass siphon resting on the upper edge of the jar, 
the siphon having been rendered non-emptying by having its outer end 
bent upward. The inner end of the siphon was enclosed by a bolting 
cloth bag. Fresh seawater was added by a siphon extending to the 
bottom of the culture jar from a supply jar placed at a higher level. 
By this method a constant level was maintained in the culture jars ; the 
old water was drawn off from the top while the new water was added 
at the bottom. A third method was to have inside the culture jar a 
tantalus siphon emptying through the side of the jar near its bottom. 
Fresh water was siphoned into the culture jar from supply jars placed at 
a higher level. When the water in a culture jar had reached the level 
of the upper curvature of the siphon, it began to run out and continued 
to flow until the level of the open end of the siphon in the jar was 
reached. The jar was then refilled by the afferent siphon until the level of 
the upper curvature of the tantalus siphon was again reached, when the 
water again began to flow out. This method caused a rise and fall 
in the level of the water. A fourth method was to cut the bottom out 
of a culture jar and to place the glass collar thus produced over a tile 
in a jar of larger diameter, the bottom of which had previously been 
covered with sand to a depth of an inch or slightly more. The tile 
and its surrounding collar were sunk into the sand until the upper sur- 
face of the tile and the upper surface of the sand were level with each 
other, while the level of the upper edge of the collar remained slightly 
higher than that of the enclosing jar. Water was siphoned into the 
collar from supply jars, and filtered through the sand filling the space 
between the collar and the side of the inclosing jar. When the level 
of the upper edge of the jar was reached, the water overflowed. This 
method maintained a constant level of water, drew off old water at the 
bottom, and added new water at the top. 

All four methods were successful, but as the second was somewhat 



Anthozoa 147 

the more convenient it was, as stated above, the one used in most of the 
experiments. 

After the planulae had attached themselves to the terra-cotta discs 
and had begun to form small colonies, the discs were affixed to the heads 
of iron stakes driven into the sea bottom at convenient places where the 
discs would always be submerged at the lowest tides. Through the end 
of the stake, which extended through the central perforation of the 
disc, there was a hole in which an iron pin was placed. This pin held 
the disc firmly on the head of the stake. By this means colonies of 
both Favia fragum and Porites astrcoides were reared to an age of five 
years. The diameter of the colonies of Favia fragum for colonies five 
years old ranged from 28.5 to 38 mm., that of the colonies of Porites 
astrcoides ranged from 41 to 99.75 mm. The height of the colonies of 
Favia fragum at five years of age ranged from 13.5 to 22 mm., that of 
colonies of Porites astrcoides ranged from 18 to 54.4 mm. 

Bibliography 

Vaughan, T. W. 1910. The recent Madreporaria of southern Florida. Carnegie 

Inst, of Washington, Year Book 9:135. 
1 91 1. The Madreporaria and marine bottom deposits of southern Florida. 

Ibid. 10:147. 

1919. Corals and the formation of coral reefs. Smithsonian Inst. Publ. 



2506, Report of 1917:189. 



Phylum V 

Plathelminthes, Class Turbellaria 



Order rhabdocoelida, Family catenulidae 

CULTURE OF STENOSTOMUM OESOPHAGIUM 

Margaret Hess, Judson College, Marion, Alabama 

CULTURE medium for Stenostomum oesophagium is prepared in the 
following manner: 

Boil 250 cc. of water with 8 to 10 grains of wheat for one minute; 
allow to stand exposed to the air for 24 hours; remove about two-thirds 
of the wheat grains and inoculate with a mixed laboratory culture of 
Protozoa. 

Introduce Stenostomum oesophagium into this culture 24 hours or 
more after the addition of the Protozoa. The presence of other Turbel- 
laria, oligochaetes, and small Crustacea has no harmful effect on Sten- 
ostomum oesophagium. 

Varying temperatures have little effect on Stenostomum oesophagium, 
although room temperature has been found the most satisfactory for 
rapid growth and multiplication. Likewise they are able to withstand 
variations in hydrogen-ion concentration. The best cultures show a pH 
range of 5.8 to 7.6.* 

CULTIVATION OF STENOSTOMUM INCAUDATUM** 

T. M. Sonneborn, Johns Hopkins University 

THIS turbellarian may readily be cultivated in mass or in isolation 
pedigree cultures if fed copious supplies of the ciliate Protozoan, Col- 
pidium campylum. The basic medium is prepared by boiling for 10 
minutes 15 grams of whole rye grains in one liter of spring water. This 
infusion is filtered while hot, cooled, inoculated with 1 cc. of a similar 

♦Editor's Note: Concerning two other species of Stenostomum, Jeanette Seeds Carter 
states (/. Exper. Zool. 65:159, 1933) that S. grandc is naturally cannibalistic and that in 
5. tenuicauda cannibalism is not a normal phenomenon. J. G. N. 

** Condensed by the author from: Genetic studies on Stenostomum incaudatum {nov. 
spec). I. The nature and origin of differences among individuals formed during vegetative 
reproduction. J. Exper. Zool. 57:57, 1930. (See pp. 62 and 63.) 

148 



Micro st omidae 149 

i-day old infusion, and allowed to stand at io° to 14° C. for 1 day. Then 
the ripened infusion is inoculated with Colpidia, placed in a 9-inch petri 
dish, and allowed to stand for 4 days. After this time the culture is 
centrifuged for 30 seconds at 1800 revolutions per minute. The Col- 
pidia form a dense, almost solid mass at the bottom, where they may 
be separated from the supernatant fluid and the middle layer of debris. 
The concentrated Colpidia should then be diluted to about 15 cc. with 
1 -day old rye infusion. 

Stenostomum incaudatum may be cultivated in isolation pedigree lines 
by placing one Stenostomum in a single drop of this fluid on a depres- 
sion slide. Temperatures of 20 to 2 6° C. are favorable. The culture 
fluid should be renewed daily, by transferring one Stenostomum to a fresh 
slide with fresh, concentrated Colpidium fluid. Mass cultures may be 
reared in similar rye infusion to which heavy growths of Colpidium have 
been added. These cultures must be renewed before the supply of 
Colpidia gets low. 

References 

For the culture of Stenostomum see also pp. 136 and 142. 
For the culture of Catenula see p. 136. 

Family Microstomidae 

THE CULTURE OF MICROSTOMUM 

M. Amelia Stirewalt, University of Virginia 

MICROSTOMUM is particularly sensitive to very small traces 
of such poisons as are used in chemical reagents. In the selec- 
tion of glassware to be used in the culture of these animals, therefore, 
care must be taken that dishes, pipettes, etc., have had no contact with 
fixing and staining reagents. Petri dishes have proven most successful 
as aquaria because the large, flat, bottom surface presents ample space 
for the benthal habits of Microstomum. In these dishes the animals may 
easily be seen with the naked eye, especially if the culture is placed over 
a dark background. Both stender dishes and larger culture dishes, 
however, will serve as aquaria, though the small capacity of the former 
necessitates frequent change of the culture medium, and the large size of 
the latter makes close observation of particular animals impossible. 

Into the aquarium selected, in the approximate proportion of 9-1, place 
spring water (from a non-limestone district) and water containing small 
detritus from the bottom of the stream or pond in which the Microstoma 
were collected. It is best that no animals be present which can be seen 
with a magnification of 20. To this medium may then be added such 
Cladocera as Cypris, Daphnia, and their relatives [see pp. 207-220], and 



150 Phylum Plathelminthes 

such Copepoda as Cyclops [see p. 227]. Of aerating value are a few 
branches of Elodea or a small mass of Spirogyra or both. In this culture 
the Microstoma may then be placed. Microstoma living in the presence 
of water plants are more active and of larger size than those living in 
control cultures without these plants. In such a culture the animals may 
be expected to thrive indefinitely, if at intervals some of the detritus be 
drawn off with a clean pipette and replaced with fresh water, and if 
food be supplied regularly. 

The food which serves best, in my experience, is the annelid Dero 
[see p. 143]. Every two or three days the Microstoma should be fed 
small, freshly cut sections. Enough should be placed in the culture to 
supply each Microstomum with three or four pieces, for the animals 
eat voraciously when in a healthy condition. The Dero may be cut 
easily by means of two small needles used in criss-cross fashion. The 
pieces should not exceed the size of the Microstoma and must be freshly 
cut. If the food has been prepared for an hour or more before feeding, 
the wounded surfaces of the annelid heal, thus cutting off the flow of 
the fluid by which the Microstoma sense the presence of the food most 
readily. In such case, or whenever the Microstoma seem insensible to 
the presence of food, they may find and eat it if several pieces are freshly 
cut in the culture. 

Other conditions being favorable to growth, the size of the Microstoma 
is directly related to the amount of food consumed. Animals with six 
zooids often occur in vigorous cultures. On the other hand I have had 
Microstoma live in favorable cultures for two weeks without feeding. 
Under such conditions they become progressively smaller until they die 
from starvation. 

The foods eaten by Microstomum under my observation, listed in 
order of preference, are: Dero, Hydra, liver (tadpole and mammalian), 
Cypris, Daphnia, Cyclops, Difflugia, Pristina, egg yolk, Stentor, desmids, 
and Nematodes. 

Hydra, as food for Microstomum, deserves special mention. It seems 
to act as a tonic for animals which are not in good condition as evi- 
denced by their lack of response to food. If, as sometimes occurs, the 
Microstoma cannot sense food, or refuse it, fragments of hydra, fed in 
the same way as the annelids, will rejuvenate them. 

Reference 
For the culture of Microstomum see also p. 136. 



Planar iidae 15 1 

GEOCENTROPHORA APPLANATUS 

William A. Kepner, University of Virginia 

THIS member of the group Alloeocoela may be cultured in spring 
water to every 200 cc. of which 5 cc. of wheat infusion has been 
added. (Wheat infusion: 10 seeds wheat in 250 cc. spring water. Boiled 
one minute. Set aside for one wee*. This infusion may be used there- 
after for two months.) 

The specimens have been fed with food described by Margaret Sans- 
low in the Bull, of Averett College (Danville, Va.), Vol. 1, No. 4, IQ35-* 
To a small dish containing 205 cc. of water there has been added as 
much food as will cling to the moist tip of a very small scalpel. This 
food is cut into very short lengths and then ground in a depression slide 
with the rounded end of a small glass rod, after which it is placed in the 
culture dish. The specimens will find this food within fifteen minutes 
if it is not widely distributed. 



Order tricladida, Family planariidae 

CULTURE OF PLANARIA [=EUPLANARIA] AGILIS 

Rosalind Wulzen, Oregon State College and 
Alice M. Bahrs, St. Helen's Hall Junior College 

PLANARIA for use in nutrition experiments are collected in the 
field by placing small lumps of fresh liver in shallow water at the 
edges of ponds or streams where they are known to be present. They 
gather on the liver in a short time and may be rinsed off into collecting 
jars. They should not be crowded in the jars or they will be dead before 
the laboratory is reached. Likewise, they should not be crowded in the 
laboratory containers. For stock containers we use white enameled 
milk pans, because in these the worms may easily be inspected to deter- 
mine their condition. A city water supply containing chlorine is not 
to be trusted. For some time it may appear to be harmless but when 
one observes that the worms are more restless than usual, that is, all the 
worms in a container are in motion for an extended period of time, 
one should suspect that the tap water contains too much chlorine. The 
restless stage is followed by one in which the worms secrete large quan- 
tities of mucus and roll away from contact with the container. They 
gather in writhing masses and will disintegrate if they are not put into 

♦Editor's Note: This food consists of shrimp, corn flakes, shredded wheat, lettuce, 
spinach, and sea lettuce. These last two ingredients are dried quickly in a flower press. 
All the materials are separately powdered to medium grains with mortar and pestle and 
mixed together in amounts such as to provide 50% protein, 31% carbohydrates, 2% 
fats, 12% minerals, and 5% bulk by volume. M. E. D. 



!^2 Phylum Plathelminthes 

chlorine-free water. We use river or well water collected directly from 

the source. 

After the worms have been distributed in the stock pans they must 
be treated as though in quarantine for about a month. Every day they 
must be inspected carefully and any worm showing the slightest irre- 
gularity in outline or surface texture must be removed. We have found 
that the worms come to the laboratory infected with parasitic diseases 
which develop quickly under the abnormal conditions of a laboratory 
environment. These diseases are capable of spreading and of annihilat- 
ing a large part of the stock. If one is to have stock reliable enough for 
experimentation, all disease must be eliminated, and with care this 
is easily accomplished. We always boil all water to be used on the 
worms in order to avoid the introduction of any disease-producing para- 
site. If at all possible, the water used should be perfectly clear because 
we have found that even slightly muddy water reacts unfavorably on the 
worms. 

The laboratory routine in the care of the stock is as follows. The 
worms are washed every other day. This is done by plunging the 
hand into a lysol solution and then rinsing until no odor of lysol re- 
mains, for very slight amounts of lysol are highly poisonous for the 
worms. Then with the fingers the pan is wiped over its whole surface 
to loosen the dirty slime which always gathers. The worms settle at once 
to the bottom and all the water is poured away and replaced by fresh 
water. If the pan does not appear clean, this is repeated. Once a week 
the worms are washed into freshly sterilized pans and the dirty pans are 
thoroughly cleaned and sterilized. 

Our stock worms are fed exclusively on raw liver and they continue 
in vigor and health for an indefinite period of time. The source of the 
liver must be considered because not all liver has correct nutritional 
value for the worms. For example, rat liver is poor food while beef 
liver is almost always excellent. We use liver taken from freshly-killed 
guinea pigs which are in prime condition. This has been found the best 
stock feed we have tried because we can control the diet of the animals 
furnishing the liver, and this is the determining factor in the production 
of nutritionally correct liver. If the stock is merely being maintained 
it is sufficient to feed once a week. The feeding is done by placing 
a small piece of liver in each stock pan. The worms feed readily and 
the liver is left with them for 3 or 4 hours. It is then removed and the 
stock pans are thoroughly washed. If one wishes to develop the stock 
rapidly, the worms should be fed twice a week. 

To rear new worms for experimental purposes it is of course only 
necessary to cut the stock worms into pieces of suitable size and to allow 
time for regeneration. We always cut off the posterior extremities of 



Planariidae 153 

the worms, the length of the piece cut off depending upon the size of the 
worm. We separate these tail pieces into pans by themselves and allow 
them to stand without feeding for a period of 4 weeks, at which time 
they have attained the adult shape and are ready for nutritional ex- 
periments. 

We keep our experimental worms in an incubator with a temperature 
of 24 C, but the stock and regenerating worms are kept in the labora- 
tory with the heat turned on at night during the cold season. If the 
worms become thoroughly chilled many or all of them will die. This 
happens above the freezing point and makes it wisest to keep the stock 
pans away from cold windows. 

PLANARIA 

William LeRay and Norma Ford, University of Toronto 

FOR eight years a culture of Planaria [=Euplanaria] maculata has 
been kept under observation and fed very successfully with enchy- 
traeid worms. The animals were collected in a large pond and a selec- 
tion was made of the individuals which would accept the enchytraeid 
worms as food. 

The planaria are kept in a wooden tub (24 inches in diameter and 
n inches deep), charred on the inside, and are fed about once a week. 
Approximately 2000 individuals are maintained in this space. When 
fed, a level teaspoonful of worms is dropped over the bottom of the 
tub. Several planaria will cluster over each worm and so share the food. 
The amount of food given is regulated by the growth of the planaria: 
if they are getting smaller, more enchytraeid worms are offered; if more 
individuals are needed, additional food will speed up their growth and 
reproduction. 

When large numbers of planaria are needed for class material, we 
are careful not to disturb the tub for two or three days. A film then forms 
over the surface of the water. To bring the planaria to the surface, the 
sides of the tub are tapped with a hammer. Each animal is then picked 
out with a fine glass rod which is slipped under its dorsal side as it floats 
ventral side up. The planaria folds its dorsal surface around the rod 
and it is then dropped quickly into a dish for study. Without the film 
on the surface of the water the planaria will not stick to the rod. 

COLLECTION AND CULTURE OF PLANARIA 

George R. La Rue, University of Michigan 

BAITING for planaria by the method described by Hyman (Trans. 
Amer. Micr. Soc. 44:79, 1925) may not always be practicable. 
Planaria often occur in abundance, sometimes by hundreds, on the lower 
surface of stones or submerged boards in swiftly flowing water below 



1 54 Phylum Plathelminthes 

dams, in the riffles of streams, or on wave-washed shores of lakes. If 
such waters are not frozen over, collections may often be made in mid- 
winter. Stones or boards should be removed from the water and the 
lower surfaces examined for planaria. When found in suitable numbers 
scrape the worms off into dishes or pails of water, or bring in the small 
stones with worms adhering. 

Planaria may usually be secured in considerable numbers by bringing 
in masses of submerged vegetation. Cover this material with water, 
preferably pond water or untreated tap water in large glass jars or 
aquaria, and allow decay to start. The worms will collect on the surface 
of the water and on the sides of the vessel. Transfer them to finger 
bowls or larger vessels of clean water, and keep the dishes in a darkened 
place. 

Feed planaria on tubificid worms, giving only as many tubificids as 
will be eaten in 3 or 4 hours. Since these worms are completely ingested 
and live until eaten, the dishes need not be cleaned as frequently as when 
liver is fed. Dishes should be washed once or twice a week, but soap and 
other chemicals must be avoided. 

References 

For the culture of planaria see also p. 142. 
For the culture of triclads see also p. 136. 

PLANARIANS 

Libbie H. Hyman, New York City 

Collection. Different species of planarians live in different sorts of 
habitats. Some, notably our most common species, Euplanaria macu- 
lata, live in ponds, lakes, and the slow parts of rivers on the vegetation 
and on the under surface of stones, leaves, or other objects. They may 
be obtained by turning over stones and fallen leaves and washing the 
animals into a pan by means of a strong squirter made of an atomizer bulb 
and glass tube. Their presence on vegetation may be ascertained by 
shaking small samples of the vegetation in a vessel of water. If they are 
present large quantities of the vegetation should be gathered and placed 
in pans with a small amount of water. As planarians cannot endure 
stagnant water, they soon come to the top and may be picked off. 

Other species, notably the large dark forms such as Euplanaria agilis 
and E. dorotocephala, live in springs and spring-fed streams and marshes. 
Their presence may be discovered by baiting a suspected habitat with a 
piece of raw meat placed along the edge, not in the current. After 15 or 
20 minutes, the piece of meat should be turned over and planarians, 
if present, will be found attached to the under side. The entire habitat 
should then be baited with meat. Fresh raw beef is best and should be 



Planariidae 155 

cut into pieces 1 or 2 inches wide and 2 or 3 inches long. Such pieces 
should be distributed throughout the edges of the habitat so as to rest 
partly in the water, partly above the water. At intervals of 15-20 min- 
utes the pieces should be picked up with a long forceps and shaken off 
into a jar of water. With a few trials of this sort the best spots in the 
habitat are soon discovered and all of the meat may be moved to such 
spots. In preparing such collections for transport back to the laboratory, 
they should be washed free of bits of meat by several rinsings, and the 
jars filled not more than % full with fresh clean water from their habitat. 
A depth of not more than an inch of planarians should be allowed 
to a pint jar. In bringing them in, care must be taken to avoid high 
temperatures. The jars must not be set on the floor of a car which is 
apt to become hot from the engine. 

Baiting with meat is usually ineffective with pond habitats and com- 
monly succeeds only with species which live in flowing water. Some 
species, however, even in flowing water, respond poorly to this method 
and must be picked from stones and water weeds by hand. Among the 
species which the author has personally seen or knows may be collected 
successfully by baiting are: Euplanaria agilis, E. dorotocephala, Fonti- 
cola velata, and Phagocata gracilis. Euplanaria maculata and Procotyla 
fluviatilis usually respond poorly to meat baits. 

Laboratory maintenance. Planarians of practically any species may 
be kept successfully in the laboratory in glass or crockery containers or 
enameled pans. These should be darkened by means of suitable covers. 
Treated city waters are not very suitable, but most species will live in 
such water for a considerable time. Spring or well water is desirable. 

Those species mentioned above as collectable by baiting with meat are 
also the ones which may be kept most easily and successfully in the 
laboratory. They are fed two to three times a week on beef liver (pig 
liver is not suitable). Before feeding, the water in the pan should be 
lowered to a depth of several inches. The liver should be cut into long 
thin strips and disposed over the bottom. The pan is then covered 
and left undisturbed for 2 or 3 hours, after which the liver is removed, 
the pans thoroughly rinsed, and filled with fresh water. Even if the 
animals are not fed, the water should be changed two or three times 
weekly as planarians are very susceptible to fouling of the water. All 
food fragments should be carefully removed. E. dorotocephala, E. agilis, 
and Curtisia foremanii are very easily kept by this method for long 
periods of time in the laboratory; Fonticola velata and Phagocata gracilis 
may also be maintained on liver, although not so well as the first-named 
species. In place of liver, pieces of earthworm, clam, etc., may be used; 
some forms prefer such food. Yolk of egg drawn out with a dropper 
into a strand on the bottom has been employed successfully. 



156 Phylum Plat helmint he s 

E. maculata does not feed very well on beef liver and is less easy to 
maintain in laboratory culture than the preceding species. It is neces- 
sary with this species to grind the liver in a meat grinder and wash it 
thoroughly in running water. Small bits of such washed liver will usually 
be accepted as food. This species, however, in general prefers pieces 
of invertebrate flesh or liver or other flesh of tadpoles, fish, etc. 

Procotyla fluviatilis is the most difficult of our common species to keep 
under laboratory conditions as it will eat nothing but live prey, such as 
daphnids, amphipods, and isopods. It will sometimes accept blood clots, 
but in general it is impractical for laboratory purposes. 

Sexual material. Zoologists at times desire sexually mature material 
for class or experimental purposes. In general those species which re- 
produce extensively by asexual methods are seldom found in the sexual 
state; this statement applies to E. dor otocephaly, E. agilis, and Fonticola 
velata. E. maculata and its various varieties are commonly sexual 
throughout the U. S. in the summer time and numerous egg capsules will 
be found on the under side of the stones in the habitat of this species. 
It is a curious fact, however, that E. maculata is apparently never sexual 
in some localities or regions while always sexual in the summer in others. 
Sexual specimens will continue to lay eggs under laboratory conditions. 

Species which do not reproduce asexually are commonly in the sexual 
state at some definite season of the year. For Procotyla fluviatilis, which, 
owing to its transparency, is our most suitable species for preparing slides 
showing the reproductive system, the time of sexual maturity extends 
from September into the winter or even, in some localities, into spring. 
Fonticola morgani (=Planaria truncata) is sexual in summer, as is also 
Polycelis coronata of mountain streams of the northwestern U. S. 

The only species which may be depended upon to lay egg capsules 
regularly under laboratory conditions is Curtisia foremanii (=Planaria 
simplissima) . This species occurs throughout the Atlantic coast states, 
may easily be cultivated in the laboratory on beef liver, and will lay egg 
capsules continuously for a long period. The young soon grow up to 
sexual maturity and also lay in their turn so that a continuous supply of 
capsules is assured with this species. 

Class Trematoda 

THE PARASITIC FLATWORMS 

H. W. Stunkard, University College, New York University 

REPORTS on culture methods for different species of parasitic flat- 
^ worms, similar to those described for free-living invertebrates, can 
not be made, because at the present time there is no known culture 



Trematoda 157 

method by means of which a parasitic flatworm may be maintained in 
artificial media. Attempts to grow the parasites in vitro have resulted 
in failure, largely because there is no adequate knowledge of their meta- 
bolic requirements. Their physiology has been studied very little and 
the factors which determine host-parasite specificity are quite unknown. 
The basis of the relationship is chemical and the adjustment has de- 
veloped gradually during a long period of association. Accordingly, the 
only course of procedure is to maintain these worms in or on appropriate 
hosts. The life cycles of most species consist of two or more successive 
generations which may infest different host species. Certain parasites 
manifest very close host-parasite specificity while other may complete 
their development in a variety of different hosts. All members of a 
natural family follow a similar course of development and it has be- 
come clearly evident that types of life cycle are closely correlated with 
phylogenetic and systematic relations of the worms. A brief account 
is here given of attempts to culture the trematode, Cryptocotyle lingua, 
and the cestode, Crepidobothrium lonnbergi. C. lingua was selected be- 
cause this species manifests little host-parasite specificity, and C. lonn- 
bergi because it is relatively common and, being a parasite of cold blooded 
hosts, may be studied at room temperature. 

The writer (1930, 1932) has reported attempts to culture Cryptoco- 
tyle lingua and Crepidobothrium lonnbergi. The metacercariae of 
C. lingua were washed in dilute seawater and placed in an isotonic salt- 
dextrose medium. As a result of a series of experiments it was determined 
that a pH of 6.8 is the optimum hydrogen-ion concentration for the sur- 
vival of the worms. Specimens remained alive in this medium for 12 
days, the medium being changed daily. During this time the young 
worms did not develop; on the contrary they slowly diminished in size 
and at the end of the experiment were only about % as large as when 
removed from their cysts. It seems that the tissue of the body was utilized 
in metabolism and that the specimens actually starved to death. When 
kept at 38 , the worms lived only 6 days, but this result is significant, 
since sexual maturity is attained in the vertebrate host in about 6 days. 
Keeping the worms under reduced oxygen pressure did not appreciably 
alter the degree of activity or time of survival. The worms may live for 
long periods of time in solutions from which the oxygen has been re- 
moved. 

To supply accessory food substances, veal was digested and the result- 
ing extract was filtered, adjusted to a pH of 7, and sterilized. Various 
amounts of this material were added to the salt-dextrose solution to form 
a culture medium. With the addition of protein material, the media 
rapidly disintegrates as a result of bacterial growth. The worms did not 
develop at room temperature. In one experiment the worms were put in 



158 Phylum Plathelminthes 

nutrient fluid for 3 hours in the morning and for 3 hours in the afternoon, 
and during the remainder of the day were maintained in the salt-dextrose 
solutions. By this method the young worms were kept alive for 6 days in 
the incubator and for 14 days at room temperature, but they did not 
grow and apparently lived no longer in the veal broth than when the 
products of protein decomposition were absent. 

Specimens of Crepidobothrium lonnbergi were removed from the in- 
testine of Necturus and washed in sterile Ringer's solution. They were 
then transferred to a sterile isotonic salt-dextrose solution. The medium 
was modified by the addition of different amounts of Hottinger broth 
prepared by the digestion of veal, and a series of cultures were prepared 
with pH values from 6 to 8. The cultures which varied around pH 7.3 
seemed most favorable. The worms were kept in small petri dishes at 
room temperature and transferred to new media every 12 hours. In one 
experiment, young specimens were kept alive for 32 days. During this 
time they increased 3 to 4 times in length and the terminal portion of the 
strobila became definitely segmented, but the proglottids were abnormal 
and sterile. Addition to the media of salt extracts of the intestinal mu- 
cosa, pancreas, and liver of Necturus, sterilized by filtration, did not 
appreciably alter the rate of growth or time of survival. The exclusion 
of free oxygen by anaerobic culture methods did not affect the result. 
Fresh serum from Necturus was definitely toxic to the worms. 

Bibliography 

Stunkard, H. W. 1930. The life history of Cryptocotyle lingua, with notes on the 

physiology of the Metacercariae. J. Morph., 50:143. 
1932. Attempts to grow cestodes in vitro. J. Paras. 19:163. 

Order monopisthodiscinea 

EPIBDELLA MELLENI 

Theodore Louis Jahn, State University of Iowa 

Occurrence. Epibdella melleni is ectoparasitic on the eyes and epi- 
dermis and sometimes in the gill and nasal cavities of numerous marine 
fishes of the order Acanthopteri. It is believed to be a West Indian 
species, but it now occurs in the New York, Chicago, and Philadelphia 
public aquariums. A list of susceptible and of non-susceptible fishes was 
given by Jahn and Kuhn ( 1932 ) , and this has been checked and extended 
by Nigrelli and Breder (1934). 

Life History. The anatomy of the adult and the complete life history 
of the species were described by Jahn and Kuhn (1932). The eggs are 
tetrahedral and are shed singly into the seawater. These may fall free 
of the fish or may be caught on the gills, scales, etc., by means of filaments 



Monopisthodiscinea 159 

and may accumulate in large numbers in the gill and nasal cavities. In 
5-8 days ciliated larvae about 225 microns in length are hatched. These 
swim rapidly, and some eventually become attached to susceptible fishes. 
Development into the adult is direct. 

Collection and culture. Apparently the organisms may be cultured 
in any balanced or well aerated closed-system aquarium which contains 
susceptible fishes, and the problem present in the public aquariums 
mentioned above is how not to culture rather than how to culture them. 
However, after infection an immunity is developed by certain species 
which makes the continual introduction of new hosts advantageous. This 
is discussed by Nigrelli and Breder (1934). In mild infections the 
cornea is attacked and sometimes destroyed. Loss of eyes due to 
secondary bacterial invaders may follow. In heavy infections the epi- 
dermis may be considerably injured, and the scales may be removed from 
large areas of the body. Over 2,000 worms have been found on the body 
of a single fish. Severe infections usually result in death of the host. 
At the New York Aquarium treatment of infected fishes consists of 
dipping in "sol-argentum" or similar substances or of raising the density 
of the seawater by addition of salt (Nigrelli, 1932). 

In the work of Jahn and Kuhn the adults and the attached larval stages 
were obtained by scraping mucus from the eyes and body surface of in- 
fected fishes with a sharp scalpel. The mucus was transferred to stender 
dishes. In about 10 minutes the worms became attached to the bottom 
of the dishes, and the mucus was pipetted off and the seawater renewed. 
The process of egg laying, and the movements of the digestive system, 
etc., were observed with a dissecting microscope within a few hours 
after collection. The eggs were removed immediately after laying and 
kept in fresh seawater which was changed several times a day until hatch- 
ing occurred. The free-swimming larvae were isolated with a pipette. 
In aquarium systems which contain a filter, the eggs and larvae may be 
found in the filter chambers in considerable numbers. 

This species offers a very good source of live demonstration material 
for the life history of monogenetic trematodes, and the above methods 
seem advisable for the investigation of any monogenetic life history which 
is similar to that of Epibdella. The application of these methods to other 
species should be of special interest, for there are no other life his- 
tories known for the order Monopisthodiscinea. The scarcity of 
observations on the life histories of members of this group is probably due 
to the scarcity of well kept closed-system salt water aquaria containing 
susceptible fishes. Under natural conditions, of course, the free- 
swimming larvae could be collected only rarely in plankton nets, and 
this material ordinarily would not be sufficient for the study of life 
histories. 



160 Phylum Plat helminthes 

Bibliography 

Jahn, T. L., and Kuhn, L. R. 1932. The life history of Epibdella melleni Mac- 

Callum 1927, a monogenetic trematode parasitic on marine fishes. Biol. Bull. 

62:89. 
Nigrelli, R. F. 1932. The life-history and control of a destructive fish parasite 

at the New York Aquarium. Bull. N. Y. Zool. Soc. 34:123. 
Nigrelli, R. F., and Breder, C. M., Jr. 1934- The susceptibility and immunity of 

certain fishes to Epibdella melleni, a monogenetic trematode. /. Parasit. 20:259. 

Class Cestoidea 

INTERMEDIATE STAGES OF CESTODES 

Reed O. Christenson, University of Minnesota 

THE most available cestode for general laboratory use is Taenia 
pisiformis which occurs in the body cavity, about the mesenteries, 
or in the liver of native rabbits as a cysticercus, and comes to maturity 
in the intestinal tract of dogs and related carnivores. By autopsy of a 
dozen or so cottontail rabbits, ample material may usually be obtained 
to establish a permanent laboratory supply. 

The parasites, in the infective larval stage, appear as small (pea- 
sized) vesicles enclosing a head and encased in an adventitious con- 
nective tissue capsule. Occasionally worm-like motile stages are found 
free in the body cavity, or the parasites may be seen as regular white 
blotches in the liver. These are developmental stages and are not suit- 
able for infection. 

Encapsulated cysts are fed with meat to tapeworm-free dogs. In 
about 5 or 6 weeks an examination of the feces will disclose the terminal 
segments discharged from the worms. These contain ova composed of 
the hexacanth embryos covered by the striated embryophore as char- 
acteristic of the true taeniae. The ova are easily demonstrated by 
macerating a segment in water, mounting a drop on a slide, covering and 
studying microscopically. To infect rabbits, the ova are added to a moist 
bran mash and fed directly to young animals. 

When the ova enter the alimentary canal the hexacanths free them- 
selves of their covering, penetrate the host tissues to the blood stream, 
and are carried to the liver. Here they migrate about in the tissues 
for a while and by the 24th day have come to the surface where they 
appear as regularly contoured white blotches. They migrate again, 
leaving the liver, and come to lie free in the peritoneal cavity as elongate, 
worm-like bodies. These ultimately assume a spherical shape and be- 
come encapsulated, thus reaching the infective stage. 

The behavior of the cysticercus upon liberation is of some interest. 
They may be studied by teasing away the connective tissue capsules, care 



Cestoidea 161 

being taken not to injure the caudal vesicle of the worms, and placing 
them in warm saline solution. Many of the parasites will evert their 
heads and crawl about actively in the container, using their hooks and 
suckers as they would when liberated in the intestine. These specimens 
make ideal whole mounts either with or without staining. 

The behavior of the hexacanth embryo as it leaves its embryophore 
may be demonstrated by using Hymenolepis nana of rats and mice. 
Terminal segments of the parasite are chilled in a refrigerator (about 
20° C.) for a few hours and are then teased apart in a drop of saline 
on a slide and covered. The slides are gradually warmed to about body 
temperature and then studied microscopically. This procedure stimulates 
many of the hexacanths to action and they may be seen jabbing their 
minute hooklets at the inner membrane of the embryophore. Occa- 
sionally they entirely free themselves. 

Cysticercoid stages of the smaller cestodes are often difficult to ob- 
tain. They may be found in naturally infected intermediate hosts in areas 
of high frequency of the adult parasite in its primary host. Our method 
of obtaining these stages is to kill a series of rats from various localities 
to find the highest incidence and greatest intensity of infection with 
Hymenolepis diminuta. When this is ascertained, beetles (Tenebrio 
molitor) which serve as the common intermediate host are sought and the 
adults teased apart in a saline filled Syracuse crystal under a binocular 
dissecting microscope. The cysticercoids fall away from the surround- 
ing tissues and may be identified by their inverted heads, broadly oval 
bodies, and elongate caudal processes. As high as 20% natural infection 
has been found in this way, with as many as 28 cysticercoids recovered 
from a single beetle. 



Phylum VI 

Nemertea 



METHODS FOR THE LABORATORY CULTURE OF 

NEMERTEANS 

Wesley R. Coe, Yale University 

NEMERTEANS are found under extremely diverse environmental 
conditions. Many species are strictly littoral, living in the mud and 
sand, or beneath stones between tide-marks, or in shallow water along the 
seashore. Others are found upon the sea-bottoms at moderate depths. 
Still others swim freely suspended as bathypelagic organisms 1,000 meters 
or more beneath the surface of the great oceans, while a few freshwater 
species are to be found in pools and streams in all temperate and tropical 
regions or in water-holding leaves of tropical plants. Several species 
have acquired terrestrial habits, living in moist earth in tropical or semi- 
tropical lands, whence they have been accidentally transported to green- 
houses in all parts of the world. 

The nemerteans have a distinct advantage over many other groups 
of invertebrates for laboratory culture because, although they are essen- 
tially carnivorous, the individuals of many of the small littoral species 
are able to live for a year or more without other food than that which 
may be obtained from their own tissues. Several of these may reproduce 
asexually in the meantime, but sexual reproduction does not occur under 
such conditions. 

LITTORAL SPECIES 

The slender Lineus socialis of the Atlantic coast, or the similar 
L. vegetus of the Pacific coast, found beneath stones between tide-marks, 
is easiest to culture since it requires only a covered dish of seawater with 
a bottom layer of pebbles and sand mixed with a little mud, freshly 
brought from the nemeateans' natural habitat. This material will 
supply the necessary small Crustacea, nematodes, and other small in- 
vertebrates to keep the animals in good condition for a year or two if 
the water lost by evaporation is replaced from time to time. Asexual re- 
production will occur frequently and egg clusters may be deposited in 
late winter or early spring, but only if the water is kept below 15 C. 

162 



Heteronemertea 163 

For a study of asexual reproduction or regeneration each worm or tiny 
fragment may be kept in a separate, cork-stoppered, 6- or 8-dram vial 
containing clean seawater only, changing the water every day or every 
few days. 

The smaller representatives of each of the orders, except the parasitic 
Bdellonemertea, may be kept in the same manner but somewhat less 
successfully. The larger forms naturally require vessels with a generous 
supply of water and a thicker layer of bottom material. 

FRESHWATER SPECIES 

Species of Prostoma (Stichostemma) are found adhering to the leaves 
of aquatic plants in pools and quiet streams in nearly all parts of the 
United States. They thrive in aquaria containing a good growth of 
vegetation if supplied with minute Crustacea, nematodes, turbellarians, 
and other small organisms, but the water must be free from bacterial 
decomposition. The plants will require a thin layer of soil on the bottom 
of the aquarium. Excessive evaporation is prevented in arid rooms by 
partially covering the aquarium with a pane of glass. Prostoma thrives 
best at temperatures of about 20 C. Egg clusters are deposited along 
the sides of the aquarium at all seasons of the year. 

TERRESTRIAL SPECIES 

Greenhouses having soil, temperature, and moisture conditions suit- 
able for the cultivation of ferns and moisture-loving tropical plants are 
suitable for the culture of Geonemertes. The worms are merely allowed 
to burrow in the moist soil of the pots or boxes containing the plants. 
They are protandric, with sexual reproduction only. Egg masses are 
deposited on or near the surface of the soil. 

OBTAINING EGGS FOR EXPERIMENTAL PURPOSES 

As stated in the preceding sections, the smaller species, such as Lineus 
and Prostoma, deposit their eggs in gelatinous clusters when the nemer- 
teans are cultured in the laboratory, and in both these genera the eggs 
develop readily in the jars or aquaria without special precautions. For 
studies on maturation and fertilization and especially for experimental 
work where very large numbers of ova are necessary, the larger littoral 
forms, such as Cerebratulus or Micrura, are easily secured in the breeding 
season. 

A very large female Cerebratulus lacteus which occurs in the intertidal 
zone along the entire Atlantic coast of the United States may produce at 
one time upwards of fifty million eggs. These are mature in early spring 
along the Carolina coasts, during May and June in Long Island Sound, 
in July at Woods Hole, and during July and August in Massachusetts 



164 Phylum N em ertea 

Bay and on the coast of Maine. Several species of the same genus on 
the Pacific coast from San Diego to Alaska furnish equally beautiful ova. 

The female Cerebratulus may be kept in a vessel of clean seawater for 
3 weeks or more, if the water is changed daily and the temperature is held 
at about io° C. But the eggs are less suitable for experimental work 
after the first week. They are spawned spontaneously under such condi- 
tions. 

To obtain large numbers of ova or young larvae it is necessary to free 
the eggs from the body. This is best done by taking a small fragment of 
the body, placing it in a dish of cool seawater and making a longitudinal 
slit with sharp knife or scissors on the dorsal surface on each side of the 
median line. The muscular contractions of the fragment will soon force 
the ripe ova into the water. After a few minutes as many eggs as are 
wanted are drawn into a pipette and expelled into a dish of clean seawater. 
They are thereby washed free of most of the body fluids. 

The eggs on reaching the water still have the germinal vesicle intact 
and are not yet ready for normal fertilization. Immediate fertilization 
usually results in polyspermy. The stimulus of the water soon results in 
the formation of the first polar spindle which proceeds to the metaphase 
and then rests. This stage is reached in 10 to 30 minutes after the egg 
reaches the water, the time depending both on the temperature and on 
the ripeness of the eggs. The eggs are then ready for fertilization. 

To obtain the sperm a small fragment of a male (which may be dis- 
tinguished from the female by its brighter color) is placed in a dish of 
clean seawater and a puncture made through the dorsal body wall. 
The sperm ooze out in a dense mass. A surprisingly minute quantity of 
this, when expelled from a pipette into a dish containing the ova, will 
suffice for complete fertilization. If larvae are desired the fertilized eggs 
must be provided with a generous supply of clean, cool seawater. 

REARING LARVAE 

Prostoma and other hoplonemerteans develop directly into young 
worms without the intervention of a free-swimming larval stage such as 
is characteristic of Cerebratulus. In Lineus an intermediate condition 
known as the Desor larva occurs. These larvae require no special feed- 
ing, but the pilidium larva of Cerebratulus or of Micrura may be reared 
to the adult form only by the most careful attention. 

The difficulty lies in providing suitable nourishment during the 20 or 
more days which the swimming larva requires before metamorphosis is 
completed. Small diatoms and other minute algae may be supplied 
daily, with frequent changes to clean seawater. A more reliable method is 
to feed regularly from a pure culture of the smallest obtainable diatoms. 
The temperature of the water should not exceed 20 C. 



Heteronemertea 165 

METHOD FOR STUDIES ON REGENERATION 

In nearly all species of nemerteans the body quickly restores a missing 
posterior extremity. In some forms this ability is limited to the posterior 
half of the body, but in others the head and a small portion of the foregut 
region, or even the head alone, without any part of the alimentary canal, 
can regenerate all the missing parts. Anterior regeneration is usually 
limited to the head in front of the brain but a few species, particularly 
those of the genus Lineus, can reproduce the entire body in miniature 
from any small fragment except the minute piece anterior to the brain. 
Even a small sector of a fragment, if it contains a tiny piece of the lateral 
nerve cord, is likewise endowed with the capacity for complete regenera- 
tion and reorganization. Curiously enough, individuals of some species 
live longer in captivity with the head removed than with the entire body 
intact, for the reason that the decapitated body is less restless. 

Operations for regeneration experiments are usually performed with- 
out the use of an anesthetic, although the head may be removed if neces- 
sary. The worm is placed in a small pool of cool water upon a beeswax 
plate having a suitable concavity. Under the binocular dissecting micro- 
scope the desired cuts may be made with a cutting blade sharply ground 
from a curved needle. The fragment is then placed in a vessel or vial 
of clean water. 

In some species the fragments are less restless, and consequently re- 
generate better, if a few bits of shells or small pebbles or sand grains are 
placed in the vial or dish. This procedure may sometimes make the 
difference between the success or failure of the experiment. Food may be 
supplied after the mouth and digestive tract become functional. 



Phylum VII 

Nemathelminthes, Class Nematoda 





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RECOVERING INFECTIVE NEMATODE LARVAE FROM 

CULTURES 

G. F. White, U. S. Bureau of Entomology and Plant Quarantine 

THE method outlined here for recovering infective nematode larvae 
from cultures makes use of the often-observed fact that toward the 
close of their free-living period the larvae migrate from the medium in 
which they have been growing. The simple apparatus used traps the 

migrating larvae in 
water (White, 1927). 
Convenient and 
sufficient equipment 
consists of crystal- 
lizing dishes 125 to 
150 mm. in diameter, 
watch glasses slightly 
larger than these di- 
mensions, petri dishes 
100 to 125 mm., 
test tubes 20 by 
150 mm., filter papers 
9 to 12 cm., a spatula 
with a 4-inch blade, a test tube rack, a three-quart boiler with cover, 
animal charcoal, and sterile water. Brief steaming in the covered vessel 
suffices for the disinfection that is needed. 

The charcoal and feces with water added are mixed properly and con- 
veniently in one of the larger watch glasses and transferred to the half 
of a petri dish, with a moistened filter paper covering the bottom. Sterile 
water is poured into a crystallizing dish to cover the bottom and into it is 
placed the half petri dish containing the culture. A watch glass is used 
as a cover, the apparatus (Fig. 45) after labeling, is placed for incubation 
preferably where a high humidity may be maintained. 

Many of the larvae on approaching the third larval stage migrate from 
the culture and are trapped in the water surrounding the petri dish. In 

166 



Fig. 45. — Apparatus used for culturing nema- 
tode larvae, a, Crystallizing dish; b, Petri 
dish with charcoal-feces mixture; c, watch-glass 
cover. Water surrounds the petri dish equal 
to about one half its depth. 



Nematoda 167 

collecting them the watch glass cover is removed and the half petri dish 
with the charcoal culture is lifted out, preferably with forceps to avoid 
infestation. The water containing the larvae is poured from the crystal- 
lizing dish into a test tube which is then placed in the rack. The worms 
soon gravitate to the bottom of the tube, after which the water above 
may be pipetted off, leaving the larvae concentrated. The apparatus with 
the charcoal mixture may then be reassembled and steamed. 

A number of modifications of the apparatus and the method may be 
made to meet the worker's special needs. When there is but a small 
amount of culture it is well to use the half petri dish with the bottom up. 
A Syracuse watch glass or the top of a Coplin jar serves well in place of 
the petri dish. Room temperature, especially in summer, may be substi- 
tuted for the more constant one of an incubator. 

A modified form of the apparatus has been used to reduce somewhat 
the amount of fungous growth in the culture when this seemed desirable. 
An aluminum pan of the diameter of the crystallizing dish, with the in- 
clined side perforated, is placed beneath the watch glass cover and sup- 
ported by the edge of the dish. Into the pan is poured a few cc. of an 
aqueous solution of formalin. A 15% solution has been employed suc- 
cessfully, but the optimum strength should be determined by each worker 
to meet his own needs. 

While using the method one soon learns of its limitations and its ad- 
vantages. The larvae are recovered from the cultures in relatively clean 
water. Only infective forms are obtained. A considerable leeway is per- 
mitted as to the time larvae may be collected from the apparatus. The 
first larvae trapped may be poured off, more water added, and the appa- 
ratus reassembled for later migrations. Frequently additional ones may 
be had by transferring the charcoal culture to the Baermann apparatus 
(Darling, 191 1). 

The method was found to be convenient and efficient in studies on and 
in the diagnosis of hookworm and other nematode infestations (Fulleborn, 
192 1 ) and in studies on the biology of the causal parasites (Cort, Stoll, 
and Grant, 1926). 

In making studies on the migration of nematode larvae, Looss (1911) 
trapped them in water but apparently did not employ the observation 
in devising a routine method for obtaining larvae from cultures. 

Among those who have taken advantage of the migrating tendency of 
larvae in devising methods suitable for their studies is Darling (1911), 
who used Syracuse watch glasses in the center of which he placed 3 to 
5 cc. of stool and added sterile water until the feces were surrounded 
with fluid. The worms for study were taken from this margin of water. 
Fulleborn (1921) also made use of this habit, employing agar plates. 
The charcoal-feces mixture was placed on this medium in the center of 



1 68 Phylum N emathelminthes 

the petri dish. The infective larvae migrating from the culture over the 
agar cause their trails to be inoculated with bacteria. By the growth of 
these the courses taken by the worms are readily observed. 

Bibliography 

Cort, W. W., Stoll, N., and Grant, J. B. 1926. Amer. J. Hyg. Monog. Ser. 

7:19. 
Darling, S. T. 1911. Strongyloides infections in man and animals in the 

Isthmian Canal Zone. J. Exper. Med. 14:1. 
Dove, W. E. 1932. Further studies on Ancylostoma braziliense and the etiology 

of creeping eruption. Amer. J. Hyg. 15:664. 
Fulleborn, F. 192 1. Nachweis von Ankylostomum durch Plattenkot-cultur. 

Vorl. Mifteilg. Arch. f. Schiffs und Tropenhyg. pp. 121-123. 
Looss, A. 1911. The anatomy and life-history of Anchylostoma duodenale Dub. 

Reeds, of Egypt Govt. Sch. of Med., Cairo, IV. 
White, G. F. 1927. A method for obtaining infective nematode larvae from 

cultures. Science 66:302. 
White, G. F., and Dove, W. E. 1928. The causation of creeping eruption. 

J . A.M. A. 90:1701. 
1929. A dermatitis caused by larvae of Ancylostoma caninum. Arch. 

Dermal, and Syph. 20:191. 

Order hologonia, Family trichinellidae 

REARING TRICHINELLA SPIRALIS 

Reed O. Christenson, University of Minnesota 

THE difficulty of isolating developmental stages of Trichinella spiralis 
has restricted its laboratory use often to mere demonstration of the 
cysts and, more rarely, the adults. It is feasible, however, with a little 
experience to have on hand for laboratory use ample material of all stages 
of the parasite in the living condition. The main difficulty lies in ob- 
taining the original infection. Usually a routine examination of wild 
rats about abattoirs or packing houses will yield material. If this source 
fails, infected animals may often be obtained from other laboratories. 

When trichinosed tissue is obtained a number of animals should be fed 
to maintain a supply. Rats are ideal, but guinea pigs and rabbits may be 
used with good results. Cats are more difficult to handle but are ideal 
animals in which to maintain the infection over long periods of time. 

Shortly after the ingestion of infective meat the larvae are liberated 
from their cysts by the action of the digestive juices. This may be done 
experimentally by placing infected tissue in an artificial gastric juice com- 
posed of water, 1,000 cc; hydrochloric acid, 10 cc; scale pepsin 
(U. S. P.), 2.5 grams; and sodium chloride, 5.0 grams. Small quantities 
of finely cut meat are stirred up in the mixture and incubated at 38 ° to 
40 C. for 18 hours. 



Trichinellidae 169 

Under natural conditions larvae liberated in the stomach soon make 
their way to the intestine where they mature in upwards of 48 hours. 
The isolation of intestinal forms may be achieved with little difficulty. 

Nearly grown rats are maintained without food for 48 hours and are 
then each fed separately a thumb-sized piece of heavily trichinosed tissue. 
At the desired time (after 48 hours to get early adults) an animal is 
killed with chloroform and the intestine removed to a stender dish con- 
taining warm saline solution. Previous workers have opened the intestine 
with enterotomy scissors and have searched for the worms in the mucosa. 
Our method is to cut the intestine, without slitting it, into lengths of 
about two inches. These are held at one end with forceps and the con- 
tents stripped out by compressing a second pair tightly against the in- 
testine near the held end and drawing them downward. The parasites 
and the slight amount of debris present are caught in a crystal containing 
saline. The worms may be strained free from the debris using a 40-mesh 
copper screen, or they may be picked out with a finely drawn pipette. 
Using this method the author has recovered more than 5,000 adult para- 
sites from a single rat. 

It is difficult to obtain larvae from the circulating blood. They are 
small and considerable quantities of blood must be collected, centrifuged 
and studied before the larvae are found. Fiilleborn accomplished this by 
centrifuging the blood with a mixture composed of 5% formalin (95 cc), 
glacial acetic acid (5 cc), and concentrated alcoholic gentian violet 
(2 cc). The larvae were recovered from the sediment. Using this 
method Fiilleborn isolated larvae from the blood of the ear and heart until 
the 20th day after infection. 

It is quite instructive to present, at least for demonstration, larvae of 
trichinae migrating between the muscle fibers. By killing heavily in- 
fected animals 18 to 20 days after the initial feeding and compressing 
small bits of diaphragm between glass plates held together by screw 
clamps, the larvae near the edge of the tissue are forced free and may 
be studied. 

The parasites have come to lie in the muscle in their encysted stage 
2 1 days after infection, and at this point they are again infective. The 
connective tissue capsule has begun to form and the worms are relatively 
quiescent. By 35 to 40 days the connective tissue sheath is quite pro- 
nounced ; this is the best time to obtain encapsulated trichinae. 

By maintaining infected animals, rabbits or cats, for three or four years 
the ultimate calcified stage may be obtained. 



1 70 Phylum N emathelminthes 

Order telogonia, Family ancylostomidae 

THE GROWTH OF HOOKWORM LARVAE ON PURE 
CULTURES OF BACTERIA* 

OVA of the dog hookworm, Ancylostoma caninum, have been ob- 
tained free from feces and sterilized. These sterile ova have been 
inoculated onto agar cultures of various bacteria, and the larvae have 
hatched normally and grown to the infective stage with bacteria as their 
sole source of food. 

The method employed for freeing the ova from the feces consists in 
thoroughly mixing up about 25 grams of freshly passed feces from a 
heavily infested dog in 500 cc. of water. The mixture is then washed 
through a series of copper-wire sieves ranging up to a mesh of 100 wires 
to the inch. The larger particles in the feces are caught in the sieves but 
the ova readily pass through with the filtrate. This filtrate is allowed to 
stand in a large sedimenting cone for about an hour while the ova and 
heavy debris settle to the bottom. The supernatant fluid is then poured 
off; the sediment is transferred to a 50 cc. centrifuge tube and repeatedly 
washed with water, the solid matter being thrown to the bottom each 
time by centrifuging at a speed of 1,000 revolutions per minute. After 
the supernatant fluid from the washing has become practically clear, 
saturated salt solution is poured into the tube and the contents are again 
centrifuged at the same speed. This time the ova come to the surface and 
may be collected by removing the surface film with the open end of a 
piece of large glass tubing. If the material is centrifuged four or five 
times, a majority of the ova present may be recovered. If much solid 
material comes to the surface with the ova, it may be necessary to refloat 
the ova in saturated salt solution a second or even a third time in order 
to get rid of the foreign material. This method is tedious and time- 
consuming, but if the feces of a heavily infested dog are used, large quanti- 
ties of ova may be obtained almost entirely free from fecal material. 

Ova collected by this method may be sterilized by treatment with a 
5% antiformin solution in 10% formalin. From 10 to 50% of the ova 
remain viable after this treatment. The ova are washed several times 
with sterile distilled water and are then ready for inoculation onto the 
agar cultures. During the process of sterilizing and washing, the ova 
are best kept in a sterile 50 cc. centrifuge tube closed with a cotton plug. 

Cultures were made up in 250 cc. Erlenmeyer flasks stoppered with cot- 
ton plugs, and consisted of 20 cc. of ordinary bacteriological agar which 
had been diluted with three parts of water. The flasks were autoclaved 
and inoculated with bacteria 24 hours before the ova were introduced. 

* Reprinted, with slight changes, from Science 69:74, 1929, by Oliver R. McCoy, 
School of Hygiene and Public Health. 



Ascaridae 171 

Since at ordinary room temperature the ova do not hatch for an addi- 
tional 36 hours, there was a heavy growth of bacteria in the cultures by 
the time the larvae were ready to begin feeding. The sterile ova were in- 
troduced into the flasks in 1 cc. portions of an aqueous suspension, several 
thousand ova usually being put in each flask. 

In experiments so far carried out larvae have grown to the infective 
stage in the normal period of about 7 days on pure cultures of Bacillus 
coli, B. subtilus, B. prodigiosus, B. lactis aerogenes, Staphylococcus au- 
reus, Spirillum metchnikovi, S. rubrum, and Micrococcus citreus. Ova 
which were put on plain agar without bacteria hatched normally and lived 
for as long as ten days, but did not grow. If bacteria were then introduced 
into the flasks, the larvae grew to the infective stage. 

Not all bacteria are suitable food for hookworm larvae, since they 
failed to grow on cultures of Bacillus pyocyaneus and Sarcina lutea, and 
growth was very much retarded on cultures of B. cereus and B. mega- 
therium. Larvae, however, grew normally on a mixed culture of Bacillus 

cereus and B. coli. 

Bibliography 

Looss, A. 191 1. The anatomy and life history of Anchylo stoma duodenale Dub. 
Part II. The development in the free state. Reeds, of Egypt Govt. Schl. of Med. 
4:163. M. E. D. 



Family ascaridae 



CULTURING EGGS OF THE FOWL NEMATODE, 
ASCARIDIA LINEATA 

J. E. Ackert, Kansas State College 

THE eggs may be secured from either live or dead worms, but cul- 
tures from live worms are much to be preferred. The anterior end 
of the worm is excised and the internal organs pressed into a sterile 
petri dish ; the uteri are isolated and transferred to another sterile dish. 
At various points the uteri are punctured for the liberation of eggs with 
characteristic light centers which are known to be fertile. The portions 
of uteri containing fertile eggs are transferred to a final sterile petri dish, 
and the eggs pressed out and covered with sterile distilled water. To 
prevent bacterial or fungous growth 4 or 5 drops of 2% formalin are 
added. The cultures are incubated approximately 3 weeks at 27 to 
30° C 

For best results the culture dishes should be opened daily and agitated 
to facilitate entrance of oxygen. If patches of fungus appear they 
should be removed. As the eggs usually adhere to the bottom of the 
dish, the medium must be poured off and a new supply added. 



172 Phylum N emathelminthes 

The amount of dilute formalin that may be added to inhibit fungous 
growth depends upon the permeability of the egg envelopes, which differ 
in the various species of nematodes. 

THE LIFE HISTORY OF THE SWINE KIDNEY WORM* 

UNDER laboratory conditions, at a temperature of about 2 6° to 
2 7 C, the preparasitic stages of the development of Stephanurus 
dentatus were completed in from 5 to 6 days. Eggs obtained from gravid 
females and cultured in water or on a charcoal and feces mixture 
hatched in from 24 to 48 hours, and the larvae reached the first lethargus 
about 24 hours after hatching. The second lethargus was reached about 
48 hours later, and the infective stage, that of the third stage larva, was 
usually attained about 24 hours after the onset of the second lethargus. 
Low temperatures have been found to retard the development of the eggs 
and larvae, and at temperatures sufficiently low not only was development 
arrested but the vitality of the eggs and larvae was destroyed. 

When infective larvae of S. dentatus were placed on the scarified skin 
of pigs or when they were injected subcutaneously infection resulted, the 
course of development being similar to that which followed the adminis- 
tration of larvae by mouth. 

Stephanurus has been reared experimentally in guinea pigs in which 
animals they have been found to attain a considerable growth and 
development. 

M. E. D. 

Family oxyuriformidae 

CULTIVATION OF A PARASITIC NEMATODE 

William Trager, Rockefeller Institute for Medical Research 

THE nematode parasite (Neoaplectana glaseri) of the Japanese beetle 
(Popillia japonica) was cultured by Glaser (1931, I93 2 )- Plates of 
the culture medium are prepared by mixing in a sterile 5M>-cm. petri dish 
about 8 cc. of melted veal infusion agar (pH 7.4) with 2 cc. of a 10% 
dextrose solution. The surface of the cooled medium is flooded with a 
concentrated water suspension of a pure culture of baker's yeast and the 
plate is incubated 24 hours at room temperature. The yeast growth 
inhibits later bacterial growth and serves as food for the nematodes. 
These are obtained from diseased Japanese beetle grubs and, after having 
been sedimented and washed three times in water, are placed on the yeast- 
culture plate. The culture is incubated at room temperature. 

♦Abstracted from an article in Science 70:613, 1929, by Benjamin Schwartz, U. S. 
Bureau of Animal Industry. 



Anguillulidae 173 

The nematodes pass through one generation every 4 or 5 days, and 
must be subcultured every 2 weeks, by which time most of the yeast in 
the culture has been consumed. 

Most of the nematode strains died out after the seventh or eighth 
transfer, i.e., after 14 to 16 weeks in culture, during which time from 21 
to 32 generations had been produced. The cultures died out because, 
although the females of the last culture passage were normal in size and 
shape, their ovaries failed to mature and no young were produced. 
However, by permitting worms from the sixth transfer to infect Japanese 
beetle grubs and to go through several generations in their natural hosts, 
worms could be recovered which could again be grown in culture through 
7 or 8 transfers. 

Recently (Glaser, unpublished) it has been found possible to culture 
the worms indefinitely by adding small amounts of powdered ovarian 
substance to the culture plates. 

Free-living soil nematodes do not grow at all under the conditions 
suitable for Neoaplectana. [See p. 174.] 

Bibliography 
Glaser, R. W. 1931. The cultivation of a nematode parasite of an insect. Science 

73:6i4- 
1932. Studies on Neoaplectana glaseri, a nematode parasite of the 

Japanese beetle (Popillia japonica) . N. J. Dept. of Agric. Circular No. 211. 

CULTURING PARASITIC NEMATODE LARVAE FROM 
SILPHIDS AND RELATED INSECTS 

C. G. Dobrovolny, University of Michigan 

THE larvae of these oviparous nematodes are parasitic in the body 
cavity of silphids and the mature worms are free-living. The worms 
were successfully reared in cultures. 

The larvae were freed from the body cavity of the silphids and cul- 
tured in tap water. Most of the worms died in too dilute and in too 
concentrated cultures. It was further observed that the mortality of 
larvae cultured in Syracuse watch glasses which were kept full of water 
was 100%. Best results were obtained by keeping the larvae in culture 
media of wet sand and macerated beetles. 

Family anguillulidae 

ANGUILLA ACETI* 

Anguilla aceti is a very satisfactory subject for type study in biology, 
zoology, and parasitology classes. It may be procured at any time of 

♦Abstracted from an article in Science 74:390, 1931. by George Zebrowski, Buck 
Creek, Indiana. 



1 74 Phylum N emathelminthes 

the year from a corner grocery by asking for bulk cider vinegar. It will 
live indefinitely in the laboratory if transferred to fresh vinegar every 
two weeks. Since it is viviparous and transparent, all stages of develop- 
ment may be examined in utero. Anatomical details, such as the ali- 
mentary tract, nerve ring, spicules and sperms of male, uterus and 
uterine development in female, and all young and intermediate stages 
can be seen with a 4 mm. objective. Placed in a well slide and projected 
on a daylight screen with a micro-projector, many anatomical features 
may be shown on the screen under suitable magnification. 

M. E. D. 



ARTIFICIAL CULTIVATION OF FREE-LIVING NEMATODES* 

Asa C. Chandler, Rice Institute 

THE free-living nematodes of soil and of water may be studied to 
great advantage by culturing them on ordinary nutrient agar plates. 
A single isolated adult female of Rhabditis sp., placed on an agar plate 
with a drop or two of dirty water to supply a bacterial growth, in a period 
of 5 days will produce thousands of offspring which swarm all over the 
plate. In 10 days the offspring will number many thousands, — males, 
females, eggs, and young in all stages of development. 

For class demonstration of soil nematodes, a student may place a 
small quantity of soil, preferably manured soil, in a piece of gauze or in 
a fine sieve, and wash it in a beaker of warm water; in a few minutes 
the majority of the nematodes present will have fallen to the bottom of 
the beaker. If a drop or two of water from the bottom of the beaker is 
then placed on the surface of a nutrient agar plate, and the plate covered 
and left at room temperature for from 5 to 10 days, an enormous number 
of nematodes of several species will usually be found. The majority of 
the individuals move about on the surface of the agar, but some burrow 
into it also. The movements on the agar are sufficiently impeded so that 
they may be watched after the fashion of a slow-moving picture. The 
swallowing of bacteria and fungus spores, the excretion of waste matter 
from the anus, and every detail of locomotion may be observed under 
ideal conditions. Species of Rhabditis and Cephalobus, and others not 
positively identified, have been cultured in this manner. The method 
suggests a great range of possibilities in the way of study and experi- 
mentation, e.g., on foods, effects of hydrogen-ion concentrations and of 
chemical substances, resistance to desiccation, tropisms, effects of various 
modifications in environment on rate of reproduction and development, 
etc. The extremely rapid rate of reproduction and ready inbreeding also 
suggests possibilities in genetic experiments. 

♦Recast of article in Science 60:203, 1924. 



A nguillulidae 175 

References 



For the culture of free-living nematodes see also p. 173. 
For the culture of fresh water nematodes see p. 136. 
For uses of nematodes as food see pp. 162 and 203. 



Phylum VIII 

Trochelminthes, Class Rotatoria 



A CULTURE MEDIUM FOR HYDATINA SENTA* 

Josephine C. Ferris, University of Nebraska 

BRING to the boiling point in ioo cc. of water i gram of urea crystals, 
i gram of dried blood, and i gram of dried ox gall. Filter and add 
3 cc. of this triple solution to ioo cc. of tap or rain water. This solution 
is very satisfactory for pedigree cultures of this rotifer in watch glasses if 
fed on Polytoma cultures. [See p. 6 1.] If the Polytoma is cultured in 
a bone meal and hay solution it should be washed at least twice by 
centrifuging and decanting. 

References 

For the culture of Hydatina asplanchna see p. 210. 
For the culture of Philodina see p. 143. 

Class Gastrotricha 

METHOD OF CULTIVATION FOR THE GASTROTRICHA 

Charles Earl Packard, University of Maine 

MANY specimens of Lepidoderma squamatum were reared in a 
0.1% malted milk solution for a period of 22 months. 

One half gram of Horlick's malted milk was dissolved in 500 cc. of 
clear water from a spring-fed river tributary, usually unfiltered, and 
boiled for 5 minutes. This was left exposed for 24 hours before being 
used. Sometimes the solution was filtered, though usually not. Ani- 
mals were raised in rectangular, covered refrigeration dishes in quantity 
lots and in depression slides singly or in mass. In most cases Lepido- 
derma squamatum adapted itself readily. When once a culture was 
started, a small amount of fresh solution was added daily to keep up a 
supply of animals. Excess fluid was drawn off to prevent overflowing. 

Lepidoderma continuum, and a species of Chaetonotus, a genus char- 
acterized by cuticular spines, were also raised in the same medium for 
several weeks, adaptation being less easily accomplished with these forms. 

*See Biol. Bull. 63:442, 1932. 

176 



Gastrotricha 177 

They survived and reproduced in small numbers, however. Small 
amoebae, some Heliozoa, many tiny colorless and pale green flagellates, 
Entosiphon, Halteria, and several types of rotifers thrived at certain 
periods. Malted milk as a successful medium for the growth of Para- 
mecium is mentioned by Pixell-Goodrich in Lee (1928). 

Bibliography 
Lee, A. Bolles. 1928. The Microtomist's Vade-Mecum, 9th ed. p. 411. 



Phylum IX 

Bryozoa, Class Ectoprocta 



BUGULA FLABELLATA AND B. TURRITA 

Benjamin H. Grave, De Pauw University 

THE breeding season of these Bryozoa at Woods Hole, Mass., extends 
from June i or June 15 to Nov. 1. 

The complete life history of Bugala flabellata is readily observed 
because of the ease with which larvae are obtained and kept under 
laboratory conditions until colonies of several individuals are established. 
The larvae, which resemble the trochophores of annelids, are given off 
by the parent colonies at dawn. To obtain them in abundance sexually 
mature colonies should be collected late in the afternoon and placed in 
dishes of seawater. These are left over night near a window. Larvae 
issue from the colonies early in the morning and continue to be liberated 
from 5 to 10 a.m. They promptly swim to the lighted side of the dish, 
where they may be taken in a pipette and transferred to fresh dishes of 
seawater for study. Finger bowls or stender dishes are recommended 
for the purpose. 

At first the larvae, as indicated above, are strongly and positively 
heliotropic but later a change occurs and they become negative in their 
response to light. Still later they make permanent attachment to the 
walls of the dish and proceed to develop into colonies. The larva re- 
quires no food because it has no digestive tract, but as soon as attachment 
occurs the containing dish should be placed in running seawater to secure 
aeration and food for the developing colony. 

The swimming period of the larva is about six hours in duration. The 
first individual of the colony becomes a complete feeding polypide within 
two days and a colony of eight is established in one week if conditions 
are favorable. The colonies under natural conditions bud rapidly and 
become sexually mature in 1 month. They continue to grow for ap- 
proximately three months which is the approximate duration of life of 
colonies established early in the summer. Colonies established late in 
the summer live over winter. 

The rate of growth of colonies may be observed under natural con- 
ditions from the establishment of the colony to maturity by placing 

178 



Ectoprocta 



179 



wooden floats in the region where the species occurs normally. Wooden 
crosses measuring three or four feet in length have proved excellent for 
the purpose (Fig. 46). They were tied to a dock and allowed to float on 
the surface. So constructed they do not turn over in storms and thou- 
sands of colonies soon appear on their lower surfaces; from these samples 
may be taken for study from day to day. Sexually mature colonies 
which liberate great numbers of larvae daily may be grown in this way. 
Bugnla turrita has the same life history but it grows in somewhat 
different situations. The larvae show interesting structural differences, 
especially in the presence of visible pigmented eye spots, but they respond 





J— J 




1 


3?B 


1 


y 







Fig. 46. — Type of wooden cross used to grow colonies of Bugula 
flabellata and other sessile organisms, such as barnacles, as- 
cidians, and hydroids. (Construction: spruce or pine, 2x4; 
three feet long with cross bar 1x6, two and one half feet long, 
sunk flush with the surface.) 

to the same treatment, their behavior being the same in all respects, 
larvae of B. turrita may be liberated in the afternoon.* 



The 



CULTURING FRESHWATER BRYOZOA 

Mary Rogick, College of New Rochelle 

FRESHWATER Bryozoa are among the most common of aquatic 
invertebrates. Ponds, rivers, streams, lakes, harbors, bays, and 
quarries abound with specimens. Bryozoa may readily be collected 
throughout the spring, summer, and fall in either one or two of the fol- 
lowing stages: colony, statoblast, or hibernaculum. Hibernacula are pro- 
duced by colonies of Pottsiella and Paludicella while statoblasts are 
produced by the Plumatellas, Cristatella, Pectinatella, Lophopodella. 
Lophopus, and Fredericella. Houghton and Marcus admit the possibil- 
ity of over-wintering in the case of Fredericella and Lophopus under 
favorable conditions. 

Collection of Bryozoan forms is simple. Floating statoblasts may be 
obtained by skimming over the surface of the water with a silken dip-net. 

* A more complete account of the behavior of the larva and the rate of growth of the 
colony may be found in /. Morph. 49:355, 1930. 



180 Phylum Bryozoa 

Attached or enclosed statoblasts, hibernacula, and colonies may be ob- 
tained by scraping with a scalpel or sharpened spatula the under side of 
rocks, lily pads, submerged objects such as boards and rubbish, floating 
objects such as sticks and logs, leafy aquatic vegetation such as Vallis- 
neria, Elodea, Potamogeton, Ceratophyllum, Myriophyllum, or Scirpus, 
and by scraping shells of Unionidae, Astacus (Abricossoff, 1925) and cer- 
tain tortoises (Annandale, 1912). Records exist for Bryozoans from 
various depths. Hand collecting is resorted to in the shallows, but at 
depths beyond three feet the use of double rakes or dredge is necessary. 

Some of the enemies of Bryozoa which may occur in the collections and 
which should be carefully excluded, are planarians, various gastropods, 
insect larvae such as chironomids and caddis worms (Hydropsyche, 
et al.), oligochaete worms, small crustaceans, and arachnids. 

The problem of culturing freshwater Bryozoans has been a rather 
difficult one. They are very voracious. They feed upon diatoms, 
desmids, Oscillatoria, Ciliata, Flagellata, some rhizopods (Arcella), small 
rotifers, etc. Marcus mentions the following forms as of use in feeding 
Bryozoa: Euglena, Colpoda, and Chlamydomonas. Brown has been 
successful in using "Geha" fish food, size 000, in culturing Plumatella, 
keeping the colonies alive and reproducing for four weeks. 

In culturing Bryozoa shallow dishes or finger bowls should be used if 
one wishes to observe the organisms very frequently. In larger aquaria 
they may not be easily examined. The water in the finger bowls should 
be replaced every 2 or 3 days with fresh pond water or with tempered tap 
water. The following diets were tried out on Lophopodella carteri and 
found to be unsuccessful: Paramecia, Euglenae, green algae, dried and 
powdered Elodea, Vallisneria, algae, malted milk, nutritive broth (de- 
hydrated), and fish food (not powdered). As a last resort, greenhouse 
water which contained a great amount of organic debris or detritus was 
tried. This was collected from around the stems and bases of large 
aquatic plants which had been planted in the greenhouse tanks. Some of 
the mud and decaying vegetation at the base of the plants was included. 
Planaria, chironomids and other offenders were removed from the culture 
as far as possible. The debris was removed and new material added 
every two or three days. This proved to be an ideal medium for rearing 
colonies. Lophopodella colonies collected in Lake Erie in August and 
September, 1932, and cared for in the laboratory released statoblasts in 
October and November. These statoblasts hatched in November, De- 
cember, January, and February, giving rise to small colonies. These new 
colonies gave rise to statoblasts in March and April. Some of these 
statoblasts germinated in April (1933). The culture dishes were kept 
in the laboratory under ordinary room conditions of light and tempera- 
ture (approximately 22 C). 



Ectoprocta 181 

Frequently in some of the cultures there appeared a gelatinous coating 
over the bottom of the dish, covering the colonies. If this were not 
removed, the colonies would die from suffocation, lack of food, etc. This 
scum occurred more frequently in flat stender dishes than in finger bowls 
and Pyrex glassware. When such a condition occurs, the Bryozoa must 
be transferred to another dish, the scum must be removed from them with 
dissecting needles, scalpel, or other suitable instrument. They should 
then be placed in a sterilized finger bowl in fresh pond water. 

Lophopodella will tolerate stagnant and polluted water to a surprising 
extent. A number of colonies were placed in an aquarium with two 
large stones which had a small amount of dirt and algae on them, some 
broken lily pads, and a small amount of Elodea, to which was added 
lake water (about 5 gallons). At the end of ten days, the water was a 
cloudy yellow in color, turbid, and gave off a very bad odor. The col- 
onies however were in excellent condition. Pectinatella, Cristatella, and 
some of the other Bryozoans could not tolerate such conditions so well. 

Bibliography 
Abricossoff, G. G. 1925. The materials for the fauna of the Bryozoa of the 

government of Moscow. Arb. Biol. Sta. Kossino, Lief. 2:81. 
Annandale, N. 1912. Fauna Symbiotica Indica, No. 3. Polyzoa associated with 

certain Gangetic tortoises. Rec. Ind. Mus. 7:147. 
Brooks, C. M. 1929. Notes on the statoblasts and polypids of Pectinatella mag- 

nifica. Proc. Acad. Nat. Sci. Phila. 81:427. 
Brown, C. J. D. 1933. A limnological study of certain fresh-water Polyzoa, etc. 

Trans. Amer. Micr. Soc. 52:271. 

1934- Internal budding: with suggestions for a laboratory study of fresh- 
water Polyzoa. Ibid. 53:425. 

Davenport, C. B. 1904. Report on the fresh-water Bryozoa of the United States. 

Proc. U. S. Nat. Mus., 27:211. 
Harmer, S. F. 1913. Polyzoa of Waterworks. Proc. Zool. Soc. London, pp. 426. 

1 93 1. Recent work on Polyzoa. Proc. Linn. Soc. London, Session 143, 

part VIII, p. 113. 

Henchman, A. P. and Davenport, C. B. 1913. Clonal variation in Pectinatella. 

Amer. Nat. 47:361. 
Houghton, W. i860. Note on Fredericella, etc. Ann. Mag. Nat. Hist. (3) 6:389. 
Kraepelin, K. 1885. Die Fauna der Hamburger Wasserleitung. Abh. Naturwiss. 

Ver. Hamburg, 9. 
Marcus, E. 1925. Bryozoa in P. Schulze's Biologie der Tiere Deutschlands. Lief. 

14, Teil 47, pp. 1-46. 

1934. Uber Lophopus crystallinus (Pall.). Zool. Jahrb., Abt. Anat. 

Ont. Tiere, 58:501. 

Rogick, M. D. 1934. Studies on fresh-water Bryozoa, I. Trans. Amer. Micr. Soc. 

S3:4i6. 
Schodduyn, R. 1923. Materiaux pour servir a l'Etude biologique des Cours d eau 

de Flandre Francaise. Ann. biol. lac. 12:121. 
Tanner, V. M. 1932. Ecological and distributional notes on fresh-water sponges 

and Bryozoa of Utah. Utah Acad. Sci. 9:113- 
Williams, S. R. 1921. Concerning "larval" colonies of Pectinatella. Ohio J. Sci. 

21:123. 



Phylum XIII 

Annelida, Class Polychaeta 



Order polychaeta errantia, Family nereidae 

NEREIS LIMBATA 

Benjamin H. Grave, De Pauw University 

Nereis limbata, which occurs abundantly in Eel Pond at Woods Hole, 
is the one species in this vicinity which is known to have a distinct and 
unmistakable lunar periodicity in spawning. Eggs may usually be had 
in great abundance roughly from the full moon until new moon during 
all of the summer months and not to any considerable extent at any other 
time. 

The eggs and spermatozoa are extruded at night from 9 to 10 p.m. as 
the sexually mature worms swim at the surface of the sea. As the 
males and females come into contact with each other they are stimulated 
to expel their gametes vigorously. The stimulus, however, is chiefly 
chemical rather than physical. At this time in the month the body 
cavities of the worms are distended with eggs or spermatozoa and after 
they are expelled nothing but the ghost of a worm remains. 

METHOD OF COLLECTING 

A small dip net and a lantern or flashlight are needed in collecting. 
The worms are attracted to light and may be dipped up and placed in 
suitable dishes of seawater. Females should be kept separate from 
males, otherwise they spawn at once. 

METHOD OF SECURING EGGS 

Select a distended female, place her in a clean dish of seawater, and 
with scissors cut across her body to allow the eggs to escape. In the 
same way cut a male in two in a dish containing 25 or 50 cc. of seawater. 
After washing the eggs once or twice by pouring off the water and re- 
filling the dish with fresh seawater, add three or four drops of sperma- 
tozoa and agitate gently. Within five minutes after insemination the 
eggs extrude a jelly in which they lie embedded. The eggs are thus 

182 



Nereidae 183 

readily fertilized artificially and development proceeds. Usually 100% 
of the eggs cleave and, barring accidents, nearly all develop into normal 
embryos. 

CARE OF CLEAVING EGGS 

The cleaving eggs require no further attention except that the water 
should be changed several times during the next 12 or 15 hours. In the 
meantime the eggs cleave and acquire cilia. After 24 hours the embryos 
may be separated from the jelly and transferred to a clean dish of sea- 
water either by pouring or by using a wide-mouthed pipette. Care should 
be exercised to get rid of all decaying organic matter as soon as possible, 
and this must be accomplished within 36 hours after insemination or 
before. Cleaving eggs that are allowed to develop without frequent 
change of water usually develop abnormally or die. 

SCHEDULE OF DEVELOPMENT 

The embryo becomes an early gastrula 12 or 15 hours after fertiliza- 
tion, with the four large macromeres constituting the principal part of 
the endoderm. It is ciliated and rotates in the jelly. It is a late gastrula 
after 24 to 30 hours, the rate of development depending upon the tem- 
perature. Between 36 and 48 hours the larva is a trochophore, at first 
spherical but later somewhat elongated. On the third day the first three 
segments of the worm body are completed and no additional segments 
are added for several days although the embryo increases in size. It is 
possible to keep the embryos until other segments grow but to do so 
requires special feeding methods. The Nereis larva is unusually hardy 
and easily cared for. The trochophores and early segmented larvae are 
active swimmers but as the ciliary mechanism becomes inadequate they 
depend more and more upon wiggling and creeping. 

CARE OF LARVAE 

The egg of Nereis is large and well supplied with yolk and oil so that 
the larvae require no feeding during the first five or even seven days of 
development. They are easily cared for because after two days they 
have a tendency to settle to the bottom on one side of the dish and may 
be transferred to a clean dish of seawater with a wide-mouthed pipette. 
This should be done once per day or more frequently in hot weather. 
After five days they may be fed upon diatoms but if it is desired to keep 
them for several weeks it is best to transfer them to a large cylindrical 
balanced aquarium, containing a dense culture of developing diatoms 
which adhere to its sides. E. E. Just reared Nereis megalops to maturity 
in such a jar of diatoms. The original stock of diatoms came from the 
Fisheries Laboratory at Beaufort, N. C. 



1 84 Phylum A nnelida 

Bibliography 

Lillie, F. R., and Just, E. E. 1913. Breeding habits of Nereis limbata at Woods 

Hole, Mass. Biol. Bull. 24:147. 
Just, E. E. 1922. On rearing sexually mature Platynereis megalops from eggs. 

Amer. Nat. 56:471. 
Wilson, E. B. 1892. Cell lineage of Nereis. J. Morph. 6:361. 

A METHOD FOR REARING NEREIS AGASSIZI 
AND N. PROCERA 

John E. Guberlet, University of Washington 

THE writer has reared two species of Nereis to sexual maturity in the 
laboratory by the use of the following method. One species, Nereis 
agassizi, was reared to sexual maturity during each of two consecutive 
years and in the third attempt the larvae were maintained for a period of 
nearly 14 months but due to unfortunate circumstances did not reach 
sexual maturity. A second species, Nereis procera, was successively cul- 
tured in the laboratory for a year. At the end of that period the worms 
had reached sexual maturity. 

While the worms are "swarming" at the surface in their seasonal 
spawning it is a comparatively easy matter to capture both males and 
females. Better results may be obtained if the males and females are 
kept separate during capture and transfer to the laboratory. They 
should spawn in separate dishes in sufficient seawater to keep them well 
covered and to maintain a fairly even temperature. A small amount of 
water containing spermatozoa is added to the eggs and thoroughly mixed. 
Care should be taken not to use too great excess of spermatozoa. The 
dish containing the fertilized eggs should be allowed to stand for a few 
minutes and then be emptied into a larger container (battery-jar) con- 
taining 2 or 3 liters of seawater. This jar is placed in running water or in 
a suitable location to maintain a fairly constant temperature. The degree 
of temperature best suited to a particular species would seem to be that 
of the environment from which the adult worms were taken. After the 
eggs have settled to the bottom, as much of the water as possible should 
be siphoned off to remove the excessive spermatozoa from the culture and 
fresh seawater should be added. Polar bodies begin to appear after 1 
to 1% hours and cleavage starts after 2% hours. The cleavage rate is 
usually fairly rapid and movement of the larvae begins in 12 to 15 hours. 
The trochophore stage is reached in about 36 to 48 hours and larvae with 
three pairs of setigerous appendages appear in between 3 and 4 days. 
It is highly important that the temperature be kept constant. The 
water should be changed daily and agitated at least once each day to 
provide aeration. When the larvae have developed setigerous append- 
ages they will soon be provided with jaws and are then ready to begin 



Serpididae 185 

feeding. The larvae will consume very readily small diatoms which are 
then added to the culture. For this purpose it was found that species 
of Navicula and Nitzschia [See p. 34.] were of suitable size. They were 
eaten in large quantities. The larvae grow rapidly and when they have 
developed 6 or 8 segments they form mucous tubes within which they 
hide themselves. When the larvae reach an age and size having 10 or 
12 segments they will consume other food in addition to diatoms. At 
this time Ulva and brown kelp, such as Nereocystis, may be added to the 
diet. At this time they may consume other small worms and any mate- 
rial that they can devour. The worms extend themselves almost their 
entire length from their tubes and reach for food. 

After the young worms develop tubes it is not necessary to change the 
water so frequently. A change every 2 or 3 days is sufficient, and at a 
later date, once a week will suffice, provided there is plenty of green 
vegetation in the water. 

Several important factors must be kept in mind: 1) there must not 
by overcrowding of organisms; 2) fresh water must be provided to 
allow for a sufficient supply of oxygen; 3) care must be taken to prevent 
contamination; 4) the temperature must be kept within a limited range; 
and 5) a sufficient quantity of suitable food must be provided at all 
times. 

Order polychaeta sedentaria 
Family serpulidae 

HYDROIDES HEXAGONUS 

Benjamin H. Grave, De Pauw University 

Hydroides hexagonus, a serpulid worm, secretes a calcareous tube 
which adheres firmly to shells of mollusks, stones, and wooden structures. 
The breeding season at Woods Hole opens between June 10 and June 15 
and closes between October 15 and November 1. 

METHOD OF OBTAINING EGGS AND SPERMATOZOA 

To obtain eggs or sperm it is advisable to remove the worms from 
their calcareous tubes and place them in stender dishes or Syracuse watch 
crystals filled with seawater, one worm per dish. When so treated they 
always spawn immediately if they are sexually mature. The sexes are 
separate and the gametes, whether eggs or spermatozoa, are carried free 
in the coelomic cavities from which they are extruded through nephridio- 
pores located along the sides of the body. During the early part of the 
breeding season over 50% of the spawned eggs are immature and under- 
size. Later on they are nearly all mature and fertilizable. Maturation 



1 8 6 Phylum A nnelida 

does not take place until the spermatozoon enters the egg, the germinal 
vesicle being indistinctly visible through the yolk. 

EMBRYOLOGY 

It is desirable to delay insemination of the eggs for half an hour after 
spawning has occurred because the spermatozoa are quite immobile 
when first extruded. Under the stimulus of seawater they gradually be- 
come activated but are relatively inactive at best. After allowing time 
for activation, remove the eggs by means of a pipette to a fresh dish of 
seawater and add four or five drops of sperm. 

After fertilization the eggs develop within ten hours into actively 
swimming gastrulae and therefore rise from the bottom. They may now 
be poured into a clean dish, thus discarding the eggs which failed to 
develop. 

Within 24 hours the embryos have become transparent trochophore 
larvae which continue to swim actively. They remain in the trochophore 
stage of development for 10 days or two weeks showing little external 
change except a slight slender outgrowth at the posterior end which 
constitutes the beginning of the worm body. They feed readily upon 
diatoms by means of a ciliary mechanism and may be kept indefinitely 
under laboratory conditions. 

Because they at no time settle to the bottom, it is difficult to keep the 
water changed but they remain in good condition if poured daily into 
clean dishes discarding the bottom layers. They have a tendency to 
collect at one side of the dish and may be transferred to clean dishes of 
seawater by means of a pipette, but this method involves the loss of 
many embryos. Zeleny (1906) has reared them to metamorphosis in 
aquarium jars. 

POST EMBRYONIC DEVELOPMENT 

The trochophore finally develops a slender worm body, settles per- 
manently, and secretes a calcareous tube. By placing mollusk shells or 
stones in a cage and sinking them in shallow water which is known to 
contain breeding Hydroides worms it is possible to secure many young 
worms and study their further development. Studies of this character 
carried on at Woods Hole for several years have shown that they become 
sexually mature in seven or eight weeks before they are half grown. 
They become fully grown in two years. Most of the worms ordinarily 
collected are only one year old and it is likely that many if not most of 
them die during the second year. 

The shell of an average-sized worm after one year's growth, measures 
65 or 70 mm. in length and 3 or 4 mm. in widest diameter. The largest 
worms may reach 120 mm. in length and 5 mm. in greatest diameter. 



Sabellariidae 187 

Bibliography 

Grave, B. H. 1933. Rate of growth, age at sexual maturity and duration of life 
of certain sessile organisms at Woods Hole, Massachusetts. Biol. Bull. 65:380. 

Hatscheck, B. 1885. Entwicklung der Trochophora von Eupomatus uncinaliis 
(Serpula uncinata) . Arch. Zool. Inst. Wien. 6. 

Shearer, C. 1911. On the development and structure of the trochophore of Hy- 
droides uncinatus. Quart. J. Micr. Sci. 56 : 543. 

Zeleny, C. 1906. The rearing of serpulid larvae. Biol. Bull. 8:308. 



Family sabellariidae 

SABELLARIA VULGARIS 

Alex B. Novikoff, Brooklyn College 

Sabellaria vulgaris is a sedentary polychaete found along the Atlantic 
coast from North Carolina to Cape Cod (Pratt, 1935). The observations 
recorded here are based on experiments with worms dredged from a depth 
of sixty to eighty feet in the waters of Tarpaulin Cove in Vineyard 
Sound, near Woods Hole, Massachusetts.* The animals are abundant 
and are easily collected in this rather limited area. 

The worms live within sand tubes which they build on stones, empty 
sanddollar and oyster shells, and occasionally on Bryozoa nodules and 
Limulus shells. Males and females are about equal in number in the 
collections. The sexes can be recognized externally only in those fully 
mature animals which contain a large number of either eggs or sperm. 
The abdominal segments, which are greatly distended, are dense white 
in the male and a decided pink in the female. The eggs or sperm are 
shed almost immediately after the animals are removed from their tubes. 
The number of gametes shed is greatest from those animals which have a 
pronounced color in the abdominal segments, but even animals which 
show neither the distinct white nor pink color may shed abundantly. 

Animals collected throughout the greater part of the summer showed 
no apparent differences in the condition of their gametes. Eggs from 
worms dredged at irregular intervals during the periods, August 22 to 
September 7, 1934 and June 24 to September 9, 1935, developed nor- 
mally in more than ninety-five per cent of the cases.** The same high 
percentage of normal development usually followed from eggs of animals 
that had been kept in aquaria with running seawater for as long as 
nine weeks. 

♦This work was carried on at the Marine Biological Laboratory, Woods Hole, Massa- 
chusetts. 

** Verrill (1874, p. 317) states that "eggs are laid in May and June," and Waterman 
(1934, p. 98) says that "the normal shedding time of May and June is followed by a 
second but shorter period extending from about August first to fifteenth." 



1 88 Phylum Annelida 

OBTAINING EGGS AND SPERM 

Uninjured animals are most easily obtained using the following pro- 
cedure: (i) remove the sand tube from the shell or rock, (2) break 
away enough of the tube to make the head and tail visible within, 
(3) carefully force the animal from the tube by inserting a blunt probe 
into the head end of the tube. 

The sex of the animal removed from the tube is ascertained, if the 
color of the abdominal segments is not definitely white or pink, by 
placing it in a few drops of seawater until it begins to shed. The sperm 
usually pour out of the male in dense white clouds. The masses of eggs 
shed by the female break up in the water into small groups. The male is 
placed into four drops of seawater until it has completed shedding. The 
female is placed into a finger bowl containing about 200 cc. of clear sea- 
water. After it has shed for a few seconds, it is moved to a new position 
in the finger bowl and allowed to shed in that place for the desired length 
of time, depending on the number and kind of eggs desired. The first 
eggs to be shed are not generally used because of the possibility of their 
having been on the surface of the animal while exposed to the air. 

When preparing the eggs for fertilization, it is best to allow them to 
remain in the seawater for about fifteen minutes. Towards the end of 
that time, the sperm suspension is prepared. One drop of the sperm shed 
into the four drops of seawater is diluted with four drops of seawater, 
and one drop of this diluted suspension is then added to a finger bowl of 
seawater (about 260 cc). The eggs are drawn up with a narrow medi- 
cine dropper and transferred into this suspension.* 

The original sperm suspension (made by allowing the male to shed in 
four drops of seawater) may still be used after three to four hours and 
longer if evaporation of the water is prevented. The eggs may be 
fertilized immediately after shedding, but it is best to allow them to 
remain in the water for about fifteen minutes before they are fertilized, 
if eggs of the same stage of development are desired. 

EARLY DEVELOPMENT 

Just after being shed, the egg is very irregular in shape and contains 
a large clear germinal vesicle. A few minutes later, the egg begins to 
round out, a clearly visible membrane is raised from the surface, and the 
large germinal vesicle breaks down to form a spindle which extends across 
not quite half the diameter of the egg. The average diameter of the 

* This procedure is quite different from that of Waterman (1934) who placed the male 
and female together in a finger bowl of seawater. The method used by me has the fol- 
lowing advantages: 1 — The eggs obtained are more nearly alike with respect to the stage 
of development. 2 — The eggs are obtained free from the debris which clings to the bodies 
of the animals. 3 — The use of the small quantity of sperm gives higher percentages of 
normal development. Polyspermy, which might otherwise be encountered, is avoided. 



Sabellariidae 189 

rounded egg is fifty-six micra; the membrane, usually wrinkled, is about 
twelve micra from the egg surface. Fine protoplasmic processes are 
seen to extend from the egg surface as the membrane is raised. The 
elevation of the membrane is apparently due to the swelling of a rather 
dense jelly situated between the egg surface and the membrane. This 
jelly is ordinarily invisible but it can be demonstrated by removing the 
outer membrane (see p. 190) and placing the eggs in a dense suspension 
of Chinese ink. 

The process of fertilization has been described by Waterman (1934) 
and will be further discussed by me in a future publication. The sperm 
attaches to the egg within the first minute after mixing the eggs and 
sperm, but the exceedingly large fertilization cone may not be com- 
pletely retracted for a period of twelve to fourteen minutes. Some time 
during the course of sperm entry, the protoplasmic processes are with- 
drawn into the egg. The egg is too opaque to allow for the direct ob- 
servation of any internal processes other than those connected with the 
spindle area. 

Preceding the formation of the first and second polar bodies there is a 
distinct flattening of the egg at the region where the polar bodies will 
be extruded. The first polar body separates at about nineteen to twenty- 
three minutes after fertilization; the second comes off nine to eleven 
minutes later. At fifty to fifty-five minutes after fertilization, a large 
anti-polar lobe is formed and the cell divides into two (sixty-five to 
seventy minutes) . After the division the polar lobe goes into the CD cell. 
Ten to fifteen minutes later, a smaller lobe is given off at the anti- 
polar end of the CD cell and the second cleavage occurs. At the com- 
pletion of the division the lobe flows into the D cell. The first set of 
micromeres, which are only slightly smaller than the basal cells, comes 
off in the usual dexiotropic fashion. During the course of this division 
a third lobe forms in the D cell. The later cleavage has not yet been 
described. 

At five and a half hours, the developing embryo begins to move about 
by means of cilia and at eight hours the apical tuft and prototroch are 
well formed. The larvae live fairly well on a diet of the diatom Nitzs- 
chia. One larva was raised, without much care, to the beginning of meta- 
morphosis (seven weeks) at which time it was fixed. Wilson (1929) has 
given a detailed description of the larvae of the British Sabellarians with 
which those of Sabellaria vulgaris agree very closely. 

The exact time relations in development and the effect of change of 
temperature on such relations have not been studied. The schedule of 
events as given above is only approximate for room temperatures 
varying from nineteen to twenty-five degrees C. 



i go Phylum Annelida 

REMOVAL OF OUTER MEMBRANE 

The outer membrane of the egg may be removed by following a slight 
modification of a procedure described by Hatt (1932). Two solutions. 
A and B, are prepared as follows: 

Solution A. — A given volume of a solution of i gm. of Na 2 CO3 
in 1000 gm. of isotonic NaCl (or KC1). This gives a solution with 

pH 9.6. 

Solution B. — 0.45 cc. of 1.0 n HC1 are added to 100 cc. of iso- 
tonic NaCl (or KG). This solution has enough HC1 so that when an 
equal amount of the solution is added to a given amount of Solution 
A the resulting mixture is at the pH of seawater (8.2). 

The eggs are placed in a Syracuse dish and as much of the seawater as 
possible withdrawn, without injury to the eggs. A small amount (three 
to four cc.) of Solution A is poured on them and the dish rotated. After 
about one minute, the eggs are allowed to settle and the solution with- 
drawn. Now a carefully measured amount (five cc.) of Solution A is 
pipetted into the dish. The eggs clump into large masses, but as the 
membranes dissolve the eggs separate and the masses break up. The 
disappearance of the egg membranes may be followed under the micro- 
scope. When they have disappeared from most of the eggs (about five 
minutes) five cc. of Solution B is quickly added. The force of the 
stream from the pipette is sufficient to mix the solutions. After the eggs 
have again settled, they are transferred to fresh seawater in Syracuse 
dishes. The bottoms of the dishes have been previously coated with 
a thin layer of agar to prevent the adhesion of the eggs to the glass 
surface. 

This treatment removes the outer membrane but the jelly still remains 
about the egg. 

Bibliography* 

Dehorne, A. 1910a. La division longitudinale des chromosomes dans la sperma- 
togenese de Sabellaria spinulosa. Compt. Rend. Acad. Sci. 150:1195. 

1910b. Le valeur des anses pachytenes et le mecanisme de la reduction chez 

Sabellaria spinulosa. Ibid. 150:1625. 

191 1. Recherches sur la division de la cellule. II. Homeotypie et Hetero- 



typic chez les Annelides polychaetes et les Trematodes. Arch, de Zool. Exp. S. 
9,1:1. 

1 913. Nouvelles recherches sur les mitoses de maturation de Sabellaria spin- 



ulosa. Compt. Rend. Acad. Sci. 156:485. 
Faure-Fremiet, E. 1924. L'oeuf de Sabellaria alveolata L. Arch. d'Anat. Micr. 
20:211. 

*The literature on the European species of this genus is included in the bibliography. I 
will shortly publish accounts of (i) the details of fertilization, (2) the later cleavage, 
and (3) the results of experiments with centrifugation and isolation and transplantation of 
blastomeres. 



Enchytraeidae 191 

Harris, E. J. 1935. Studies on living protoplasm. I. Streaming movements in 
the protoplasm of the egg of Sabellaria alveolata L. J. Exper. Biol. 12:65. 

Hatt, P. 1922. La fusion experimental d'oeuf de Sabellaria alveolata L. et leur de- 
veloppement. Arch, de Biol. 42:303. 

1932. Essais Experimentaux sur les Localizations Germihales dans l'oeuf 

d'un Annelide, Sabellaria alveolata L. Arch. d'Anat. Micr. 28:81. 

Pratt, H. S. 1935. Manual of the Common Invertebrate Animals. 

Verrill, A. E. and Smith, S. I. 1874. Report upon the Invertebrate Animals of 
Vineyard Sound and Adjacent Waters. 

Waterman, A. J. 1934. Observations on reproduction, prematuration, and fer- 
tilization in Sabellaria vulgaris. Biol. Bull. 67:97. 

Wilson, D. P. 1929. Larvae of the British Sabellarians. J. Mar. Biol. Assoc. 
30:221. 



Class O/igoc/iaeta, Family enchytraeidae 

CULTIVATION OF ENCHYTRAEUS ALBIDUS 

Raymond F. Blount, University of Minnesota Medical School 

SUCCESS in the cultivation of Enchytraeus albidus is largely de- 
pendent upon constant care and attention to small details in the 
condition of the culture. The principal things to be considered in this 
are the medium; moisture, and food. 

A dishpan is very satisfactory for a container although a tight wooden 
box may be used. The latter is not so satisfactory since air and moisture 
exchange at the sides and bottom make regulation difficult. In any case 
the use of several small cultures rather than a large one is advisable. 
There should be a cover to lessen evaporation. In addition to this the 
surface of the medium may be partially covered by small pieces of 
slate or stones, or glass jar covers. 

The medium is a light loam soil of such a character that it does not 
easily harden when dry, while on the other hand it should not be sandy. 
It should not be too great in quantity, as the worms should congregate 
in masses for breeding. A depth of 2 or 3 inches is best, for if too 
shallow it is difficult to regulate the moisture. 

The culture should be kept in a place where the temperature is such 
that multiplication is encouraged, but relatively low. The range is wide 
but the optimum is probably around 20 C. 

There should be sufficient moisture to allow free motility of the worms 
but not enough to bring them to the surface except as they may travel 
on the under side of pieces of slate or stone resting on the surface of the 
dirt. They are visible here if glass is used for this purpose. They 
should not congregate in this position however. In a rich culture the 
movement of the worms is audible. The addition of water is by 
sprinkling rather than by pouring and should be frequent and light. 



192 Phylum Annelida 

A laundry sprinkler in the neck of a bottle is useful. The regulation of 
moisture may be aided by removing the cover for a time as necessary. 
The surface of the earth should be level and pressed down, not too 
firmly, to leave no lumps above to dry or mold. The fragments of slate 
or stone on the surface aid greatly in producing an optimal condition 
beneath. There should be no collection of water in the bottom of the 
pan. If this occurs it is best to tip the pan up, with something under 
one edge, to allow the water to drain to one side so that the greater 
part of the pan may dry. If excessive water is present the earth should 
be held in place and the pan drained on its side. 

The food should be placed where the worms are found as they have 
usually congregated in favorable localities. However some should be 
scattered throughout to reach those worms which are dispersed in the 
culture. It is best to place the food in small masses as the cocoons are 
deposited on these and the young may be securely attached to them. 
Cereals, such as oatmeal, are convenient foods. Pieces of boiled potatoes 
with the skins attached are also excellent. Bread or other materials 
may be used. It is best to vary the diet. The amount should be such as 
to be consumed within a reasonable time, frequent rather than large 
feedings being needed. 

Mold may often be present but does not seem to interfere if the food 
masses are not large. Removal of surface growth and taking the cover 
off to allow short drying periods will help keep it in check. A culture 
which attracts Drosophila is often in optimal condition and a few of 
these flies are usually present. However a souring culture is to be 
strictly avoided. 

In starting a "culture the worms should be placed in only a few places 
rather than scattered throughout the dirt, and food placed with them. 
Breeding is facilitated if large numbers of worms are together. Since a 
culture usually presents cycles of abundance and scarcity of worms it is 
advisable to have several cultures if a constant supply of worms is needed. 

In securing worms for use, masses of them may be removed with 
forceps and placed in water. If small worms are desired it is best to 
remove also some dirt and food. At times they may be secured by 
washing the under side of the slate or stones on the surface. Repeated 
decanting of the water and the addition of more will wash most of the 
light dirt away and the worms may be transferred with wide-mouthed 
pipettes through several wash waters to clean them. 

It should be emphasized again that constant daily attention to the 
cultures is essential. 



Enchytracidae 193 

LABORATORY CULTURE OF ENCHYTRAEUS 

Victor Loosanoff, U. S. Bureau of Fisheries 

THESE small Oligochaetae are extensively used for feeding small 
fish and amphibians kept under laboratory conditions. Since it is 
comparatively easy to grow them and because they are excellent food for 
lower vertebrates, it is desirable to maintain an abundant supply in each 
laboratory where work on fish or Amphibia is carried on. 

To grow Enchytraeus collect rich, dark garden soil and place in a large 
dish pan. Pulverize a few milk crackers or dry pieces of bread, and mix 
this powder with the soil, which has previously been rendered moist. 
After three or four days seed the soil with Enchytraeus taken from 
another culture or obtained from a biological supply house. Add more 
cracker powder. The soil must always be kept moist but not wet. It has 
been observed by the writer that the best growth of worms is obtained 
if the culture is kept at a temperature of about 20 C. The culture must 
be kept covered with a lid but not too tightly, allowing free access of 
air. Addition of small quantities of crushed bone powder helps to keep 
the culture in good condition. Crushed crackers or bread crumbs should 
be added to the soil every 4-6 days. 

ENCHYTRAEID WORMS 

William LeRay and Norma Ford, University of Toronto 

A SPECIES of enchytraeid worm, which is used in our department, 
was originally obtained from a dealer. These worms are kept in 
boxes 1% x 2 feet in size, over the bottom of which is spread 4 inches of 
rich soil, consisting largely of decayed leaves. The worms, are fed on 
white bread soaked in milk, buried in furrows across the box. A tem- 
perature of about 6o° F. is desirable, although some strains of this worm 
will withstand higher temperatures. 

Enchytraeid worms are used to feed a great variety of animals, in- 
cluding leeches, crayfish, etc. 

References 

For the culture of Enchytraeus albidus see also p. 196. 

For the culture of several species of Oligochaetes see p. 136. 
Family Aelosomatidae 

For the culture of Aelosoma see p. 143. 
Family Naididae 

For the culture of Nais see p. 143. 

For the culture of Dero see p. 143. 



194 Phylum Annelida 

Family tubificidae 

TUBIFICIDAE 

George R. La Rue, University of Michigan 

OLIGOCHAETE worms of the family Tubificidae form an excellent 
food for many kinds of laboratory animals including planaria, 
leeches, dragonfly and damselfly nymphs, aquatic beetle larvae, and 
many fishes. A star-nosed mole kept in the laboratory ate them vora- 
ciously. The ease with which these worms may be collected in quantity 
and kept for months in the laboratory, and the avidity with which 
they are eaten, make them important laboratory animals. 

Tubificidae occur in a considerable variety of freshwater habitats. 
They may be found most readily and most abundantly in the mud, or in 
muddy borders, of streams or ponds where considerable organic matter is 
undergoing decay. When in shallow water they may often be seen with 
their tails waving in the water, their heads buried in the mud. The 
location of tubificids in soft muddy borders of streams or ponds may 
often be determined by noting numerous small casts on the surface. 
When their presence is suspected take up a small quantity of mud on a 
trowel and examine it for worms. If they are abundant determine how 
deeply they are embedded in the mud, then scrape or scoop up the layer 
of mud containing them with trowels or small shovels and put in 10- or 
12-quart pails. 

At the laboratory put the contents of a 12-quart pail in a shallow gal- 
vanized pan measuring about 15 x 12 x 3 inches deep. Set the pan on 
a drain table in a cool room and allow a very small stream of water from 
a faucet to flow continuously through a rubber tube into the pan. The 
worms will come to the surface in a few hours. During the night small 
masses of worms tend to migrate out of the pan and onto the drain table 
if the pan is overcrowded. These may be used for feeding purposes until 
the stock is reduced. 

If the mud rises in the pan because of the formation of gas, prick holes 
in it and press it down. 

In quiet rivers receiving sewage or in ponds to which manure has 
been added to increase productivity of fish food, tubificids often collect 
in masses as large as a man's fist, or larger, at the surface, either on or 
near the decaying material and frequently near the margin or even upon 
the muddy border. They occur when the water is warm and disappear 
when it gets cold in the fall. Such worm masses are collected with a large 
tea strainer, dipper, or long-handled fine-meshed dip net. In the labora- 
tory they may be put with mud and decaying vegetable matter, or with 
manure, potato, or other food material free from mud. 



Lumbricidae 195 

To feed tubificids the following materials have been used: fresh horse 
manure, baked potatoes cut in halves, boiled potatoes, butts from head 
lettuce, masses of bran, and bread. Press food down into the mud. The 
horse manure and bran should be buried in the mud. 

The worms are removed in small masses by means of a pair of forceps. 
Wash them to remove mud before feeding them to other animals. 

Reference 
For the culture of Tubificidae see also p. 142. 



Family lumbricidae 



EARTHWORMS 

Walter N. Hess, Hamilton College 

SINCE living earthworms are useful in the laboratory for demon- 
strating behavior, and since freshly killed earthworms are far 
superior to preserved specimens for the study of certain organ systems, 
especially the digestive, circulatory and excretory systems, many lab- 
oratories need a supply of living worms for mid-winter use. Brief con- 
sideration will be given to the culturing of three species. 

Specimens of Lumbricus terrestris, the common earthworm of the 
United States, must be gathered at night and preferably between 10:00 
and 12:00 o'clock during or following a drizzling, warm rain when the 
ground is thoroughly soaked. The worms come to the surface of the 
ground in large numbers at such times and may be captured easily with 
the assistance of a strong flash light or an acetylene lantern. The best 
collecting grounds are closely cut lawns where the soil is rich. 

When the worms have been collected they may be left for the re- 
mainder of the night in a cool place in a pail containing a small quantity 
of freshly cut grass. The next morning the worms should be carefully 
sorted, and all injured or abnormal specimens should be removed. If 
they are washed and placed, a few at a time, in a dish of water those that 
are injured may easily be detected. 

Earthworms feed very largely on dead and decaying leaves and, like 
chickens, they digest their food better if there is a certain amount of 
grit in their diets. We have obtained best results by keeping earthworms 
in large boxes filled about 1 2 inches deep with approximately equal parts 
of old leaves and leaf loam gathered in the woods. Under no conditions 
should heavy clay soil be used. The worms need no other food, as they 
feed on the dead leaves. The material should be kept moist but not 
saturated with water. Unless extreme care was exercised in removing 
all injured worms, the boxes should be inspected after a week and all 



196 Phylum Annelida 

dead and dying worms removed. Should it happen that the worms are 
not keeping well those that are healthy should be removed and placed in 
a fresh box of leaves and loam. 

Earthworms also keep well in very light, loamy soil. If this is used it 
is often advisable to feed the worms. Bread crumbs or corn meal make 
excellent food. The food should be moistened with water, spread spar- 
ingly over the top of the soil every 2 or 3 weeks and covered with about 
an inch of loam. Feed sparingly and not too often or the food will spoil 
and the worms may die. 

Avoid trying to keep too many worms in one box. A cubic foot of 
culture material, after it has settled, will be sufficient for about 50 worms. 
Cover the boxes with panes of glass and keep cool. Temperatures above 
6o° F. usually prove fatal. 

While cocoons of this earthworm are not easily obtainable, a few of 
them may usually be found by carefully sorting over the loamy material 
in the boxes after the worms have been stored in it for a month or so. 
The young worms emerge from the cocoons in a few weeks and thrive 
under the same treatment as that given the adults. 

The fecal earthworm, Eisenia [=Allolobopkora] joetida, which is 
much hardier than Lumbricus terrestris, and which is rather extensively 
used for experimental purposes, keeps very well in partly rotten cow and 
horse manure. The worms may usually be found here in abundance. 
They copulate and form large numbers of cocoons in the laboratory, if 
kept in containers supplied with this material. 

The small white earthworm, Enchytraeus albidus, lives well in the 
laboratory. In addition to being an excellent food for small fish and 
Amphibia it may be narcotized with chloretone and used to demonstrate 
many annelid structures with the aid of a binocular microscope. Keep 
in boxes filled with black loam and feed sparingly with bread soaked in 
milk or water. After each feeding the food should be covered with about 
an inch of loam. The material should be kept moist but not soaked. 
The worms prefer temperatures around 55°-6o° F. Higher temperatures 
should be avoided. 

Reference 

For the feeding of earthworms see also note on p. 49. 

CULTURE OF ALLOLOBOPHORA 

R. N. Dantelson, University of Minnesota 

Allolobophora sp. has been cultured through an adaptation of an old 
fisherman's trick. Worms collected in the fall were placed in a mixture 
of black soil and leaf mold in a covered, galvanized iron can, such as a 
garbage can. Through the winter, the contents of the can were kept 



Gephyrea 197 

moist but not wet, and additions of spent coffee grounds were made at 
intervals of a month or so. Excessive moisture was corrected by leaving 
the cover slightly raised. The culture has lived through the three 
summer months without any attention, but was rather weak in the fall.* 



Class Gephyrea 



CULTURING LARVAE OF URECHIS CAUPO 

G. E. MacGinitie, California Institute of Technology 

THE larvae of the echiuroid, Urechis caupo, may be successfully 
reared by the method given below. This method has also been used 
to rear the larvae of the phoronid, Phoronopsis viridls, the polychaete, 
Halosydna brevisetosa, the tectibranch mollusk, Tethys calijornicus, and 
sand dollars and sea urchins. No doubt it will serve as a method for 
rearing many others as yet untried. The following materials will be 
required: Syracuse watch glasses; finger bowls with glass covers; sea- 
water, preferably filtered not long before use; pipettes or medicine 
droppers with the small end pulled out to % or % of its original opening; 
at least two pipettes more finely drawn out with which to tap the animals 
for eggs and sperm, and a diatom culture. 

COLLECTING THE ANIMALS 

Urechis caupo inhabits the salt water estuaries of the west coast from 
San Diego northward. They build elongated U-shaped burrows with 
two openings to the surface of the mud. These openings are on an 
average about 30 inches apart, and the burrow itself is from 12 to 16 
inches deep. Although the mud flats of estuaries are perforated with the 
openings of burrows of a great variety of animals, each and every opening 
in some way gives a clue to the underground inhabitant that made it. 

♦Editor's Note: Mr. Ralph W. Moltke of 106 Broadway, Peoria, Illinois, issues a small 
bulletin on permanent-bed culture of Allolobophora [—Eisenia] foetida that he sells to 
fishermen and to dealers in baits for fishing. It contains detailed practical instructions. 
With his permission we summarize here the main features of his plan. 

1. Select a shady place in well-drained soil for a bed, say 3x6 feet. 

2. Dig out a foot or more of soil and fill with well rotted manure. 

3. Wet it down thoroughly and introduce the worms by distributing them over the 
surface. 

4. Cover them with a sprinkling of dirt and allow them time to burrow. 

5. Add some pieces of hard stale bread soaked in water and cover these with dirt. 

6. Feed the worms once a week by spreading over the surface of the dirt corn meal, 
old bread, or vegetable refuse from the kitchen, covering this with dirt each time and adding 
a layer of straw or gunny-sacking to retain the moisture. Sprinkle the bed with water 
whenever it shows signs of getting dry. 

7. Remove worms for use by turning over surface layer with a hand digging-fork, 
digging in a new place each time. 

J. G. N. 



1 9 8 Phylum Annelida 

The burrows of Urechis may be distinguished in the following manner: 
The opening of the burrow varies somewhat in size according to the 
size of the animal within, but is always smaller than the deeper portions. 
The diameter of the opening varies from the size of a lead pencil to 
some twice that size, and is always well smoothed and has a somewhat 
slick appearance due to the mucus secreted by the animal, and which 
holds the sand and mud in place. At one or the other of the openings 
castings usually will be found. These castings are of equal length and 
smoothly rounded at each end. A casting of average size will be about 
i mm. in diameter and i cm. in length. 

Since Urechis is one of the few animals which has two openings to its 
burrow, one may feel quite sure that it is a burrow of this animal, if, 
after locating two openings about 30 inches apart, one steps quickly on 
one hole, or jabs a shovel handle into it, and the water squirts out the 
other hole. This also locates the animal underground in so far as it must 
be somewhere between the two openings. This test may not be repeated, 
for, after being disturbed, Urechis will tightly block the burrow. 

In bringing living specimens of Urechis to the laboratory it is necessary 
that they be kept cool to prevent harm both to the animals themselves 
and to the sex products that they contain. If they are to be transported 
for any distance the container in which they are carried should be packed 
in ice. By using ice I have transported them a distance of 250 miles 
without ill results. Males and females should be segregated by testing, 
and kept in separate aquaria so that when used in the future the par- 
ticular sex needed may be known readily. 

MAINTENANCE IN THE LABORATORY 

If the animals are to be used for only a short period of time they 
may be left in the bottom of an aquarium with a good supply of running 
salt water. However, if they are to be kept for longer periods of time it 
is much better to keep them in glass tubes. These tubes may be bent 
U-shaped to simulate their natural burrows, in which case the animals 
may be kept for years. But even straight tubes in which they may be 
confined by corking the ends with single-holed corks will keep the ani- 
mals in good shape for a much longer period than usually will be the 
case when they are kept free in the aquarium. In the U-shaped tube 
they will feed and carry on all the activities of their natural habitat. In 
general they will not feed in a straight tube, although an occasional 
individual may do so. The glass tubing in which they are confined 
should be of ample dimensions, as when one is handling the worms they 
lose their respiratory water and may be confined in a space much smaller 
than they actually require. Especially in the southern limits of their 
range Urechis is often spawned out in summer, so that in order to insure 



Gephyrea 199 

material to cover this period it is necessary to collect the animals in May 
and keep them for rather long periods of time in the manner just de- 
scribed. 

FERTILIZING THE EGGS 

Eggs or sperm may be removed from any of the six gonopores, which 
lie in three pairs just posterior to the oral setae, by inserting into one of 
these pores a finely drawn out pipette, which, of course, has been fire- 
ended. Use one pipette for males and another for females to avoid 
premature fertilization. Four or five hundred eggs should be placed in 
a finger bowl and covered with % inch of filtered seawater, and then 
fertilized by the introduction of sperm, at the same time mixing well by 
squirting water in and out of the pipette. A minimum amount of sperm 
for insuring fertilization should be used, as polyspermic conditions and 
abnormal larvae nearly always result from an excess of sperm. 

CARE OF THE LARVAE 

These fertilized eggs should be covered and left to develop to the 
trochophore stage, or from 18 to 24 hours. Then it is advisable to take 
from these larvae those which are actively swimming near the surface 
and distribute them to several finger bowls with approximately 50 to 
each bowl. Remove any abnormal larvae whenever any are observed. 
Put glass covers on the finger bowls and set them in a cool place. From 
this time on the larvae should be carefully inspected and fed each day, 
and changed to fresh filtered seawater about once each week. 

The above directions may need to be modified somewhat for the larvae 
of other animals. 

It should be remembered that the length of the larval stage before 
metamorphosis is not a criterion of what it would be under natural con- 
ditions for obvious reasons. Since feeding must be carefully done to 
prevent fouling, it is probable that the amount of food is quite different 
from that of the open ocean. When raised in the above manner Urechis 
larvae usually require from 40 to 60 days before they metamorphose 
into burrowing worms. 

DIATOM CULTURES 

We have used several diatom cultures in the feeding of marine larvae, 
but we are unable to give a technical name for any of the species used. 
The first culture I used was one which I believe originally came from the 
Plymouth Marine Laboratory, and which was already at the Hopkins 
Marine Station of Stanford University when I first went there. Later I 
cultured a single-celled green alga which grows quite abundantly on the 
wet sand of the beaches. Later still a splendid form was cultured from 



200 Phylum Annelida 

the diatoms of the ocean here at our laboratory. In addition to this 
another culture was sent to me by Dr. S. C. Brooks a year ago from 
Woods Hole. Of these cultures the most suitable for food for growing 
larvae are the first and third, as these are small in size and stay in sus- 
pension. 

FEEDING 

Because it is impracticable to know the concentration of the diatoms 
within any culture, and because of the varying ability of the animals 
to use certain amounts of the diatoms as they grow, and also because 
the number of larvae in the dishes varies somewhat, it is impossible to 
state definitely what amount of diatom culture should be given to the 
feeding larvae each day. 

Only that amount of diatoms should be fed to the larvae each day that 
they will clean up well by feeding time on the following day. Due to the 
transparency of the trochophores it is a simple matter to ascertain if they 
are feeding well, for the ingested diatoms may be seen massed within the 
gut. 

Be sure, however, that the amount of material is sufficient to last 
until the time of the next feeding. An average feeding from an average 
diatom culture at the beginning of the feeding stage would be about % cc - 
to each finger bowl containing 50 larvae. It is better to feed less than 
may actually be used in the beginning, and increase this amount until 
the correct amount is known. 

SUMMARIZED DIRECTIONS 

A number of important factors to be considered are given below, and if 
these are carefully observed I think successful results would be quite 
certain with any free-swimming marine larva that is fairly hardy. 

1. The seawater should be carefully filtered to remove all plank tonic 
organisms, and unless one is quite certain of the non-toxic condition 
of the salt water pipe installation of the laboratory, the seawater should 
be carried in from the outside in a clean glass container. 

2. The temperature should be kept as low as possible, preferably at 
or below that of the outside ocean water. Guard against any sudden 
change of temperature. 

3. Not more than 50 to 75 larvae should be kept in a finger bowl at 
the beginning, and this number should be reduced to 25 or 30 after 
feeding has become well established and growth has commenced. 

4. Do not put more than three-fourths inch of water in each finger 
bowl, as more increases the ratio of volume to surface and hence allows 
less oxygenation. 

5. A good diatom culture for feeding is necessary. The diatom should 



Hirudinea 201 

be quite small and should be one which will stay in suspension. 

6. Each day, and this is most important, the bowls should be care- 
fully inspected to note: 

a. Are the larvae quite active? 

b. Are they feeding? 

c. Is the diatom culture well under control by the feeding larvae? 
Perhaps nothing will insure failure more surely than feeding too much 
diatom culture, or allowing the culture to reproduce in the finger bowls. 
A sufficient amount of it should be fed once each day so that upon in- 
spection the following day at feeding time practically all has been used 
by the larvae as food. 

7. If there are signs of any fouling, or if the diatoms become too 
numerous, the larvae should immediately be taken out with a small 
pipette and transferred to a clean bowl with freshly filtered seawater. 

8. Each day, just after feeding, gently aerate the water in each bowl 
with three or four pipettefuls of water from the same bowl. Better still, 
just before feeding carefully remove three or four pipettefuls of water 
from each bowl, feed the larvae, and then squirt three or four pipettefuls 
of freshly filtered seawater into each bowl. Either of these methods 
serves to aerate the water and to distribute the diatoms throughout the 
water in the bowls. 

Bibliography 

Fisher, W. K., and MacGinitie, G. E. 1928. A New Echiuroid Worm from Cali- 
fornia. Ann. and Mag. Nat. Hist. 1:199. 
1928. Natural History of an Echiuroid Worm. Ibid. 1:204. 

Class Hirndi?iea 

LABORATORY CARE OF LEECHES 

J. Percy Moore, University of Pennsylvania 

BESIDES serving for problems peculiarly their own, leeches offer 
suitable subjects for the study of certain general problems of 
physiology and behavior and some of them provide beautiful material 
for embryology, both observational and experimental. For the latter 
purposes common species of the Glossiphonidae, such as Glossiphonia 
complanata, Helobdella stagnalis, and Placobdella parasitica, are to be 
recommended, as the eggs and embryos are carried in large numbers by 
the female parent either in delicate capsules or uninclosed. 

Except for certain of the fish leeches (Ichthyobdellidae) culture is 
simple and easy. Many of them will live under almost anerobic con- 
ditions and the sanguivorous species especially will thrive for a long time 
on a single meal of blood, or even without feeding at all. I have kept 



202 Phylum Annelida 

Macrobdella decora in perfectly good condition without food for as long 
as 14 months. Most of them require little space and they are resistant 
to a wide range of temperatures, especially at the lower ordinary registers. 
But cleanliness is requisite as most species soon succumb to foul or over- 
heated water. They are also extremely sensitive to many mineral and 
organic poisons and even minute traces of copper sulphate, calcium 
chloride, chloroform, nicotine, etc., may be quickly fatal if the leeches 
cannot escape. 

The fish leeches (Ichthyobdellidae) are mostly marine and have been 
little cultured. Some of them have been kept in the aquaria at Naples 
and Plymouth where their natural hosts are available. The freshwater 
Piscicola and related genera may be kept in aquaria, either balanced or 
with running water, with sunfish, goldfish, or other small fishes as hosts. 
They will attach their egg capsules to aquatic plants, to stones, or to the 
glass. They are sometimes exceedingly numerous and harmful in the 
artificial ponds and tanks of trout and other fish hatcheries. 

But most useful are the common species of Glossiphonidae. The 
smaller species may be kept indefinitely, preferably in a moderately cool 
and light place out of direct sunlight, in finger bowls or small crystallizing 
dishes, with a few sprigs of Elodea or similar aquatic plants. A few 
living water snails, such as Physa or Lymnaea, should be added from 
time to time for food. Pond or spring water should be used, as tap 
water is frequently chlorinated or otherwise treated and as a consequence 
is likely to be injurious. Any dead or dying leeches or snails and their 
feces should be removed promptly. The water should be changed if it 
shows any indications of contamination. This is easily done as the leeches 
usually cling firmly to the sides of the vessel. If, in order to stimulate 
the growth of the plants, it is desired to place the dishes at a window ad- 
mitting some sunlight, a few pieces of shale, clam shell, or dead leaves 
should be added to afford concealment and protection. Twenty or thirty 
of such small species as Helobdella stagnalis or H. lineata (jusca) or half 
as many Glossiphonia complanata will thrive in a finger bowl, exchange 
spermatophores and produce fertile eggs in abundance throughout spring 
and summer. The young are easily raised but care should be taken to 
avoid overcrowding or undue disturbance as they often die if detached 
from the mother before most of the yolk is absorbed. After the young 
become free it is best to remove them to a separate dish. Should it be 
desired to keep large numbers of these leeches, small balanced aquaria 
with a bottom layer of sandy soil and with plants with ensheathing leaf 
stalks, like Sagittaria, in addition to Elodea or Myriophyllum, and a 
supply of snails will serve admirably. 

The larger glossiphonids (Placobdella parasitica, P. rugosa, P. multi- 
lineata, P. montijera, etc.) may be kept indefinitely in finger bowls or 



Hirudinea 203 

other flat glass or clay dishes but only 2 or 3 to 5 or 6, according to size, 
should be placed in a vessel. Most of these normally feed on the blood 
of snapping or other water turtles (the last named species on frogs and 
toads) and a supply of these animals should be available. A meal at 
intervals of a month or two is sufficient, and it is better to prevent the 
leeches from becoming too heavily gorged. A meal of blood in early 
spring is usually followed closely by egg-laying, and often reproduction 
may be initiated during the winter by placing the leeches in a moderately 
warm room and permitting them to feed on a turtle or frog. 

The Erpobdellidae (various species of Erpobdella, Dina, and Nephelop- 
sis) are equally easy to keep if certain precautions are taken. As they 
are much more active than most of the Glossiphonidae, they require more 
spacious quarters and the vessels should be securely covered to prevent 
them from escaping, particularly at night. These leeches are largely 
nocturnal, predacious and incline to be amphibious. Consequently they 
often leave the water at night in search of earthworms and similar food. 
They are also scavengers and will feed on dead or wounded fish, frogs, 
etc. In confinement they are best fed with small earthworms, the larger 
aquatic Oligochaeta, insect larvae, or finely chopped fresh meat. Plants 
are usually unnecessary for leeches of this group, but the water should be 
kept clean, especially after feeding. Plenty of small pieces of stone, bits 
of bark, and dead leaves should be provided as places of concealment and 
for the attachment of egg capsules, which are flat, purse-shaped struc- 
tures, each containing several eggs attached by a flat side to any firm 
substratum. Egg capsules are produced in great numbers during the 
spring and summer. 

The true blood sucking and medicinal leeches and the related so-called 
horse leeches, belong to the family Hirudidae. These include the largest 
of our freshwater leeches. During the middle decades of the last century 
when leeches were employed medicinally in great numbers they were 
extensively cultivated in so-called leech farms, especially in France.* 

On a small scale these leeches may be raised in tanks or aquaria or 
even in earthenware jars. The Indian medicinal leech (Hirudinaria) is 
cultured largely in this way, the reproducing leeches being placed in a 
jar containing some wet clay, the egg capsules removed daily and placed 
in moist clay cups until the young hatch when they are transferred to 
water and after a time cautiously fed. Our American leeches of this type 
(Macrobdella, Philobdella) are best kept in low-sided aquaria or tanks 
with a sloping bank of sandy earth at one end and shallow water at the 
other. A cover of thick moss on the earth is desirable, as well as some 

* The technique of leech culture was elaborately described in many books published 
mostly in France, of which Ebrard — Novelle Monographic des Sangsues Medicinales, 1857, 
is a good example. 



2 04 Phylum A nnelida 

stones or pieces of wood under which the leeches may hide. No aquatic 
plants are required as these leeches spend much of their time hanging 
from the sides of the vessel above the water and exposed to the air. A 
small number may be kept alive indefinitely in a glass or crockery jar 
with a small amount of water and some Sphagnum moss. The leeches 
of these genera and especially Philobdella, are less strictly sanguivorous 
than Hirudo, etc., and add to their normal diet frogs' eggs, aquatic larvae, 
Oligochaetes, etc. A meal of blood about every six months is sufficient. 
This may be taken from frogs or small fishes, but mammalian blood is 
better as being greater in amount and percentage of solid matter. Care 
should be taken to avoid overfeeding, which checks breeding. The egg 
capsules are deposited in the earth just above the water level and are 
best slit open with fine scissors to secure the eggs. Our largest leeches 
belonging to the genus Haemopis are more strictly predacious and often 
wander at night a considerable distance from the water in search of food. 
One subspecies has become practically terrestrial, living in garden soil 
and feeding upon earthworms which are the best food for all species in 
confinement, although they also eat insect larvae, smaller leeches, snails, 
and almost any animals of suitable size. Except for feeding, culture 
methods are similar to those recommended for Macrobdella. All leeches 
of this family are given to wandering, and the vessels should be securely 
covered, preferably with fine fly screen. 



Phylum XIV 

Arthropod a, Class Crustacea 
Subclass Entomostraca, Order branchiopoda 



A METHOD FOR REARING ARTEMIA SALINA 

R. M. Bond, Santa Barbara School, Carphiteria, California 
Artemta salina is an anostracan phyllopod crustacean about 12 mm. in 
length. This genus with practically world-wide distribution may be di- 
vided into several forms, some of which are parthenogenetic. The tax- 
onomy of the genus is at present in a confused state. The form found in 
North America is sexual, and occurs naturally in Epsom Lake, Washing- 
ton; in certain natural salterns along the California coast from San 
Francisco southward; in Mono Lake, California; in Little Soda Lake, 
San Luis Obispo County, California; in Great Salt Lake; and probably 
elsewhere. It has also appeared in numerous man-made salterns, espe- 
cially where salt is extracted from seawater by solar evaporation. 

Resting eggs float in brine and do not hatch until after drying. They 
are carried by the wind to the lee side of the saltern and are there 
piled (mixed with debris) in windrows. They may be collected with a 
shovel and buckets. Artemia eggs mixed with salt, etc., may be obtained 
from dealers in tropical fish, from the Leslie Salt Company, Redwood 
City, California, and from San Francisco Aquarium Society, at a price of 
about $0.50 an ounce. The dried eggs remain viable for several years. 

For physiological experiments, or for other purposes, it is often desir- 
able to separate the eggs as completely as possible from foreign matter. 
This may be conveniently done as follows: Dry the eggs in air and sift 
through a 10-mesh sieve; suspend in 10-20 volumes of 15% NaCl in a 
large separatory funnel, and shake well. Heavy substances will settle and 
salts will dissolve. The brine should be changed about twice a day till 
it remains clear (about 6 changes, and the final washing should be 
drained off as completely as possible. The eggs should then be washed 
in the funnel with distilled water, caught on a 100-mesh sieve, allowed 
to drain for an hour, spread out, dried in a current of air at 25-30 C. till 
thoroughly dry (about 30 hours), and then put through a 50-mesh 
sieve, through which they will just pass. In the distilled water some eggs 

205 



206 Phylum Arthropoda 

will sink and some will float. The former are nearly 100% viable. A 
majority of the floaters will also hatch, but the percentage of viability will 

be smaller. 

For transportation, young Artemia must be placed in open contain- 
ers, since they are very susceptible to accumulated C0 2 . 

The hatching medium may be almost any salt solution not containing 
much potassium, and ranging in concentration from about 0.1% to 6%. 
Natural or artificial seawater is as good as anything. After hatching, 
the nauplii may be transferred directly into more concentrated solutions, 
but in the higher concentrations the mortality may be great unless the 
change is made gradually, either by stages or by evaporation. Older 
animals are much less resistant than nauplii, and are killed by large 
changes in concentration of the medium, unless the changes are very 
gradual. When a culture is established it is continued by viviparous 
reproduction. The first batch of eggs produced by the females usually go 
at once to the nauplius stage in the brood pouch and then escape. Subse- 
quent batches of eggs from the same females are resting eggs and must 
be dried before hatching. Under most favorable conditions, a generation 
takes about 3 weeks. 

It is possible to raise the animals in a wide variety of salt mixtures, from 
about 4% to concentration, always provided that the concentration of 
potassium is not too high in proportion to other salts present. For getting 
the animals to reproduce as rapidly and as vigorously as possible, I 
have found nothing better than seawater with 5 to 8 gm. NaCl per 100 cc. 
added. Artificial aeration of cultures in unfavorable media is helpful. 

For food, particulate matter is required. Ordinary yeast is excellent 
and convenient. It should be suspended in enough freshwater to make 
up for evaporation and floated on the salt medium. For starting cultures, 
however, and for unfavorable media, a rich culture of a one-celled green 
alga such as Dunaliella salina, D. viridis, or Platymonas subcordaeformis 
will be found very helpful. [For culture see p. 134.] Food should be 
added in small quantities every day or two. 

The optimum temperature is about 30 C, but Artemia will live at 
temperatures as low as io° and as high as 37 . 

Reference 
For the culture of Artemia see also p. 215. 



Sididae and Daphniidae 207 

Families sididae and daphniidae 

CULTURE OF CLADOCERA 

A. M. Banta, Brown University 

THESE small animals are useful as food for cultures of hydra, young 
and older aquarium fish, and larval salamanders. More directly as 
scientific material, they are very useful for laboratory teaching and 
for research purposes. 

They provide a favorable laboratory type for illustration of the struc- 
ture of an entomostracan for which, because of their transparency, they 
may readily be used alive to demonstrate most of the morphological 
structures and in addition several physiological activities (respiration 
feeding and egestion, circulation, reproduction). They are also ad- 
vantageous for the study of animal behavior, adjustment to environment 
adjustment to the annual seasonal cycle, etc. They are unexcelled for 
the study of parthenogenetic reproduction and the environmental control 
of sex of offspring. 

As material for physiological studies or experimentation a clone of 
cladocerans provides the almost unique advantage of genetic uniformity, 
which is practically guaranteed by their diploid parthenogenetic repro- 
duction. 

For studies in the genetics of a pure line or clone they are probably 
unequalled among metazoans. Also, thanks to the technique developed 
by Miss Thelma R. Wood, they may be used for studies of genetics in 
sexual reproduction, an essential supplement to the analysis of the genetics 

of the clone. 

Live animals with which to start cultures may ordinarily be obtained 
from small ponds or lakes during the open season and frequently through 
the ice in winter. Daphnia longispina and the species of Simocephalus 
may be secured in moderate numbers in many sections the year round. 
Simocephalus also occurs occasionally in dense vegetation in relatively 
quiet portions of freshwater streams. Daphnia pulex is frequently 
abundant especially in spring and early summer in clear and often in 
rather dirty pond water. The species of Moina are found in late spring 
and summer in pig-lot or stable-yard puddles and in other situations 
where the content of organic matter in the water is high. 

The essential food of most Cladocera is bacteria or single-celled, not 
filamentous, algae. For Daphnia magna, algae are sometimes more 
advantageous than bacteria although usually D. magna does very well in 
manure culture medium. But for the other species of Daphnia, for 
Simocephalus, and some of the Sididae, bacteria seem equally good or 
better. For Moina, bacteria of the colon group (which generally prevail 



208 Phylum Arthropoda 

in manure solutions) seem to be best; and live bacteria appear to be 
essential (Stuart, McPherson, and Cooper, 1931). 

The manure solution or "stable tea" to be described below has, in the 
writer's experience, proven a most satisfactory culture medium. It would 
be misleading, however, to encourage the worker to think that this cul- 
ture medium is infallible and that every make-up of medium is equally 
good. It is, however, a highly successful method. 

The manure solution or stable tea medium may be made up in accord- 
ance with either of the following formulae in battery jars 9 inches in 
diameter by 12 inches high or in stone jars or enameled containers 
approximately that size. Galvanized or copper containers are to be 
avoided for handling the pond water inasmuch as zinc and copper are 
extremely toxic to Cladocera. 

Formula I. Garden soil 2 lb. 

Horse manure 6 oz. 
Pond water 2^2 gal. 

The ingredients should be placed in the container in the order named. 
The container may be kept in a cool place or surrounded by running 
water to keep the temperature 15 to 18 C. After 60 to 72 hours (or 
48 hours if the temperature has been higher) the floating manure, if any, 
is removed and the super-natant liquid strained through a silk bolting 
cloth (about 130 meshes per inch) or other similarly porous, smooth- 
threaded cloth. With the final liter or so of the liquid a quantity of 
silt is placed within the straining cloth and enough silt is worked through 
by active stirring or gently rubbing through the cloth to produce in 
settling a layer of sediment 1 or 2 mm. thick on the bottom of the jar. 
This strained liquid constitutes the stock medium. It requires dilution 
with pond water before being used as culture medium. The proper 
dilution may vary between 1 part of the stock medium to 2 to 4 parts 
of pond water, depending upon the density in appearance of the liquid. 
The pond water used both in making up the stock medium and in the 
dilution of it is strained or filtered to avoid contamination with "wild" 
Cladocera and copepods.* 

The horse manure may be obtained from a stable and allowed to age 
for a week or ten days before use. Fresh manure, dry or moldy manure, 
or manure more than a month old is ordinarily to be avoided. It is con- 
venient to keep a small supply covered over in a wooden box or cardboard 
carton, which may be kept outdoors (but sheltered from rain) or in a 
basement location. The stock medium in process of ripening and the 

*Tap water has frequently been used after it has stood in an aquarium containing some 
fine sand or silt for at least a week or ten days. But it is questionable if tap water which 
has been heavily chlorinated or subjected to other extreme measures to render it potable 
may be so readily "conditioned" and thus rendered suitable for such use. 



Sididae and Daphniidae 209 

culture medium in use are practically odorless and may be kept in the 
laboratory. 

The soil to be used handles better if it is of somewhat sandy nature 
and of not too fine a texture. The main difficulty with soil of very 
fine particles is the slowness with which the silt becomes settled and thus 
leaves the medium transparent — but slightly reddish brown in color. 

The culture medium must be well stirred while being dipped out into 
the culture containers (which are not covered or stoppered) in order that 
some of the silt may be in each individual culture. Immediately after 
straining or within two days thereafter the stock medium is at its best. 
It is ordinarily nearly spent after 5 to 7 days. It has been found that the 
numbers of bacteria rapidly decrease after about the third day following 
straining. Presumably the accumulation of by-products of bacterial 
growth and decrease in numbers of bacteria both contribute to render the 
older solution less effective as a culture medium. 

The second formula for making up the manure solution medium pro- 
duces a much more concentrated stock medium. 

Formula II. Garden soil 2 lb. 

Horse manure 12 oz. 
Pond water 5 qt. 

The handling of this make-up differs from that of Formula I only in that 
the dilution of the resulting stock medium in preparation for use as cul- 
ture medium is much greater, ranging from 1 part of the strained 
medium with 4 to 10 parts of pond water. The writer prefers to use the 
first formula. Several workers prefer and have excellent results with the 
second formula. 

The stable tea has been used and found effective by the writer in open 
cultures (not covered or stoppered) in quantities from 25 cc. to 10 liters. 
We have made few attempts at rearing mass-cultures of Cladocera. 
Manure solution culture medium may readily be used for mass-cultures 
if every few days it is renewed or strengthened by the addition of small 
quantities of manure (preferably tied up in cloth bags and submerged). 
Others have with good results occasionally thrown a dead guinea pig or 
other small animal into a tank in which an abundance of a culture of 
Cladocera is on the decline. A more precise method for such situations 
as large laboratory tanks or outdoor tanks is the use of manure or the 
employment of Dr. Embody 's soy bean meal [see p. 218] or W. A. 
Chipman's (1934) cotton seed meal [see p. 212] culture methods. In 
all of these methods the essential feature is the provision of decaying 
organic matter upon which the proper bacteria may develop and con- 
tinue to be present in amply large numbers. 

The beginner with the manure solution medium might best first em- 



210 Phylum Art hr op o da 

ploy approximately the minimum dilutions suggested above and grad- 
ually increase the strength of the medium. Medium that is unnecessarily 
weak will give rise to small clutches of young — fewer than 8 per first 
clutch for Daphnia or Simocephalus or fewer than 12 for Moina. First 
clutches of 10 to 18 Daphnia or Simocephalus or of 15 to 25 for Moina 
are to be expected under approximately optimal cultural conditions. 
Over-dilution is also indicated by too little apparent density of the 
medium. In 200 cc. large-mouthed, tall salts bottles it should be 
moderately opaque grayish brown when first placed in the bottles and 
be fairly clear of suspended matter and of a light amber color after two 
days in the bottles. Over-strong medium is indicated if it appears too 
dense or (other conditions being normal) if a considerable percentage of 
newly transferred animals die, if the development is retarded and clutches 
of young are small and are produced irregularly, if embryos die in the 
mother's brood chamber, or (Moina) if first clutches are large but many 
of the mothers die after their young are released. A little experience will 
dictate the amount of dilution to be employed. Two successive make-ups 
may require somewhat different dilutions. The worker must decide by 
inspection when the dilution is sufficient. 

In the writer's laboratory this medium has proven eminently satis- 
factory when made up with pond water and with horse manure in the 
proper condition; when the medium has been used with some of the silt; 
and when it is properly diluted, over-strong medium being especially 
avoided. This medium has been used by us primarily for rearing to ma- 
turity or longer, without change of medium, cladocerans in 200 cc. bottles 
(half filled) in pedigree cultures of several species and many different 
clones of Cladocera. Excellent reproducing specimens for laboratory 
use may be reared from young in 6 to 8 days with 2 to 4 per bottle. 

Dr. D. D. Whitney has used a similar stable tea medium for rearing 
Hydatina asplanchna and other rotifers. In all probability this medium 
may prove useful for other aquatics {e.g., mosquito larvae, etc.) which 
may be fed primarily upon bacteria and which flourish in natural waters 
containing considerable organic matter. 

Bibliography 
Chipman, W. A., Jr. 1934. A new culture method for Cladocerans. Science 

79:59- 
Stuart, C. A., McPherson, Maurita, and Cooper, H. J. 193 1. Studies on bac- 
teriologically sterile Moina macrocopa and their food requirements. Physiol. 
Zool. 4:87. 



Sididae and Daphniidae 211 

A NOTE ON BANTA'S CULTURE MEDIUM 

George G. Snider, University of Cincinnati 

AM. BANTA (192 1 )* introduced a culture medium for clado- 
♦ cerans which has been, and still is, extensively used. 
The following modifications of Banta's medium have been used by the 
writer and found to yield even better results than those obtained from the 
original directions. First, the manure is collected in a relatively fresh 
state and permitted to dry thoroughly. It is then added (8 ozs.) to the 
garden soil in finely divided form as described. Second, after the animals 
have been in the diluted culture medium (each animal in 100 cc. solution) 
4 days, 12-15 cc - strained undiluted medium are added. In the case of 
Daphnia magna this culture medium usually results in animals produc- 
ing from 15-20 or more young in their first broods. 

CLADOCERA CULTURE 

Harold Heath, Hopkins Marine Station 

THE equipment used in the culturing of Cladocera comprises three 
aquaria each with a capacity of 16 gallons. After two of these have 
been filled with water a thin layer of sand is spread over the bottom, 
and a few aquatic plants are anchored under small stones. At this stage, 
a cloth sack, containing approximately 8 ounces of sheep manure, is 
suspended in each aquarium, and the culture is allowed to stand for 
3 or 4 weeks. Some investigators, I understand, add lettuce leaves from 
time to time, but so far as my experience goes the ordinary decomposition 
of the aquatic plants affords, with the fertilizer, a sufficient pabulum for 
the bacteria and other unicellular organisms which soon appear. Into 
this mixture a stock of Cladocera is now introduced, and where the 
manure is renewed each month or so the culture usually flourishes for 
months, in several instances for more than a year. 

To safeguard against accidents or an unaccountable disappearance of 
the crustaceans it has been our custom to keep in reserve a third 
aquarium cultured according to the foregoing method but without Clado- 
cera. Also in our series approximately % of each aquarium was 
shaded — though this may not be necessary — and the tank was kept in 
sunlight where the diurnal temperature ranged from 54° to 74 F. 
during the year. 

Furthermore, in this region (Monterey Co., Calif.) it is necessary 
to place a screen over the aquaria to prevent the entry of two types of 
insects, mosquitoes and back-swimmers (Xotonecta sp.). The first 
named organisms probably do not interfere with the Crustacea, although 
it is reasonable to presume that they do diminish the food supply. The 

*A convenient culture medium for Daphnids. Science 53:557. 1921- 



212 Phylum Arthropoda 

back-swimmers, on the other hand, destroyed several colonies before the 
trouble was discovered. 

In conclusion it may be added that no particular attention has been 
paid to the pH of the water, nor to the contained salts. In some in- 
stances the water was drawn from the municipal mains which in turn 
are supplied from numerous mountain streams. At other times pond 
water was employed. So far as could be detected no differences existed 
in the growth rate of these two types of cultures. 

Such, in brief, has been my experience in culturing Cladocera, for 
which credit is due to several investigators in other parts of the country, 
whose verbal or written statements have been followed in large measure. 

A NEW CULTURE MEDIUM FOR CLADOCERANS* 

IN recent investigations in this laboratory it has been necessary to use 
numbers of cladocerans. In order to raise these animals in quantities 
and under controlled conditions various culture media have been re- 
viewed and tested. Most of the existing media call for manure to supply 
the organic matter, but as manure is so variable the substitution of 
materials of more constant composition was tried. Wiebe (1930) has 
pointed out that soy bean meal is superior to manure for plankton pro- 
duction in pond fertilization, and more recently the U. S. Bureau of 
Fisheries has found cotton seed meal quite, if not more, desirable for 
this purpose. It seemed logical, therefore, to substitute cotton seed meal 
for manure in cladoceran culture media. This change produced a very 
satisfactory culture medium, having several advantages over the manure 
infusions as suggested by Banta (1921). [See p. 208.] 

Pond water was filtered through coarse filter paper and added to a 
mixture of fine garden soil and cotton seed meal (commercial cotton 
seed meal, as used in dairy feeds), in the proportions of 1 liter of filtered 
water to 90 grams of garden soil and 17 grams of cotton seed meal. 
After a thorough stirring, the mixture was set aside at room temperature 
in large Erlenmeyer flasks for five days. During this period the mixture 
fermented and produced considerable gas. At the end of five days the 
supernatant fluid was decanted and then strained through muslin. 
Analyses showed that the strained fluid contained an almost pure culture 
of B. coli. The strained fluid was diluted with filtered pond water before 
using and re-strained through muslin whenever bacterial masses de- 
veloped. The pH of the final diluted product was adjusted to 7.2 by the 
addition of sodium carbonate. 

In strong concentration of this medium bacterial masses formed which 

♦Reprinted, with slight changes, from an article in Science 79:59, 1934, by Walter 
A. Chipman, Jr., U. S. Bureau oj Fisheries, 



Sididae and Daphniidae 213 

interfered with the free movement of the Daphnia and often resulted in 
their death, but by a dilution of 1 part of the strained fluid as decanted 
from the original mixture with 100 parts of filtered pond water a medium 
was obtained which remained clear and in which Daphnia grew rapidly 
and produced normal clones. It has been found desirable to renew the 
media in which the cultures of animals are growing from time to time, 
i.e., at periods of a week or more, but the addition of more bacteria to 
the cotton seed medium, as suggested for manure infusions by Stuart 
and Banta, has not been found necessary. Fresh stock supplies of the 
cotton seed mixture have been prepared each week, a small amount of an 
old mixture being added each time to insure inoculation with the original 
bacteria. 

References 
For the culture of Cladocera see also p. 136. 

Bibliography 

Banta, A. M. 1921. Science 53:557. 

Stuart, C. A., and Banta, A. M. 1931. Physiol. Zool. 4:72. 

Wiebe, A. H. 1930. Bull. U. S. Bur. Fish. 46:137. 

M. E. D. 

A CULTURE MEDIUM FOR DAPHNIA* 

FLEISCHMANN'S yeast has been fed to a mass culture of Daphnia 
magna with striking results, reproduction and growth being markedly 
more rapid, and population more dense than with any of the usual media. 
About Y 4 of a fresh yeast cake is mixed into a uniform suspension with 
from 50 to 100 cc. of water, and poured into the aquarium, which con- 
tains from 60 to 70 liters of water. The feeding is repeated every 5 or 
6 days. It is necessary to have a stream of air bubbling through the 
medium at all times, or the yeast may prove lethal, probably by giving 

off C0 2 . 

The method has not been tried on other species of Cladocera, except 
Moina affinis, with which it was equally successful, nor has it been tried 
with few animals in small containers, but it is so successful in the mass 
culture that it seems wise to make the food material known. It should 
be particularly useful in physiological work, in which the usual manure 
infusion may be a source of large quantities of unknown solutes. It 
should also be valuable in raising Daphnia in large numbers as food for 
other organisms. 

M. E. D. 

♦Reprinted, with slight changes, from Science 79:60, 1934. by R. M. Bond, Santa 
Barbara School, Carpintcria, California. 



214 Phylum Arthropoda 

METHODS FOR CULTURING DAPHNIA 

Arthur D. Hasler, U. S. Bureau of Fisheries 

i. Algae Method, a. Run tap water into battery jars or butter tubs 
and allow to stand for 24 hrs. for the purpose of getting rid of air bubbles; 
otherwise the water-fleas adhere to these and are carried to the surface 
where they soon perish. 

b. Add sufficient algae to tinge the water slightly green and inoculate 
with Daphnia. 

c. The alga Coccomyxa simplex may be cultured in large quantities 
by a method developed by H. Schomer. When the culture is at its peak 
it is centrifuged with a Birge-Juday centrifuge and it is this centrifugate 
that is added to the Daphnia culture. The algae may be grown in 
battery jars. The Schomer medium consists of: 



KH2P04 


2.7 gm. 


per liter 


MgS0 4 


4.9 gm. 


per liter 


Ca(N0 3 ) 2 


4.7 gm. 


per liter 



This method surpasses any I have tried in bringing about maximum 
rate of reproduction. 

2. Yeast Method (Modified from Bond's), a. Same as (a) above. 

b. Make a thick suspension of moist Star or Fleischmann's yeast in a 
flask by shaking chunks vigorously until a suspension forms; then add 
the suspension to the culture until the water is slightly milky. When the 
water-fleas have cleared it, add the same amount again. Water should 
be completely changed once a month and the crop seined often to prevent 
crowding. Aeration is not necessary, but the maximum reproductive 
rate is reached with aeration. Stirring once a day is sufficient to keep 
the culture going satisfactorily. 

c. Augmenting the yeast diet with the algae as described above is 
recommended. If the cultures are in good light, algae generally grow 
on the sides. They may be scraped off the walls with a razor blade and 
then broken up by stirring. These serve well to augment the yeast diet. 
One may also add about a teaspoonful of dry sheep manure per 4 gal. 
every two weeks. The addition of these substances gives D. magna, for 
example, the natural red color, whereas if they are raised on yeast alone 
their color is almost white. 

3. Bant a' s Method. [See p. 208.] This method gives very satis- 
factory results. I raised Daphnia for 9 months in total darkness, using 
culture water made according to his directions. 

4. Sheep Manure method, a. Let water stand in suitable containers 
24 hours. 

b. To every gallon of water, add 1 teaspoonful of dry sheep manure, 



Sididae and Daphniidae 215 

0.5 gm. of acid phosphate and a liter of aqueous soil filtrate (water that 
has been allowed to filter through rich garden soil). 

c. Allow mixture to stand for a day and inoculate with Daphnia. 

5. Aquarium water method. Aquaria that have become very green 
with a phytoplankton growth furnish a very convenient culture medium 
for Daphnia. Just remove the fish or transfer the water to another con- 
tainer and inoculate with Daphnia. This method was recommended to a 
tropical fish fancier who was having trouble with "green aquaria." He 
removed the fish, then inoculated with Daphnia; after the Daphnia had 
multiplied sufficiently to "clear" the aquaria, the fish were put back and 
had a real feast! 

20 C. is a satisfactory temperature at which to keep these cultures. 

Artemia may also be raised by any of these culture methods. I have 
kept individual Artemias living for 4 months by using algae. The salt 
content must be regulated (2 teaspoonfuls to % pint of water). 

If it is desired to cease the Daphnia cultures during vacation periods, 
it is only necessary to chill or "crowd" the cultures and thus produce 
ephippial eggs. These may be collected and stored. To hatch them, 
place outside a window for two weeks of October weather so that they 
freeze and thaw several times, then place in water to hatch. They may 
also be artificially frozen in a refrigerator; 8 thaws and freezes give the 
maximum yield. 

Bibliography 

Banta, A. M. 1921. A convenient culture medium for daphnids. Science 53:557. 
Bauer, V. 1921. Wie ernahren sich die Wasserflohe. Allg. Fisck. Zeit. Jahrg. 

46:30. 
Geyer, Hans. 1909. Einige Bemerkungen u. die Zucht von Daphnia. Wochen- 

schrift Aquar.-Terrar. Kde. Jahrg. 26:32. 
Knorrich, Friedr. Wilh. 1901. Studien ueber die Ernaehrungsbedingungen ein- 

iger fuer die Fischproduction wichtiger Microorganismen des Siisswassers. Forsch. 

Ber. biol. Stat. Blon. 8:1. 

DAPHNIA CULTURE 

Alfred W. Schluchter, Dearborn, Michigan 

THE writer has used several methods which seem to work fairly well. 
The first is a method similar to that described by Mr. Walter Chip- 
man.* [See p. 212]. A wheat bran fermentation was used here, instead 
of cotton seed meal. This was made as follows: About 20 grams of wheat 
bran was added to 3 liters of water and allowed to ferment for about 
one week in a moderately warm place. If the Daphnia culture was to be 
carried out in the open where plenty of sunlight was present, this culture 
medium was diluted to about 1 to 100 parts of water and then % to % 

* Science 79:59. I934> 



216 Phylum Art hr op oda 

gram each of sodium chloride and calcium sulfate per liter of water were 
added. This solution was stocked with Daphnia. More infusion was 
added later as needed. 

Experiments have shown that sunlight is important in the propagation 
of Daphnia and many methods which are successful in strong sunlight 
will not work when sunlight is completely absent. 

The second method was found best and most desirable for indoor 
propagation since sunlight is not necessary. It depends essentially on a 
combination of bran infusion and liver. Although the latter may be used 
alone, experiments indicate that a combination of the two may be more 
desirable than either alone. 

Ten grams of wheat bran were added to 1500 cc. of water and fer- 
mented a week, as before. The supernatant fluid was poured off and dis- 
carded; only the bran residue was used. This was then added to a shal- 
low tank of about 100 liters' capacity, salt and calcium sulfate being 
added as before. The liver was heated as rapidly as possible until the 
proteins were coagulated. It was then cut in slices about 1 mm. thick 
and of these slices about 0.1 to 0.05 grams per liter of water were added 
to the tank containing the bran infusion. If too much bran infusion 
or liver was added, Daphnia came to the top because of lack of oxygen. 
It was then necessary to replace some of the old with fresh water. 

After the liver is heated it may be dried by keeping in a cool place and 
then used as required. It has been observed that such liver slices in con- 
tact with the water will tend to become moldy in the absence of a 
sufficient number of Daphnia, but if enough Daphnia are present the mold 
will disappear. Nearly all of this work was done with Daphnia magna, 
although the above methods were found to apply to several other Clado- 
cerans as well. 

PROPAGATING DAPHNIA AND OTHER FORAGE 
ORGANISMS INTENSIVELY IN SMALL PONDS* 

FOR the first few years we tried to keep cultures going continuously 
throughout the summer in the same pond, merely adding from time 
to time a definite amount of fertilizer. The result was that very success- 
ful cultures were maintained during May and part of June which always 
ran out in July but seemed to come back in late September and October. 
The most important causes for the diminishing supply seemed to be the 
population density, the accumulation of waste products not only from the 
Daphnia themselves but from micro-organisms also present, predatory 
enemies of Daphnia, and probably water temperatures somewhat above 

♦Abstracted from an article in Trans. Amer. Fish. Soc. 64:205, 1934, by G. C. Embody, 
Cornell University, and W. O. Sadler, Mississippi College. 



Sididae and Daphniidae 217 

82 ° F. All of these factors have been mentioned before by various 
investigators, especially by Dr. A. M. Banta and his associates. All of 
the above mentioned factors except predatory enemies favor the produc- 
tion of so-called winter eggs, which is usually an indication that the 
culture is on the wane. 

In order to have strong cultures at all seasons it becomes necessary to 
keep the individual Daphnia in the active condition of producing asexual 
or so-called summer eggs only, which demands the elimination of the 
inhibitive factors just mentioned. 

A too dense population, of course, is easily reduced by using the 
Daphnia. The excessive accumulation of waste matter is corrected by a 
change of water in the pond. Predatory enemies are controlled partly by 
sterilizing the pond bottom and sides immediately before starting a cul- 
ture and later by spraying the surface with some non-toxic animal oil, 
such as herring oil, salmon oil, or cod-liver oil. The water temperature 
cannot be controlled and, consequently, where it ranges above 82 ° F. for 
any length of time it may be difficult or impossible to produce cultures 
of Daphnia magna. 

A population density reaches the maximum under the experimental 
conditions maintained in our ponds in from 16 to 26 days with the water 
temperature varying between 70 and 8o° F. Consequently cultures are 
permitted to develop for 2 1 days, when they are fed to the fish and an 
entirely new culture with fresh water is started. The schedule of opera- 
tions is as follows: 

First day — Pond is drained, bottom and sides thoroughly disinfected 
with a strong solution of chlorinated lime, allowed to stand 6 hours; 
then refilled, fertilized, and stocked with large Daphnia from an active 
culture. 

Fifth to seventh day — Second fertilization. 

Tenth to fourteenth day — Third fertilization. 

Twenty-first day — Drawing the pond and using the Daphnia. 

In general this routine was continued through several summers from 
May to October and when a proper amount and kind of fertilizer was 
administered, the results were consistently good. 

Although we have tried to maintain pure cultures of Daphnia, other 
organisms have naturally appeared and multiplied. Some of these are 
desirable food animals but others are predators. 

Several little hard-shelled ostracods have appeared in considerable 
numbers, especially late in the culture period. We believe there is some 
connection between their abundance and a decline in the production of 
Daphnia, without as yet having direct evidence that they actually prey 
upon living Daphnia. Disinfection of the pond does not entirely elimi- 
nate them but nevertheless helps to keep them under control. 



218 Phylum Art hr op oda 

Midges of the genus Chironomus are attracted to the pond within a 
day or so after adding fertilizer, probably by the odors of fermentation, 
and deposit enormous numbers of eggs. These add considerably to the 
production of the pond. Mosquitoes likewise are attracted at about the 
same time and their characteristic floating egg masses become con- 
spicuous all over the surface. 

These three associated organisms may be considered beneficial insofar 
as they increase considerably the production of fish food. The mosqui- 
toes, however, are obnoxious as adults and since they transform to the 
adult stage long before the Daphnia culture reaches a peak, it is prob- 
ably better to exterminate them with non-toxic oil spray 4 to 6 days after 
the eggs appear. Other groups of animals which almost invariably appear 
are the back-swimmers, larvae of aquatic beetles, nymphs of dragonflies 
and damselflies, and hydra. All are predatory on Daphnia and midges. 
It is well known that the oil spray kills all insects that must come to the 
surface for air, such as the larvae of mosquitoes, beetles and adult back- 
swimmers. The dragonfly and damselfly nymphs and hydras are elimi- 
nated when the pond is drained and disinfected. 

Table I. 

Amount of Fertilizer per 100 Cubic Feet of Water and Its Apportionment 
(Experimental Ponds at Ithaca, N. Y.) 

Sheep Manure 
Soy Bean Cotton Seed Dry and 

Fertilizer Meal Meal Buttermilk Soy Bean Meal 



Initial Dose 


1 pt. 


1 qt. 


54 Pt- 


4 qts. Sh. M. 
1 qt. B. M. 


2nd Dose 


5th day — 


7th day — 


7th day — 


14th day — 




1 pt. 


1.5 pt. 


54 pt. 


y 2 pt. b. m. 


3rd Dose 


10th day — 


14th day — - 


14th day — 






1 pt. 


1.5 pt. 


A Pt. 




4th Dose 


15th day — 

l A Pt" 









Many different fertilizers have been tried, including manure from 
horses and cattle, dried sheep manure, acid phosphate, soy bean meal, 
cotton seed meal, dry buttermilk, and alfalfa meal. The alfalfa meal pro- 
duced only fair cultures and required such a large quantity of material 
that experiments were early discontinued. The dried sheep manure used 
in combination with either acid phosphate or soy bean meal produced 
average cultures consistently. The wet animal manures were from ordi- 
nary barnyard piles containing much straw and slightly rotted. The re- 
sulting cultures were about average but not always dependable. The 
fertilizers which gave cultures averaging the highest were dry buttermilk, 
soy bean meal, and cotton seed meal. Very little difference between them 
was noted. 



Sididae and Daphniidae 219 

Peak cultures seem to require from 30 to 50% more cotton seed meal 
than soy bean meal and the culture period is somewhat longer than with 
the former. On the other hand, cotton seed meal, like animal manures, 
stains the water a deep brown, resulting in deeply colored red Daphnia. 
The deep color seems also to prevent undue growth of blanket algae. The 
soy bean meal produces light gray Daphnia and an abundance of free- 
moving micro-algae which color the water green. It also encourages the 
growth of blanket algae. 

The quantity of Daphnia necessary to start a successful culture, within 
certain limits, is not so important as the physiological condition of the 
mother organisms. They should be active summer egg producers. Al- 
most always a few will be found with winter eggs. If the proportion is 
large, specimens from such a culture should be discarded. In the ex- 
periments reported here, from 25 to 100 cc. of mother organisms were 
generally used. We believe that 50 cc. is sufficient for 100 cu. ft. of water. 
They are measured by pouring water containing Daphnia into a tall 
graduate held in the sunlight. The individuals very soon settle to the 
bottom and the quantity may be determined with ease. 

The water supply of the seven concrete propagating ponds used is con- 
trolled by dams in such a way that a constant level is automatically 
maintained in each pond without overflow. The water is therefore stag- 
nant at all times. Each pond has an independent drain which leads 
directly into a bass-rearing pond. Hence all food organisms produced 
may be drained off directly into the rearing pond by removing a stand- 
pipe. During the last four years it has been customary to operate them 
in rotation, thus producing several crops in each during the summer 
season. 

M. E. D. 

DAPHNIA CULTURE 

Libbie H. Hyman, New York City 

TO CULTIVATE Daphnia, bring lettuce leaves, preferably of a green, 
leafy type of lettuce such as Boston lettuce, to a boil. Do not 
continue boiling. Place the boiled lettuce leaves in containers having 
6 to 8 inches of water in the proportions of 1 good-sizsd lettuce leaf to 
each 6 square inches of bottom. After 2 or 3 days add Daphnia. These 
may be obtained often from pet shops or dealers in biological supplies 
and also in almost any somewhat stagnant pond by means of a plankton 
net. After the lettuce leaves have disintegrated additional lettuce leaves 
should be added from time to time and also small pieces of raw liver or 
the entrails or corpses of small animals such as tadpoles, mice, rats, etc . 
In adding such material it is essential that only a moderate quantity be 
used. The water must not become foul or cloudy as this will kill the 



220 Phylum A rthropoda 

Daphnias. In general, the Daphnia culture should be kept supplied 
with fresh food, either lettuce leaves or raw flesh (preferably both) up 
to the limit possible without making the water cloudy. This limit must 
be learned by experience and depends upon the size of the container. 
Whenever it is seen that the Daphnias are not flourishing as rapidly as 
before, new food should be added. 

Daphnias may not be grown indefinitely in the same culture water. 
From time to time it is necessary to strain out the Daphnias and place 
them in freshly made cultures. When it is evident that the Daphnias are 
no longer multiplying well despite addition of fresh food, they should 
be transferred to an entirely new culture. 

References 

For the culture of Scapholeberis see below. 
For the culture of Simocephalus see p. 207. 
For the culture of Moina see p. 207. 
For the culture of Moina affinis see p. 213. 



Family 



CHYDORIDAE 



CHYDORIDAE 

Charles H. Blake, Massachusetts Institute of Technology 

IN RAISING Pleuroxus hamulatus I found that a few cc. of water in a 
Syracuse watch glass sufficed for an individual. The young were re- 
moved to separate glasses a few hours after birth, each being provided 
with freshwater which had been stored in contact with air and with 
Elodea. An aquarium provided a mixed culture of Stichococcus and 
Ankistrodesmus. [For culture see p. 227.] Enough of this was placed 
in the watch glass to form a thin, green coating on the bottom. After 
a few days most of it had been passed through the animal's gut and had 
become agglomerated and no longer suitable for food. The specimen 
was then transferred to a fresh glass made up as before. Crowding 
caused the production of two of the hitherto unknown males but all speci- 
mens were short-lived when crowded. Ephippia are formed in the absence 
of males or by crowding and, as in Scapholeberis, an individual may re- 
vert to the production of parthenogenetic eggs. Unfertilized ephippial 
eggs degenerate. Mass-cultures were not attempted with this species, 
but from observations in nature and on Alona guttata in the aquarium, it 
is apparent that a relatively great volume of water must be allowed per 
individual and an abundance of unicellular green algae as a source of 
food. The actual food material is unknown to me since Ankistrodesmus 
is apparently unharmed by its passage through the gut. Abundant 
oxygen is necessary. 



Copepoda 221 

Order copepoda 

CULTURE METHODS FOR PELAGIC MARINE COPEPODS 

George L. Clarke, Woods Hole Oceanographic Institution and Harvard University 

PELAGIC marine copepods are extremely delicate and sensitive to 
slight changes in their environment and no completely satisfactory 
technique for culturing them has yet been worked out. The methods 
described below are the result of preliminary experiments which have 
been undertaken at Woods Hole using the following species: Centro- 
pages typicus and C. hamatus, Labidocera aestiva, Acartia tonsa, and 
Calanus finniarchicus. 

Collecting. The copepods were collected by making short hauls with 
a plankton net from the laboratory power boat. Each haul was just 
sufficiently long (%-4 min.) to obtain the desired number of specimens 
(50-200), and no more, in order to avoid overcrowding. The mesh of 
the net was selected according to the size of the copepods desired, the 
coarsest possible net being used in each case so that the amount of other 
plankton material taken at the same time might be reduced to a mini- 
mum. The animals were protected from harmful crowding and abrasion 
by closing the tail of the net with a 2 -liter glass jar. At the end of the 
haul the glass jar was removed from the net and in the case of Centro- 
pages, Acartia, and Labidocera, which were obtained in Woods Hole 
Harbor and Vineyard Sound not more than 15 minutes' run from the 
laboratory, the copepods were left undisturbed in the jar (but protected 
from the sun) until the laboratory was reached. But in the case of 
Calanus, which could be obtained only in Vineyard Sound off No Man's 
Land (near the bottom) — requiring a 3 -hour trip back to the labora- 
tory — the catch was diluted and kept at a low temperature by placing 
the containers in the boat's ice box. Immediately upon arrival at the 
laboratory the copepods were transferred by means of a large-mouthed 
pipette to the containers to be used for culturing. 

Containers. The suitability of a variety of containers, including large 
battery jars, beakers, Erlenmeyer flasks, and small crystallizing dishes 
was tested. The size and shape of the container was not found to have 
any significant effect on survival as long as the animals were not unduly 
crowded. When the culture water was not changed, an allowance of 
20 cc. of water per copepod appeared to be adequate provided that the 
animals did not tend to cluster in one part of the culture dish. For the 
purpose of keeping track of the condition of individuals small containers 
were found most suitable. Erlenmeyer flasks of 250-500 cc. capacity, 
which were plugged with cotton stoppers or covered with inverted petri 
dishes, were used, and small crystallizing dishes (50 x 35 mm.), a num- 



222 Phylum Art hr op oda 

ber of which could be set in a large crystallizing dish and covered with a 
glass plate, were found especially convenient and were suitable for two 
copepods each. 

Water. The water used in the containers was ordinarily taken from 
the laboratory salt water tap. No improvement was found to result 
from using water brought in directly from the harbor or from Vineyard 
Sound. In a few cases the copepods appeared to fare perfectly well for 
a week or two without any change of water. In other cases the copepods 
were transferred after a certain number of days by means of a pipette 
to fresh containers, but this procedure was very laborious and often re- 
sulted in the loss or injury of some of the animals. A better method was 
to pour away all but a little of the original culture medium, leaving the 
copepods in the bottom corner of the dish and then to replenish with 
fresh seawater. It has not yet been determined how frequently it is 
necessary to change the culture water under various conditions. Very 
good survival was obtained in two sets of experiments, described in detail 
by Clarke and Gellis (1935), in which the culture medium was run 
continuously through flasks in which the copepods were confined. 

Stirring and Aeration. If the copepods were at all crowded they died 
off rapidly unless stirring in some form was provided. When beakers, 
or other wide-mouthed vessels, are employed, stirring may be accom- 
plished conveniently by using glass plungers activated by the Plymouth 
siphon device [Harvey, 1928, p. 57] or by an electric motor (Hagmeier, 
1930) . Such stirring provides at the same time a certain amount of aera- 
tion, which is probably sufficient in most cases. More effective aeration 
may be obtained by bubbling compressed air from a tap filter pump slowly 
through the water by means of glass tubes reaching to the bottom of the 
containers. The stream of bubbles brings about a slow circulation of 
the water which makes mechanical stirring unnecessary. This method 
is especially convenient for narrow-mouthed containers such as Erlen- 
meyer flasks. When small culture dishes were used with only a few 
copepods in each, stirring and aeration were found unnecessary. 

Temperature. Temperature was controlled by placing the Erlenmeyer 
flasks and the crystallizing dishes in a constant temperature tank where 
they rested half submerged on a wire rack. Two of these tanks were 
available and were maintained at different temperatures (kept constant 
to o.i° C.) by means of two Kelvinator cooling units operated by 
Hiergesell thermo-regulators and relays. A third and lower temperature 
(5-6 C.) was obtained by placing the containers in a large refrigerator. 
It was found that for all the species investigated the copepods died off 
rapidly if the temperature was allowed to rise above 20 C. Below 20 C. 
in the case of Calanus survival was improved progressively at lower 
temperatures down to 5-6 C, but the molting of shells became less 



Copepoda 223 

frequent. Further information on effect of different temperatures on 
other species is wanting, nor is it known to what degree constancy of 
temperature must be maintained. 

Illumination. The effect of light on the well-being of copepods has not 
been adequately investigated. Since the constant temperature tanks 
which were used in these experiments were open at the top only, the cul- 
ture dishes placed in them received indirect illumination reflected from 
the ceiling. Direct sunlight is certainly harmful and there is no doubt 
that the illumination should be weak. The copepods placed in the re- 
frigerator in complete darkness survived well for several weeks, and it 
is possible that light is not necessary for these animals at any time. 

Food. The question of what forms the chief food of copepods is a 
controversial one and is being intensively investigated at various labora- 
tories and from various angles. The matter has been reviewed by Clarke 
( 1934) and the specific experiments carried out at Woods Hole have been 
described in detail by Clarke and Gellis (1935). Since a satisfactory 
food for culturing has not yet been found, a brief statement of methods 
only will be attempted here. The amount of food and frequency of feed- 
ing required in different cases is highly variable for in early experiments 
with Centropages it was found that death supervened within a few days 
if food material was not added, whereas in the case of Calanus a few 
specimens lived for 14 days in continuously flowing water from which all 
particulate matter had been removed by a membrane filter. 

The survival of Centropages, Acartia, and Labidocera was improved 
by adding to the culture dishes planktonic material obtained by centri- 
fuging seawater, or in larger quantities, by making short hauls with a 
diatom net. "Persistent" cultures of diatoms and green flagellates grown 
in the laboratory* were also used. Experience showed that organisms 
which grow encrusted on the bottom and walls of the vessel were not 
suitable for food, probably because copepods, being filter feeders, can 
take in material in suspension only. However, the addition of none of 
these foods prolonged the life of the majority of the animals for more 
than about two weeks. When the flagellates were added to the water, 
green material could be seen in the intestines of the copepods and many 
excretory casts were found in the bottom of the container, but molted 
shells were observed only rarely. 

In the case of Calanus more elaborate experiments have been carried 
out since this article was originally prepared. Reference had best be 
made to the published reports (Fuller and Clarke, 1936, and Fuller, 
1937). Briefly, these experiments show that Calanus will live for several 
weeks and molt readily when provided with fine planktonic material. 
Bacteria are not, however, an important article of diet. Precisely which 

♦For method see Clarke and Gellis (1935)- 



224 Phylum Art hropoda 

elements of the plankton are the chief source of food in nature is still 
to be determined. When this question has been settled, it will probably 
be possible to grow the required food organisms in the laboratory and to 
keep Calanus and other copepods in culture indefinitely. (See addenda 

on p. 571.) 

Bibliography 

Bond, R. M. 1933. A contribution to the study of the natural food-cycle in 
aquatic environments with particular consideration of microorganisms and dis- 
solved organic matter. Bull. Bingham Oceanog. Coll. 4(Art. 4)11. 

Cannon, H. G. 1928. On the feeding mechanism of the copepods, C. finmarchicus 
and Diaptomus gracilis. Brit. J. Exper. Biol. 6: 131. 

Clarke, G. L. 1934. The role of copepods in the economy of the sea. Proc. Fifth 
Pacific Sci. Congress, Canada, 1933. 3^5.5:2017. 

an d Gellis, S. S. 1935. The nutrition of copepods in relation to the food- 
cycle of the sea. Biol. Bull. 68:231. 

Crawshay, L. R. 1 915. Notes on experiments in the keeping of plankton animals 
under artificial conditions. /. Mar. Biol. Assoc. N. S. 10:555. 

Dakin, W. J. 1908. Notes on the alimentary canal and food of the copepods. 
Intern. Rev. Hydrobiol. u. Hydrogr. i:77 2 - 

Esterley, C. O. 1916. The feeding habits and food of pelagic copepods and the 
question of nutrition by organic substances in solution in the water. Univ. Calif. 
Publ. Zool. 16:171. 

Fuller, J. L. 1937. Feeding rate of Calanus finmarchicus in relation to environ- 
mental conditions. Biol. Bull. 72. (In press) 

Fuller, J. L., and Clarke, G. L. 1936. Further experiments on the feeding of 
Calanus finmarchicus. Biol. Bull. 70:308-320. 

Hagmeier, A. 1930. Die Ztichtung verschiedener wirbelloser Meerestiere. 1930. 
In Abderhalden: Handb. biol. Arbeit., Abt. 9, Teil 5, Heft 4, Lief. 326, pp. 465-598- 

Harvey, H. W. 1928. Biological Chemistry and Physics of Sea Water. Cambridge 
University Press, Cambridge, England. 

Lebour, M. V. 1922. The food of plankton organisms. I. J. Mar. Biol. Assoc. 
12:644. 

Marshall, Sheina. 1924. The food of Calanus finmarchicus during 1923. Ibid. 

J 3 : 473- J . 

Murphy, Helen E. 1923. Life cycle of Oithona nana, reared experimentally. 

Univ. Calif. Publ. Zool. 22:449. 
Yonge, C. M. 1928. Feeding mechanisms in invertebrates. Biol. Reviews, 3:21. 
1931. Digestive processes in marine invertebrates and fishes. /. du Conseil 

6:175. 

Families calanidae and harpacticidae 

NOTES ON THE CULTIVATION OF TIGRIOPUS FULVUS 
AND CALANUS FINMARCHICUS 

R. M. Bond, Santa Barbara School, Carpinteria, California 

TIGRIOPUS FULVUS 

THIS harpacticid copepod is very large for the group. It has been 
reported from the Mediterranean, the coast of California, and else- 
where in warm climates. In suitable localities it is found, often in 
incredible numbers, in spray-pools above high tide mark. It may be 



Calanidae and Harpacticidae 225 

conveniently collected with a tea strainer or any sort of net. It is rarer 
and sometimes even difficult to find after a severe storm has washed out 
the pools which it inhabits, but in a few weeks is as numerous as ever. 

The animal may be raised in Syracuse dishes or in any larger con- 
tainers. The natural medium is seawater, which may be diluted as much 
as half, or concentrated to as much as 8% total salinity. The animal 
can withstand very rapid changes within these limits. I have usually 
used natural strength seawater. 

The food of these copepods probably consists of bacteria, but this has 
not been determined. They multiply rapidly if there is plenty of decay- 
ing vegetable matter in the medium, such as bits of seaweed, or even a 
piece of cheesecloth. The richest culture I ever obtained had a heavy 
growth of Platymonas subcordaejormis in it. The alga may have served 
as food, or simply to aerate the medium. 

If the food supply is at all abundant, it is necessary to use rather 
shallow containers or to employ compressed air for oxygenation. In 
either case, care must be taken to see that evaporation does not increase 
the salinity of the medium beyond the tolerance of the copepod. 

The optimum temperature seems to be about 25-30 C. 

CALANUS FINMARCHICUS, AND OTHER PELAGIC COPEPODS 

These copepods are not easily kept alive in the laboratory, and it 
is very difficult to raise adults from eggs. The greatest mortality seems 
to be in passing from the last copepodid stage to the adult. A number 
of different methods, however, have met with at least partial success. 
Miss Lebour* and others recommend the Plymouth plunger jar. [See 
p. 21.] For old stages I have had equal success with 500 cc. Erlenmeyer 
flasks containing about 300 cc. of seawater in which 4 to 6 copepods were 
kept. 

Probably the natural food consists mainly of small diatoms and other 
unicellular algae, Protozoa, and bacteria. Enough food is contained in 
seawater filtered through coarse filter paper if the water is kept flowing. 
Otherwise Nitzschia [For culture see p. 33.], Dunaliella [For culture 
see p. 134.] , or, still better, Platymonas subcordaejormis [For culture see 
p. 135] may be added (1-5% of algal culture). 

Calanus is found in waters with temperatures from about 4° to above 
20 C, but has a rather sharp upper limit of tolerance, which appears 
to be different in races from different regions. Specimens taken off 
Pacific Grove, California, lived several weeks at 17.5 C. Those from 
near Woods Hole died quickly above 15 C. It should generally be 
safe to keep the animals between 12 ° and 15 C. 

*Sci. Progr., London, 27:494, 1933. 



226 Phylum Arthropoda 

Medium and containers should be changed once a week or oftener 
to prevent the animals becoming entangled in bacteria. Calanus and 
several of the other pelagic copepods may be kept for some days or even 
weeks in a healthy condition in an icebox at 5°-6° C. if not too crowded, 
and if kept in containers wide enough to allow an ample air supply. 

References 

Family Diaptomidae 

For the culture of Diaptomus see p. 143. 



Family cyclopidae 

CULTURE METHODS FOR CYCLOPOID COPEPODS 
AND THEIR FORAGE ORGANISMS 

R. E. Coker, University of North Carolina 

Cyclops viridis, C. serrulatus, and C. vernalis have been reared very 
successfully when fed on a mixed bacteria and protozoan culture with 
occasional supplemental feeding from an algal culture; these cultures 
are prepared by one of the methods described below. 

PROTOZOAN CULTURES 

Horse Manure. Fill several quart jars each % full of fresh horse 
manure and nearly to the top with filtered tap water. After several days 
add a few pipettes of pond water or some mixed protozoan culture to the 
original jars; later jars are best inoculated from the best of the older 
cultures. Putrid conditions existing during the first few days seem 
effectively to sterilize the culture against copepods that might have been 
introduced with the tap or pond water. Rich cultures of Protozoa should 
develop within about a week; selection may then be made of the best 
as measured by richness in Protozoa and bacteria and freedom from 
cloudiness due to undesirable bacteria. A good jar may continue to 
furnish a satisfactory food medium for 1 or 2 months, but fresh cultures 
should be started at intervals of 2 or 3 weeks. 

Sheep Manure. Fill a jar % full of dried sheep manure (obtainable 
from seed or fertilizer stores) and nearly to the top with water. Treat 
like the horse manure cultures, although inoculation may not be 
necessary. 

Strong hay infusion. This is prepared by steeping hay, preferably 
timothy, in hot water for an hour or more (standard laboratory method) . 
Place the strained liquid in open dishes and, after a few days, inoculate 
as above. 



Cyclopidae 227 

ALGAL CULTURES 

Ankistrodesmus, or other unicellular alga. Culture methods may be 
found in botanical texts, but we have had best results with rich cultures 
of Ankistrodesmus that have developed naturally in aquaria in the 
laboratory in which small fish have been kept and fed with fish roe 
or in cladoceran cultures kept in bright sunlight and fed with small 
quantities of sheep manure infusion. The green water from the 
aquarium is passed through filter paper and kept in covered jars 
for observation during several weeks to guard against the presence 
of wild copepods. Occasional fertilization of these clean aquaria with 
sheep manure or with liquid fertilizers in small quantity may be de- 
sirable. 

Moitgeotia. A small dense tuft of the filamentous alga, Mougeotia, 
is held over a watch glass of water while successive close choppings are 
made with a scissors. This food medium then consists of short fragments 
of algal filaments, extruded protoplasm and water ; after stirring, a few 
drops may be used in place of the unicellular algae. Results have been 
excellent in growth and fertility of the Crustacea. 

CYCLOPS 

The amount of medium to be used and the frequency of feeding depend 
upon temperature and conditions of the experiment. In our practice a 
female with sacs is placed in a small vial with about 3 cc. of filtered 
pond or aquarium water and 1 cc. of mixed food (Protozoa with a little 
algae). As the manure cultures become older, dilution with pond water 
may be unnecessary. The culture water is changed or supplemented 
about every two days at 23 C. or about every two weeks at 6° C. The 
condition of the culture water may be appraised, and the need for sup- 
plemental feeding determined by looking through the vial toward a 
light; the liquid should neither appear cloudy nor have the excessive 
clearness indicative of food deficiency. Adjust feeding to maintain a 
reasonable abundance of Protozoa, evident to the eye, and a suggestion 
of greenness in the liquid or on the bottom of the vessel. Nauplii in 
quantity may be kept in larger containers with about 4 cc. of medium 
per nauplius and, if required, one or two supplemental feedings in the 
course of development. 

The methods employed have given maximum rates of growth (com- 
plete life cycles for C. vernalis in 7 days), high fertility through several 
generations, and virtually no disease or fungus. Copepods reared in the 
protozoan cultures without algae have lived well, and in many cases have 
grown rapidly, but they have displayed greater individual variation in 
rate of growth and less fertility than those given some plant food. 



228 Phylum Arthropoda 

References 

For the culture of cyclopids see also p. 230. 

For the culture of Cyclops see also p. 143. 
Family Harpacticidae 

For the culture of Tigriopus julvus see p. 224. 
Family Canthocamptidae 

For the culture of Canthocamptus see p. 143. 

For the culture of Copepoda see also p. 136. 

Order ostracoda 

SUGGESTIONS FOR CULTURING OSTRACODS 

Esther M. Patch and Lowell E. Noland, University of Wisconsin 

OSTRACODS are commonly found in nature at the bottom of pools, 
lakes, or sluggish streams, where there is dead organic matter 
which has passed the active fermentation stage of decay and is slowly 
decomposing in a medium where oxygen is present at least in small 
amounts. 

To rear ostracods in the laboratory it is desirable to duplicate natural 
conditions as closely as possible, for instance by a hay-wheat infusion 
made as follows: boil 2 grams of whole wheat grains and 3 grams of 
timothy hay in enough water to cover them. Pond or lake water will 
serve best since running water may carry pollutions, city water may con- 
tain chemicals used in purification, and distilled water may have un- 
desirable materials or lack desirable ones for optimum animal growth. 
Boil the hay and wheat slowly for about 10 minutes and replace the 
water with about a liter of that originally used, filtered or boiled so that 
no Entomostraca will be present. Set aside for a few days until the 
bacterial decomposition has passed from the acid to the alkaline stage, 
when the ostracods may safely be placed in the medium. More of the 
boiled wheat and hay and more of the boiled or filtered water may be 
added from time to time as required. 

Cultures of these small Crustacea thrive well in a wide dish, covered 
to retard evaporation but not so tightly as to exclude air. There are 
species differences among ostracods in the requirements for light and 
temperature as well as for food, but most of the common species do well 
at ordinary room temperature and in sunlight which favors the growth 
of algae. The above culture medium has been successful in rearing 
ostracods to feed to very small salamander larvae. Thin slices of potato 
(suggested by R. W. Sharpe in "Fresh Water Biology," edited by Ward 
and Whipple) have also been used, as well as decayed lettuce. 

Ostracods may usually be gathered with a weighted net from the mud 
or vegetation at the bottom of ponds and lakes ; at our laboratory they 



Isopoda 229 

have been obtained during the winter from mud drawn up through an 

opening in the ice. If the active forms are not available, they may be 

cultured from egg masses which are often present on submerged stones 

or plants and may even be found in viable form in the dried mud of 

seasonal pools. 

Reference 

For the culture of ostracods see also p. 136. 

CYPRIDAE 

Charles H. Blake, Massachusetts Institute of Technology 

THE usual ostracod of aquaria in the northeast is Cypridopsis vidua. 
This species may be taken from nearly any neutral or faintly acid 
pond, even if temporary, but is not likely to be found where the margins 
are marshy. 

Cypridopsis will persist for many months in fair numbers in any 
balanced freshwater aquarium. The eggs are laid on almost any solid 
object and all instars are easily found. If mud is present on the bottom, 
no other special food appears to be needed. 

Other ostracods may occur for short periods but I have not succeeded 
in keeping them more than a few weeks. They all require abundant 
oxygen. There appear to be no records of the reproduction of any 
marine form in captivity nor do young specimens survive more than one 
or two molts. 

Reference 

For the culture of Cypris see p. 143. 

Subclass Malacostraca, Order isopoda 

A SUGGESTED IMITATION OF A WOODLAND POOL 

Charles H. Blake, Massachusetts Institute of Technology 

CERTAIN animals live only in the presence of much waterlogged 
wood and fallen leaves, and in situations in which there are almost 
no algae. Early in March I brought in a small amount of mud con- 
taining a high proportion of waterlogged bits of twigs, leaves, etc., and a 
little sand. The freshwater isopod, Asellus communis, was rather com- 
mon in the situation where the material was obtained and some speci- 
mens were included in the gathering. 

An aquarium, about 6" x 18" in floor area, was set up with about %" 
depth of the mud together with such plants and animals as were present 
in the original bottom sample and 1" depth of stored aquarium water. 
The aquarium was covered with a loose-fitting glass plate as a protection 



230 Phylum Art hr op oda 

against dust. It was placed in a cool laboratory near a north window 
and shaded from other windows. By late May almost no algae have 
appeared and there has been no putrefaction. The death of various 
insect larvae and of some individuals of the amphipod, Dikcrogammarus 
jasciatus, has caused no evident upset in the equilibrium. Instead of the 
animals being balanced against plants they are balanced against the 
atmosphere. The aquarium contains a considerable number of % to 
Y 2 grown Aselli, offspring of the original individuals, and a variety of 
Entomostraca, including a few cyclopids which have maintained them- 
selves without noticeable increase in numbers. 



MAINTENANCE OF LAND ISOPODS 

John L. Fuller, Massachusetts Institute of Technology 

MOST woodlice are easily maintained in the laboratory on moist 
soil rich in humus. A wooden box 18" x 18" x 6" with a tightly 
fitting glass or wood cover will accommodate about a thousand animals. 
Land isopods should not be overcrowded, or molting individuals will be 
killed by their companions. In general, species should not be mixed, 
for the weaker species will be eliminated. If pieces of rotten wood and 
small stones are placed on the soil, the woodlice will congregate under 
them, making it easy to collect animals when desired. Water is added as 
necessary. 

A temperature of from 20 to 25 C._ is favorable for most species. 
Animals collected in the fall and kept between these temperatures will 
bear their first brood of young in February. Mating takes place readily 
when the two sexes are together; the fertilized females may be picked out 
since each carries its eggs in a ventral thoracic pouch. From 12 to 200 
young are released in about three weeks, the rate of development and 
number of young differing for different species. 

These animals appear to find sufficient nutriment in the organic matter 
of the soil, but will also eat slices of raw potato avidly. The nutritive 
requirements of the young are similar to those of adults. 

Greatest success has been obtained with Oniscus asellus, Porcellio 
scaber, Trachelipus rathkei, and ArmadiUidium vulgar e, though other 
species do nearly as well. Porcellio pic t us is less hardy, and apparently 
requires lime. Trichoniscus may be kept on wet moss in glass jars with a 
little water in the bottom. Particular care should be taken to keep 
Porcellio and Trichoniscus in a cool place. 

There appears to be no reason why healthy cultures may not be 
maintained indefinitely with a minimum of care. Lack of moisture is 
the chief danger to be avoided. 



Pandalidae 231 

Family ligydidak 

NOTE ON BLEEPING LIGIA OCEANIC A IX THE LABORATORY 

John Tait, McGill University 

KEEP animals in the darkness in a closed biscuit tin (of the English 
kind) the walls of which are lined with cardboard. Moisten the 
walls with freshwater or diluted seawater ( 1 part seawater to 3 parts of 
freshwater;. Give them a good foothold: do not keep them on glass. 
Clean the tin every four or five days by washing it out with seawater. 
Yarious kinds of animal and vegetable food were tried; the best results 
were obtained with blades of Laminaria. 

Bibliography 

Tait, J. Experiments on immersion of Ligia. Quart. J. Exper. Physiol. ,3:1. 

r- 191 7. Experiments and observations on Crustaceae. Pt. 1. Immersion 

experiments on Ligia. Proc. Royal Soc. Edin. 38:50. 

1925. The Sea-Slater, Ligia oceanica; a study in adaptation to habitat. 



The Scottish Naturalist. 151:13-18; 152:49-55. 



Order decapoda, Famih' pandalidae 

HATCHING ANT) REARING PANDALID LARYAE 
At.freda Berkeley Xeedler, Biological Board of Canada 

THE five Pacific coast species under consideration are Pandalus 
danae, P. borealis, P. hypsinotus, P. platyceros, and Pandalopsis 
dtspar. All of these species lay their eggs in the autumn, carry them over 
winter, and hatch them in the spring. The hatching begins about the 
end of February and continues until the end of May, being at its height 
about the beginning of April. 

Towards the end of February, therefore, five tanks were set up in 
the basement of the Biological Station. The tanks were made with 
wooden ends and bottoms and plate glass sides, and were about 2 feet 
long, 15 inches wide, and 18 inches deep. The wood and joints were 
covered with black asphalt paint such as is commonly used in hatcheries. 
The outlets led into beakers covered with muslin so that the larvae 
when hatched could not escape. The water came from a large concrete 
tank above the station and about 10 years old, which was daily pumped 
full of seawater from near the end of the station wharf. In previous ex- 
periments it had been found that this water often contained a fine brown 
debris which clogged the shrimps" gills and killed them. Partly to 



232 Phylum Arthropoda 

prevent this and partly to aerate the water each tank was fitted with a 
large funnel covered with muslin and with a tube leading to the bottom 
of the tank. Into each funnel a small but steady stream of water splashed 
from a tap about a foot above. When the larvae began to hatch a series 
of nursery tanks made of battery jars were fitted up in a similar manner 
and a few large beakers were also used. 

It was found necessary to have the eggs carried until hatching by the 
females, as larvae were never successfully hatched from eggs that had 
become separated. Ovigerous females were taken with a small beam 
trawl on various grounds sometimes 50 miles away or more. During 
transportation to the station they were carried in a large galvanized tub 
with changes of water at least every hour. They were placed in the big 
tanks and kept there until their eggs had hatched. During their cap- 
tivity they were fed mainly on chopped crab's liver and marine worms. 
Most of them kept healthy although some of those from deep water 
acted as if they were blind and bruised themselves badly against the 
sides of the tanks. 

In many cases the larvae appeared to be healthy for a week or ten 
days and then died before molting. A few Pandalus danae larvae reached 
the second stage but none of the others ever succeeded in passing the first 
molt (although Spirontocaris and Crago larvae were easily reared in 
beakers through several stages). 

The salinities in the tanks compared favorably with the natural ones. 
Moreover no sudden ill effects were noted on the days that the salinity 
dropped and experiments conducted in the laboratory had indicated 
that the adult shrimps at least could stand a fairly wide range of salinity. 

The pH of the water in the tanks remained close to that in the sea 
near the station. 

As the adult shrimps all came from comparatively deep water (20 to 
60 fathoms) they were subjected in the tanks to much lower pressures, 
and perhaps in some cases to stronger light than normal although the 
latter was kept subdued. The adults, however, appeared healthy during 
hatching. The larvae occur naturally from a depth of about 4 fathoms to 
the bottom, the younger stages keeping nearer the surface. It does not 
appear probable, therefore, that light or pressure were adverse factors in 
the rearing of the larvae. 

The larvae were given various foods — eggs of marine worms, plankton, 
and finely minced crab liver. Apparently they ate the food, as traces 
could be found in their alimentary tracts. Examination of larvae ob- 
tained in plankton hauls indicated that they had been feeding on the 
smaller plankton organisms. It is quite possible that unsuitable food was 
one of the chief adverse factors. 



T 



Homaridae 233 

Family crangonidae 

A METHOD FOR REARING SMALL CRANGONIDAE 

Hugh H. Darby, College of Physicians and Surgeons 
HE rearing of these organisms seems to have given trouble to 



JL several investigators. By the following technique it has been pos- 
sible to keep them alive for at least three months. Small embryological 
dishes, about 12 cm. in diameter, were used, one for each animal. If 
two are placed together in the same dish, they tear each other to pieces. 
No running water was used, but the seawater was changed once every 
2 or 3 days. Some of the small Synalpheus were kept in similar dishes 
without any change of water for 10 days without ill effects. A change 
of water just after molting is harmful. Much better results were obtained 
when the organisms were kept for at least 24 hours in the same water in 
which they had molted. During this time they devoured their own cast 
skin, and needed no other food. Ordinarily the muscles of small fishes 
or of other Crustacea were supplied for food. If kept in the ice box for 
two or three days before use, the food was devoured more readily than 
when given fresh. Very small pieces were given, so that there was no 
remnant left to become a source of bacterial infection in the culture 
dishes. Small marine algae kept in the dishes furnished both aeration 
and food. Larvae need much more aeration than adults for survival. 
Temperatures between 22 ° and 30° C. were entirely satisfactory for 
organisms found in the Dry Tortugas islands. 

Bibliography 
Darby, Hugh H. 1934. The mechanism of asymmetry in the Alpheidae. Carnegie 
Inst, of Wash. Publ. No. 435. 



Family homaridae 



HATCHING AND REARING LARVAE OF THE AMERICAN 
LOBSTER, HOMARUS AMERICANUS 

Paul S. Galtsoff, U. S. Bureau of Fisheries 

THE range of the American lobster extends from Labrador to North 
Carolina. Every spring lobsters migrate from deep water inshore 
where they remain until the onset of cold weather. At Woods Hole, 
Massachusetts, they usually appear in large numbers in May and begin 
their outward migration in October. Many of them remain in relatively 
shallow water throughout the winter. 

The copulation of lobsters occurs primarily in the spring. During this 
time the sperm is deposited in an external seminal receptacle situated 



234 Phylum Arthropoda 

between the bases of the third pair of walking legs of the female where 
it retains its vitality for several months. The eggs are fertilized after 
ejection from the oviducts and are attached to the hairs of the swim- 
merets by a cement-like substance secreted by special glands of the 
female. Observations of Herrick show that regardless of its sexual con- 
dition the female lobster may be approached by the male more than 
once. About 80% of the spawning females lay their eggs in July and 
August, the remainder extrude their eggs in the fall and winter. The 
peak of the spawning season at Woods Hole occurs during the latter 
part of July and the first half of August. In Maine it is two weeks later. 

The eggs are carried by the females from 10 to 11 months before 
they are hatched. The newly laid eggs measure from 1.5 to 1.7 mm.; 
they are of dark olive green color and are easily distinguished from the 
light-colored old eggs in which the green yolk has been absorbed by the 
embryo. In the freshly laid eggs the yolk, invested by a transparent cap- 
sule, is of uniform granular texture. In 20 to 25 hours after oviposition 
large yolk segments are distinguishable by the unaided eye. In about 
10 days the embryo reaches the egg-nauplius stage and in 26 to 28 days 
the eye pigment may be seen at the surface. Hatching eggs may be 
obtained at Woods Hole from the middle of May until the first half of 
August. According to Herrick the number of eggs laid by the female 
at each reproductive period increases in geometrical proportion while the 
length of the female increases in arithmetical proportion. Thus, females 
8, 10, 12 and 14 inches long would lay 5,000, 10,000, 20,000, and 40,000 
eggs respectively. The relation holds good up to the 14-inch size. 

Owing to the unequal rate of development the hatching period of a 
single brood may last about one week. 

The following method is used in obtaining eggs for hatching: An egg- 
bearing female (with light-colored eggs) is stretched on its back over 
the table. By cautiously moving the dull side of a knife pressed against 
the abdomen the eggs are detached from the swimmerets and are imme- 
diately placed in a hatching jar (Fig. 47 A). A strong stream of water 
delivered through a glass tubing, almost reaching the bottom, keeps the 
eggs stirred, while the newly hatched larvae are carried by the vertical 
current of water into the tank (T). A screen (S) surrounding the over- 
flow prevents their escape. 

A newly hatched larva is a transparent, actively swimming organism, 
about 8 mm. in length. Its pelagic life continues for about two weeks 
during which time it grows and molts three times. After the 4th molt 
the larva bears a striking resemblance to the adult lobster but it has the 
larval rostrum and its front abdominal somites are still without the 
appendages. At this stage its color may be bright red, green, or reddish 
brown. The manner of swimming also has changed; the larva rapidly 



Homaridae 



235 



moves forward by means of the swimmerets and darts backward by 
sudden jerks of the abdomen. It measures between n and 14 mm. 
All the traces of larval swimming organs disappear after the 6th molt. 
The ensuing molts follow at rather short intervals. It has been estimated 
that during the first year the lobster molts from 14 to 17 times and 
attains a length of from 2 to 3 inches. 

The first three stages comprise the most critical period of the larval 
life of the lobster. The natural food of the larvae probably consists of 
pelagic copepods and other crustaceans. In the hatching tanks they 
display strong cannibalistic tendencies, usually attacking their victims 
from above and nipping into the abdomen at its junction with the cara- 
pace. The self-destructiveness of the larvae constitutes the greatest diffi- 
culty in rearing them under artificial conditions. The best method of 
overcoming this consists in preventing the aggregation of the larvae on 
the bottom and by keeping them afloat. This is accomplished by a con- 
tinuous stirring of the water by means of a propeller (P) operated by 
an electric motor (not shown in Fig. 47). 




Fig. 47. — Method of hatching and rearing lobsters. A, hatching jar; P, propeller; 

S, screen; T, tank. 

Experiments carried out by the author at Woods Hole prove that 
boiled and dried fish is the best food for the larvae. A whole fish is cut 
in several pieces and boiled in freshwater for about 30 minutes, then 



236 Phylum Arthropoda 

placed in a porcelain dish and dried in an oven for about 12 hours at 
55-60 C. For feeding, a piece of the dried fish is ground in a glass 
mortar and a small amount of it distributed throughout the tank. 

Several kinds of fishes were tried. The best results were obtained 
with mackerel. Besides fish meat, dried and ground hard boiled eggs have 
been successfully used at the Rhode Island State Lobster Hatchery. 

Care must be exercised to remove the unused meat and prevent the 
decomposition of the debris and of dead larvae. 

By using fish food and exercising reasonable precautions, the author 
had no difficulty in rearing the lobsters to the 4th and 5th stages, which 
were reached in 15-17 days. 

Young lobsters may be fed on various animal food, as for instance, 
pieces of clam, oyster, crab meat, etc. They should be kept in a tank 
containing small rocks and a little sand. 

Bibliography 

Herrick, F. H. 1896. The American lobster, a stud}' of its habits and develop- 
ment. Bull. V. S. Bur. Fish. 15:1. 
Mead, A. D. 1908. A method of lobster culture. Ibid. 28:221. 

Family astacidae 

A CRAYFISH TRAP* 

IN PONDS and streams where crayfish are abundant they may readily 
be taken by means of a trap constructed as follows: 

A rectangular box of any convenient size, 16 x 24 inches for instance, 
is built of %-inch mesh galvanized screen wire. Into one end of this 
box a removable funnel of like material is fitted. This funnel should 
project about 8 inches into the box and have a flattened opening about 
4 inches wide and 1% inches deep. 

In setting the trap it should be placed in shallow water on a sloping 
bank and partially embedded in the mud or sand so that the bottom of 
the funnel is even with the bottom of the pond. The rest of the trap 
extends out toward the deeper water. A dead fish wired securely to the 
bottom of the trap makes an excellent bait. Attracted by this bait, the 
crayfish crawl into the trap and seem to be unable to find their way out. 
A single night-set with such a trap will reward the trapper with at least 
a water bucket full of crayfish for laboratory use, or for the more im- 
mediate purpose of providing the camp with an exceedingly delectable 
breakfast. 

M. E. D. 

* Reprinted, with slight changes, from Science 55:677, 1922, by E. C. O'Roke, University 
of Michigan. 



Astacidae 237 

Reference 
For the feeding of crayfish see note on p. 49. 

CULTURE METHODS FOR BRACHYURA AND ANOMURA 

Josephine F. L. Hart, Pacific Biological Station 

THE methods of successfully maintaining and rearing crabs and 
hermit crabs in the laboratory are not entirely dependent on the close 
simulation of natural conditions, but on careful feeding and preservation 
of hygiene during the progress of the experiment. 

Immediately on collection of the specimens, a mature male and female 
should be placed together in a container. The larger crabs are best 
preserved in live boxes or aquaria with circulating seawater, but the 
small ones may be placed in shallow dishes, approximately 3 liters in 
capacity, in which cases the water should be changed daily. 

It has been observed that many species of crabs, scavengers, carnivo- 
rous and herbivorous feeders, will live under such conditions, when fed on 
finely minced, fresh, clam muscle. It is probable that there are more 
suitable foods (Orton, 1927), but in the instances that have come under 
direct observation this diet has been found satisfactory. 

If sand or other natural bottom material is placed in the jars, detritus, 
rotting food, and feces may not be effectively removed, and the accumu- 
lated decomposition products tend to cause pollution and the subsequent 
growth of detrimental bacteria and Protozoa. If the specimens are kept 
in a live box or a barren aquarium with running water, these factors are 
negligible. If the individual dishes or plunger jars (Brown, 1898; 
Lebour, 1927) are used, they should be cleaned daily about three hours 
after feeding. During the cleaning process, the animals should not be 
handled, in order to avoid the possibility of physical damage to them. 

Copulation has been observed to take place under these conditions in 
Scleroplax gramdata and Lophopanopeus bellus, followed by the de- 
position of the eggs. The females were then isolated in separate con- 
tainers and the development of the eggs observed. 

The larvae, upon hatching, swim to the surface of the water and 
should be removed with a pipette and placed in large beakers of freshly 
obtained seawater, which may be aerated by stirring with a glass rod. 
The larvae should be examined daily, fresh water and food given, and 
all the dead material and sloughed skins removed. This is most easily 
accomplished by transferring the active larvae to a fresh container, 
already supplied with food material. 

The chief difficulty encountered in rearing decapod larvae is the 
maintenance of a constant supply of suitable living food. Many of the 
forms eaten in the natural state, when placed in laboratory conditions, 
soon die and their decomposition products in the water kill the larvae. 



238 Phylum Arthropoda 

Lebour (1927 and 1928) was successful in rearing three species of crabs 
in plunger jars on a diet of the larvae of Ostrea, Teredo, Echinus, and 
Pomatoceros. The megalopae and young crabs were fed on small pieces 
of the mantle of Mytilis edulis. The larvae of the native oyster, Ostrea 
lurida, have been found to be a satisfactory food, as they are held for 
some time in the mantle cavity of the adult and therefore may be ob- 
tained in quantity, yet are free-swimming when placed in the water. 
The veliger larvae of Nudibranchs and the trochophore larvae of the 
Japanese oyster were also used, but did not prove as satisfactory as 
those of the native oyster. When the larvae are no longer free- 
swimming, living food material may be replaced by minced clam muscle, 
on which the megalopa and young crabs will continue to thrive. It is 
advisable to provide shells for the glaucothoe and young stages of the 
Pagurids, and for this purpose the broken off tips of the spirals of 
Littorina were found suitable. 

The length of time spent in each stage seems to depend considerably 
on the relative abundance of food, the temperature and salinity of the 
water, and on other such external conditions. The first zoeal stage, in 
suitable natural conditions, probably lasts for 2 or 3 days, and the time 
spent in each stage increases as the larva grows. Under laboratory con- 
ditions, 4 to 5 weeks is usually required for the development from the 
egg to the young crab stage. The 6th young crab stage of Hemigrapsus 
nudus appeared 2 months later. 

These methods have been applied successfully to the rearing of both 
Brachyura (Hart, 1934) and Anomura. Some slight variations in tech- 
nique may be found advisable and these will become evident with the 
development of the experimental work. 

Bibliography 

Brown, E. T. 1898. On keeping medusae alive in an aquarium. J. Mar. Biol. 
Assoc. 5:176. 

Hart, J. F. L. 1935. The larval development of British Columbia Brachyura. 
I. Xanthidae, Pinnotheridae (in part) and Grapsidae. Canad. J. Res. 12:411. 

Lebour, M. V. 1926-27. Studies of the Plymouth Brachyura. I. The rearing of 
crabs in captivity, with a description of the larval stages of Inachus dorsettensis, 
Macropodia longirostris and Maia squinado. J. Mar. Biol. Assoc, n.s. 14:795. 

1928. The larval stages of the Plymouth Brachyura. Proc. Zool. Soc. Lon- 
don, pp. 473-560. 

Orton, J. H. 1926-27. On the mode of feeding of the hermit crab, Eupahurus 
Bernhardus, and some other Decapoda. Ibid. 14:909. 



Cancridae 239 

Family cancridae 

NOTES ON REARING THE PACIFIC EDIBLE CRAB, 
CANCER MAGISTER 

Donald C. G. MacKay, Pacific Biological Station 

THE Pacific edible crab, Cancer magister, described by Dana in 1852, 
is a large crustacean which sometimes reaches a carapace width of 22 
cm. In Alaska, British Columbia, Washington, Oregon, and California 
the species is present in large numbers. 

LIFE HISTORY 

Evidence from several independent sources leads to the conclusion that 
females become mature at a carapace width of approximately 10 cm. 
The size at which sexual maturity is attained in males has not been 
ascertained definitely but it is known that some are mature at a width of 
13.5 cm. Maturity is ordinarily attained by females in the 4th or 5th 
year though, in some instances, it is believed to occur as early as the 
3rd year or as late as the 6th year. The normal duration of life in 
British Columbia is probably about 8 years and the maximum is prob- 
ably not more than 10 years. 

The mating season is from April until September. Mating takes place 
on the tide flats and invariably occurs between a soft-shelled female and 
a hard-shelled male. The male embraces the female, sternum to sternum, 
and the two lie buried in the sand, mud, or seaweed in a nearly vertical 
position. 

The average diameter of fully developed external eggs is about 0.47 
mm. During development the ovarian eggs undergo color changes from 
white to coral red; these changes are closely correlated with the sizes of 
the eggs. The number of eggs produced by Cancer magister is large, the 
actual number depending upon the body size. In two specimens exam- 
ined the numbers were 750,000 and 1,500,000 for crabs 5.33 and 6.00 
cm. in carapace width respectively. It is estimated that one female in a 
lifetime might produce 4,000,000 eggs. 

The ovigerous period in British Columbia is from October until June. 
Hatching probably occurs from December until June and there is reason 
to believe that the season is somewhat earlier in California. 

The eggs hatch as protozoeae and pass through several zoeal instars 
and one megalops instar, after which they closely resemble the adult in 
form. 

The natural sizes of the megalops and first five post-larval instars have 
been determined and are respectively as follows: 0.28, 0.52, 0.74, 0.97, 
1.34, and 1.82 mm. Contrasted with the natural increases just men- 



240 Phylum Arthropoda 

tioned the experimental crabs are likely to become retarded in growth. 
That is to say that when they molt they are likely to increase less in 
size than would be the case in nature. This effect appears to be cumu- 
lative, crabs molting more than once falling even further behind in size. 
Large crabs kept in compartment live-wells, though comparatively more 
cramped than the small crabs kept in the laboratory, show less retarda- 
tion of growth upon molting. 

SEASONS OF AVAILABILITY 

Megalops are to be found during July and August and crabs in the 
first three post-larval instars during August and September. By the 
following spring these same crabs, then i year old, have reached the 
6th and 7th instars. 

Mating crabs are occasionally found on the tideflats among seaweed. 
Molting crabs may sometimes be discovered beneath the inverted shells 
of bivalve molluscs or otherwise hidden at extreme low tide. Crabs on 
the verge of molting appear to frequent shallow water and are to be 
found in large numbers buried in the sand in certain localities during the 
spring of the year. Except for the fact that the species characteristically 
inhabits sandy or slightly muddy regions and avoids rocky shores, we 
can give no rules for finding such areas. Crabs ordinarily found are from 
3 to 15 cm. in carapace width. For larger material the investigator must 
either depend upon fishermen or else employ traps of his own. 

METHOD OF COLLECTING 

Crabs in the megalops and early post-larval stages may be collected 
by hand in small numbers on the shore at low tide. The megalops, which 
are positively phototropic, are frequently found adhering to barnacles 
and other small objects; occasionally they are found swimming at the 
surface, sometimes in large numbers. In the latter event they are ex- 
tremely difficult to capture. 

The early post-larval crabs may also be collected by hand in small 
numbers at low tide. They are, however, ordinarily found buried in the 
sand or otherwise hidden from view. When the sand is disturbed they 
may sometimes be observed digging themselves rapidly in again. Crabs 
at this stage of development may often be taken in large numbers from 
the gear of fishermen operating on the ordinary commercial fishing 
grounds which, in Boundary Bay, B. C, are about 10 to 12 fathoms in 
depth. 

HANDLING THE MATERIAL 

Young crabs appear to be somewhat hardier than mature crabs and 
may be handled without extreme care. In all cases, however, it is im- 



Cancridae 241 

portant to avoid crowding a large number of individuals within a small 
quantity of water. The chances of survival, where crowding cannot be 
avoided, would seem to be best where water is not used. When covered 
with damp seaweed crabs will live for many hours and often for days. 

REQUIREMENTS FOR KEEPING ALIVE 

Small larval and post-larval crabs were successfully kept by the writer 
for considerable periods of time, sometimes for several months, in mis- 
cellaneous glass vessels of all available kinds. These containers were 
mostly of the type in which foods are put up and were the only vessels 
readily available in the small seaside community where the work was 
carried on. Before being used, all the containers were carefully washed 
in boiling water. In general large shallow vessels seemed to be the most 
successful. 

Into each container was placed one small crab, some clean sand, one 
or two pebbles, and a piece of seaweed. Small sticklebacks were found 
useful for keeping the water in circulation in the individual containers 
and were sometimes used for this purpose. It was, however, necessary 
to replace the small fish from time to time since, though they were con- 
siderably larger than the crabs, they were frequently caught and partially 
devoured. 

Running water was unfortunately not available but satisfactory results 
were obtained by changing the water twice daily. 

The larger crabs were kept in compartment live-wells anchored to a 
float. The live-wells were designed to facilitate the natural circulation 
of water due to the tidal currents. Compartments of various sizes were 
provided for large and small crabs. Live-wells of 3 feet x 4 feet x 9 
inches were found to be as large as could conveniently be handled and 
two of these dimensions were put in use for two seasons. 

FOOD 

The young crabs were fed mainly on small pieces of absolutely fresh 
sticklebacks, a decided preference for fresh rather than stale fish having 
been shown in feeding experiments. Sticklebacks were readily obtain- 
able with the use of a small dip net and proved very acceptable to the 
crabs. Small pieces of oyster were occasionally given but proved to be 
less satisfactory than the fish. 

Fresh fish heads and entrails were used as food for larger crabs. 

References 

Family Xanthidae 

For the culture of Lophopanopeus bellus see p. 237. 
Family Grapsidae 

For the culture of Hemigrapsus nudus see p. 238. 



242 Phylum Arthropoda 

Class Arachnoidea 

FEEDING NOTES FOR CERTAIN ARTHROPODS 

Lucy W. Clausen, American Museum of Natural History 

THE following is a list of arthropods which have been kept for com- 
paratively long periods of time in the cages containing live exhibits 
in the Hall of Insect Life of the American Museum of Natural History, 
together with the food organisms which have proven satisfactory for 
each. 

ARACHNIDA 

Scorpions (several species) have been fed on mealworms and Oriental 

roaches. 

Tarantulas will eat roaches and mealworms. 

Pholcids (false Daddy-long-legs) eat Drosophila when small and 
larger flies, such as Musca, when larger. 

The wolf spider, Lycosa carolinensis, has fed upon flies and meal- 
worms. 

The garden spider, Aranea serkata, has been fed flies and grass- 
hoppers. 

The crab spider, Olios sp., is fed mealworms. 

ORTHOPTERA 

Roaches (American, Oriental, and tropical) are fed sliced potato, 
lettuce, bananas, bread, and a piece of bacon occasionally. They seem to 
relish spinach when it is not given to them too often. 

Crickets are fed on apple, lettuce, and bread. 

Meadow grasshoppers are fed on apple and grass. 

Mantids (Chinese Mantis) eat Drosophila when small. When adult 
they eat houseflies and mealworms. The latter should be dangled before 
them on a thread to attract their attention. 

COLEOPTERA 

Tiger beetles (Cicindela sexgutata, C. dor salts, etc.) have been fed 
on mealworms cut in sections, and on apple. 

Caterpillar hunters (Calosoma calidum and C. scrutator) have been 
fed on mealworms and apples with banana occasionally. 

The ground beetle, Harpalus caliginosus, has been fed mealworms and 
apple. 

Dytiscus eats small mealworms which are dropped into the aquarium. 

Mealworms (Tenebrionidae) are grown in bran with a slice of potato 
added every other day. 

The red rust flour beetle (Tribolium jerrugineum) grows well in whole 



Theraphosidae 243 

wheat flour. Enough moisture is supplied to the culture by keeping a 
moist wad of cotton attached to the lid of the container. 

Dermestids, such as D. lardarius, are fed on felt, bones which have a 
bit of dried substance adhering, and ordinary dog biscuit (which they 
seem to like). 

In all cases moisture is extremely necessary, but care must be taken 
so that mold does not set in. 

Order araneae, Family theraphosidae 

LABORATORY CARE OF TARANTULAS 

W. J. Baerg, University of Arkansas 

TARANTULAS hatch late in summer, August or September. During 
the fall they apparently require no food. In the following spring 
a large family confined in a battery jar will begin to dwindle. In May 
and June cannibalism becomes so severe that if several specimens are 
to be saved for rearing to the adult stage they must be isolated. These 
young individuals may be kept in any sort of small jar that will confine 
them and admit some air. Small sized battery jars (4" x 6") do very 
well. A thin layer of soil (about % inch) is desirable; a deeper layer 
may crush the delicate spiders when they burrow. 

Young tarantulas will feed on any small insects they can handle. 
Termites are very satisfactory and will serve till the tarantulas are 
about 3 years old. 

Older tarantulas are conveniently kept in large battery jars which 
should be half filled with soil to satisfy their desire for digging. They 
may be fed grasshoppers, crickets, cockroaches, or a variety of cater- 
pillars, as well as various moths, butterflies, cicadas, etc. Tent cater- 
pillars and catalpa worms serve well for early spring and summer, grass- 
hoppers for late summer and fall. 

Tarantulas of the common local species Eurypelma calijornica will 
feed once a week or a little oftener; the Mexican species Dugesiella 
crinita will accept food more often and in larger quantities. Mature, 
or nearly mature tarantulas will live without food for about 2 years. 

Water should be supplied at frequent intervals. A glass dish, such as 
a small petri dish, serves well. In seasons of severe drought tarantulas 
may succumb to thirst in 2 months' time. 

During the winter the tarantulas may doubtless be kept in an ordinary 
laboratory, but to come nearer to natural conditions a basement, not 
heated, is better. A light frost will not kill them, but it is well to avoid 
temperatures below 40 F. This is for the local species E. calijornica. 
Species native farther south should be kept at higher temperatures. 



244 Phylum Arthropoda 

When approaching a molt tarantulas cease feeding. No apprehension 
need be felt over this lack of appetite even if it lasts for 2-3 weeks. 

If females heavy with eggs are brought into the laboratory they will 
in time produce cocoons. These, if kept in a sufficiently warm room 
exposed to sunlight , may produce young (600-1200). A more convenient 
method of rearing young is to bring in cocoons from which the spiderlings 
are about ready to emerge. The time required from oviposition to 
emergence of young is about 6 weeks. 

KEEPING AVICULARIA AVICULARIA IN THE LABORATORY 

Mary L. Didlake, University of Kentucky 

GLASS stender dishes have furnished satisfactory, though rather 
cramped, quarters for tropical spiders found in bunches of bananas 
and brought to me. One large Bird Spider (Avicularia avicularia) I have 
kept now for seven years. It has molted eleven times and measures 2% 
inches from the front of the cephalothorax to the tip of the abdomen. 
I use a pair of long forceps for moving the spiders to clean quarters. 
They drink readily from a smaller, 2 -inch stender dish, catch living food 
put in for them (caterpillars, grasshoppers, roaches) and will, when very 
hungry, feed on a piece of raw liver or beef, sucking it white. A fledg- 
ling English sparrow was consumed once, and on two occasions a small 

mouse. 

Specimens have usually been females and no attempt was made to 
rear successive generations. The jars were kept at room temperature. 
In very cold weather, and over week-ends when the room was likely to 
be chilly, they were set on top of an incubator which furnished a slight 

degree of warmth. 

References 
Family Pholcidae 

For feeding see p. 242. 
Family Lycosidae 

For feeding of Lycosa carolinensis see p. 242. 

Family theridiidae 

CULTURE OF LATHRODECTUS MACTANS, THE BLACK 

WIDOW SPIDER 

Elizabeth Burger, The College of William and Mary 
Caution: Extreme care should be taken in handling "black widow" 
spiders since their poison is highly toxic and may prove fatal. 

Containers. Since these spiders are cannibalistic, they must be kept 
in individual containers. Glass tumblers, covered with cheesecloth 
secured by a rubber band, with one inch of sand at the bottom, are 



A carina 245 

satisfactory. Water and food are introduced through a thistle-tube 
inserted through a hole in the cover. (This hole in the cloth is kept 
below the margin of the tumbler when not in use.) A small layer of 
non-absorbent cotton protects the spider from drowning when the sand 
is moistened. 

Food. Adult spiders should be fed upon grasshoppers and other 
Orthoptera, one insect every three or four days. When these are not 
available, cultured blue-bottle flies (Calliphora erythrocephala) are 
sufficient. [See p. 415-] 

Since the newly hatched spiders apparently are to a large extent ex- 
clusively cannibalistic, it has been considered necessary to permit them 
to feed upon each other for the first three or four days. To conserve 
stocks the spiders should be separated after this period and fed fruit 
flies (Drosophila spp.). [See p. 305.] 

Temperature. For storage: 40 to 50 F. For breeding, oviposition, 
and hatching of young spiders: room temperature. 

Light. Spiders should not be exposed to direct sunlight. 

Moisture. A somewhat damp habitat is natural. Satisfactory con- 
ditions of moisture are secured as described above. 

Breeding. Females captured in the fall will lay fertile eggs. Virgin 
females raised through the winter may be mated with males, which may 
be distinguished from immature females by their bulbous palpi. 

Egg cases should be separated from adult spiders before hatching. 
The eggs hatch within three or four weeks. 

References 
Family Argiopidae 

For feeding of Aranea sericata see p. 242. 
Family Thomisidae 

For feeding of Olios see p. 242. 

Order acarina 

CULTURE OF NON-PREDACIOUS, NON-PARASITIC MITES 
(ORIBATOIDEA AND TYROGLYPHOIDEA) 

Arthur Paul Jacot, Asheville, North Carolina 

A CELL FORMED of a microslide, a glass ring, and a large cover 
has proved most satisfactory. The rings may be 20 x 5 mm. with 
a # 2 cover glass 22 mm. in diameter. The center of the slide should 
be etched (with hydrofluoric acid blocked in with paraffin) to give it 
a rough surface to enable certain species to walk with ease. The ring 
may be fastened with Canada balsam. The top of the rim must be 
coated with a very thin film of vaseline, paraffin, or some other sub- 
stance which will make an air-tight seal with the cover, or the included 



246 Phylum Arthropoda 

moisture may escape. It is essential to keep the cells moist. In some 
cases a piece of blotting paper on the cell-floor will be satisfactory. The 
animals may be fed on bits of moss, lichens, cheese mold, or on soft, 
moist, dead wood. The cells must be inspected daily to regulate the 
moisture content and the growth of molds. 

References 
Family Tyroglyphidae 

For culture, including that of Tyroglyphns linteri, see p. 266. 



Family 



IXODIDAE 



TICK REARING METHODS WITH SPECIAL REFERENCE 

TO THE ROCKY MOUNTAIN WOOD TICK, 

DERMACENTOR ANDERSONI STILES* 

Glen M. Kohls, United States Public Health Service 

TICKS belong to the order Acarina, superfamily Ixodoidea, which 
is composed of the families Ixodidae and Argasidae. 
Tick rearing is an involved process because of complicated life cycles, 
the blood feeding habit which necessitates the use of host animals, and 
the different environmental requirements of different species during 
aestivation and hibernation and other periods when not on host animals. 
Rearing methods applicable to specific problems have been developed 
by different groups of investigators, but lack of space makes it necessary 
to limit this article to those methods and equipment now in use at the 
Rocky Mountain Laboratory for the rearing of the Rocky Mountain 
wood tick, Dermacentor andersoni. These methods have been developed 
by several investigators over a considerable period of years in connection 
with study of tick-borne diseases of the United States, and especially in 
relation to that of Rocky Mountain spotted fever and the manufacture 
of spotted fever vaccine. This vaccine is prepared from the tissues of 
infected adult D. andersoni and necessitates the rearing of this species 
in large numbers. The methods described are applicable in general to 
other species of ixodid ticks. 

D. andersoni is a three host tick, the adults of which are active in 
nature from the latter part of March to about July 1st. They are 
usually found on livestock and the larger game animals, and attach 
readily to man. When host contact is made the ticks attach and feed to 
repletion in 8 to 14 days, copulation occurring while the females are still 
attached. The latter increase enormously in size due to the blood meal, 
leave the host and, after 2 to 4 weeks, deposit 4,000 to 7,000 eggs in a 

♦Contribution from the Rocky Mountain Laboratory, United States Public Health Serv- 
ice, Hamilton, Montana. 



Ixodidae 



247 



suitable place. The larvae, hexapod and sexually indistinguishable, 
infest rodents during June, July and August, require 4 to 7 days for 
engorgement, then drop and molt to nymphs before the onset of cold 
weather. After hibernating the octopod nymphs, also sexually indis- 
tinguishable, infest rodents in April, May and June. On completing 
engorgement, which requires 3 to 10 days, the nymphs leave the host 
and molt to the adult stage during the summer. These adults normally 
do not feed until the following spring. Thus the life cycle requires a 
minimum of 2 years for completion. However, under unfavorable con- 
ditions it may be extended to 3, and rarely to 4 years, because of the 
resistance of the adult ticks to starvation. 

REARING ROOM 

The mass rearing of D. andcrsoni is conducted in a large room 
(50' x 25') specially designed to prevent the escape of ticks and to elim- 
inate, in so far as possible, places in the room in which ticks may remain 
undetected. There is but one entrance. The floor is concrete, with drains. 
The apertures surrounding all pipes passing through the floor are tick 
proofed as shown in 
longitudinal section in 
Figure 48. The walls 
are finished with ce- 
ment plaster to permit 
washing down and win- 
dows are specially de- 
signed and tightly 
screened with 18 mesh 
wire cloth. The sash 
weight boxes are tick 
tight and crevices be- 
tween the window 
frames and walls are 
packed with oakum. 
The floor and walls 
are flushed daily with 
near-boiling water 
under pressure to kill 
or remove ticks which 
may be free in the 
room. The tempera- 
ture is maintained 

automatically at 2 1 FjG 4g — showing method of tick proofing apertures 
C. or slightly above. around pipes passing through floor in tick rearing room. 



floor Leve/ 




248 Phylum Arthropoda 

Adjacent to the rearing room there is a workers' street clothes locker 
room, a shower room, a "deticking" room containing an electrically 
heated cabinet and a 3 panel full length mirror, and a work clothes locker 
room. Coming on duty the workers leave their street clothing in lockers 
in the first room and don one piece white coveralls in the fourth. On 
leaving at the close of work, the coveralls are placed in the heated cabinet 
in which the temperature is maintained between 50 C. and 65 C. for an 
hour or longer to kill any ticks that may be on the garments. The workers 
then examine their bodies for ticks before the mirror and take a shower 
bath before dressing for the street. 

REARING PROCEDURE 

Although D. andersoni can be reared through many successive genera- 
tions under laboratory conditions, stock thus carried on seems gradually 
to lose its virility. It has been found best, therefore, to start each rear- 
ing year's stock from new adult ticks collected from nature. The succes- 
sive steps involved in rearing this stock through a generation are as 
follows: (1) collection of adult ticks from nature, (2) engorging of fe- 
males, (3) oviposition and hatching of eggs, (4) engorging of larvae, 
(5) engorging of nymphs, and (6) storage of the various stages. Most 
of the essential equipment is shown in text figures. In order to have 
1,000,000 adults of the reared generation, it is necessary to collect or 
engorge approximately 5,000 females, requiring about 170 rabbits. 
These females will deposit 20,000,000 eggs yielding approximately this 
number of larvae. About 700 rabbits are required to feed these larvae, 
using about 30,000 larvae per host. Approximately 4,000,000 engorged 
larvae are obtained. These will yield about the same number of nymphs. 
Nymphal feeding requires about 2,800 rabbits using about 1,200 ticks 
per host. The approximately 1,000,000 engorged nymphs produced yield 
an almost equivalent number of adult ticks. Thus only about % of 
the larvae are brought to maturity, the principal losses being incurred in 
the feeding of the larvae and nymphs. This results in part from the 
fact that all the ticks of a given lot are not ready to feed at the same 
time. 

D. andersoni is not markedly host specific and all stages feed readily 
on domestic rabbits. In the routine feeding of D. andersoni for mass 
rearing only rabbits are used. In order that there may be as little 
moisture and waste food material as possible in the rearing cage bags 
to be later described, a scanty diet consisting solely of carrots is em- 
ployed. The larvae and nymphs will feed on almost any small mammal 
and adults can be fed on any of the larger domestic animals and even on 
guinea pigs. 

Collection of Adult Ticks. To obtain a rearing stock of D. andersoni 



Ixodidae 



249 



either unfed males and females or fully engorged females can be col- 
lected in nature in the early spring. The engorged females, ready to 
drop for egg deposition, can be secured in numbers from livestock. 

The unfed adults may be obtained by means of "flagging." A piece 
of white canton flannel about 36" square is tied to a light six foot pole 
to form a "flag" (Fig. 49). The flag is dragged over grass and low 
shrubs and ticks coming in contact with the 
cloth cling to it and are readily seen. As the 
ticks are collected on the flag they are placed 
in cork stoppered 4 dram homeopathic vials 
from which they are later transferred in lots 
of about 150 each to cardboard pill boxes for 
temporary storage. By this means an ex- 
perienced collector working in a fairly heavily 
infested area can collect 800 to 1,200 ticks a 
day. 

Engorging of Females. For engorging 
females, a capsule secured to the host by 
means of an adhesive plaster girdle is used. 
Figure 50 shows in detail the construction of 
a girdle with one feeding capsule. Two cap- 
sules may be used in the same girdle if it is 
desired to feed a larger number of ticks per 
animal. 

A circular hole slightly less than 2" in 
diameter is made toward one end of a band of 
adhesive tape B, 3" or 4" in width and long enough to encircle the 
animal and allow some overlapping. The capsule is stamped from 
20 mesh brass screening so that there is formed a circular depression 
about %" in depth and 2" in diameter surrounded by a %" rim > tne 
finished capsule having somewhat the shape of a low crowned hat. 
The crowned portion of the capsule A is then fitted into the hole 
in B so that the adhesive surface of the tape is in contact with the upper 
surface of the rim of the capsule. A slightly smaller hole is made in a 
shorter band C and the non-adhesive surface of the band is applied to 
the adhesive surface of B so that the holes in the two bands are con- 




Fig. 49. — Diagram of "flag" 
used in collecting adult ticks. 



B 



B 



C C 

Fig. 50. — Section diagram of tick feeding girdle. A, brass screen capsule; B, long 
adhesive band for girdle (fine line is adhesive surface) ; C, adhesive tape, prevents 
actual contact of rim of capsule with animal. 



V'u 



250 Phylum Arthropoda 

centric. About 40 ticks are placed in the depression of the capsule and 
the girdle applied to the animal so that the ticks may feed on a clipped 
area of the animal's belly. Two strips of %" adhesive tape, encircling the 
animal twice, reinforce the margins of the girdle. If two capsules are 
to be used the capsules are spaced so that their rims are about %" apart 
and their centers in line with the long dimension of B. A total of 80 

ticks, half males and half females, may then 

.-./" ""\ t be placed on the rabbit. A large number of 

,.:• '} females may cause the death of the host 

\, \ through exsanguination. Female engorgement 

is completed in 8 to 10 days. The rabbits are 
then sacrificed, the girdles removed and the en- 
gorged females collected. The males are 
destroyed. From 15 to 20 engorged females 
per animal are obtained from a single capsule 
girdle and 30-35 from the double. 

Oviposition and Hatching. For oviposition 
the engorged female ticks are placed individ- 
ually in glass shell vials %" in diameter and 
i%" in length, the open end being securely 
but not tightly stoppered with a plug of 
cotton (Fig. 51). The vials are then laid 
horizontally in rows on a screen tray as 
illustrated in Figure 52. The trays are placed 
almost in contact with moist sand in thermal 
cabinets operated at about 22 ° C. Under these conditions oviposition 
begins in approximately 6 days and is completed about 21 days later. 
The female dies on completion of egg laying. 

Hatching is completed and the larval ticks are ready to be fed in from 
5 to 6 weeks after the engorged females were placed in the cabinets. 
Readiness to feed is indicated by the presence of a considerable amount 
of white excretory material deposited by the larvae on the walls of the 
glass vials. 




Fig. 51. — Diagram of 
cotton stoppered ovipo- 
sition vial containing en- 
gorged female tick. 




Fig. 52.— Diagram of tray for holding oviposition vials. 



Ixodidae 



251 



Engorging of Larvae. The host rabbit is placed in 68 count white 
muslin bag 14" by 18" in size. The larvae from 4 oviposition vials are 
quickly placed in the ears and about the head of each animal and the 
bag securely tied. The bagged animals are then placed in cages sup- 
ported on frames and racks as illustrated in Figure 53. The cage frames 



52 




Fig. 53. — Diagram of cage rack and cages used in rearing of large numbers of larvae 
and nymphs. Capacity of rack is 12 cages. 



are made of %" wrought iron rods welded in place, painted with alu- 
minum, and are 15" x 17" by 18" high. Twenty- four hours later the 
animals are released and the bags and the unattached ticks clinging to 
them burned. A 10 ounce canvas bottomed bag with an 80 count muslin 
top is then placed over the supporting frame containing the cage and 
tied. The canvas bottom has a few small perforations in it to permit 
drainage into sawdust filled trays below. The frame within which the 
cage is supported serves to keep the bag from actual contact with 
the cage thus preventing holes from being gnawed in the cage bag by the 
tick host. When the feeding is done at room temperature the major 
portion of the larvae complete engorgement and drop in 5 days. Tem- 
peratures above or below shorten or lengthen the feeding period. 



252 



Phylum Arthropoda 




& 



p IG _ S4 . — Diagram of "tick picker" used in recovering fed 
immature ticks from the animal cage bags. 

removable cup C, the bottom of which is made of 
brass gauze. The screens are removed and placed 
on edge in D, the bottom of which is open, where 
they are cleaned by washing with water from a 
hose. The volume of larvae collected is meas- 
ured in cubic centimeters, i cc. equaling ap- 
proximately 700 ticks. An average of about 
5,600 ticks per rabbit is obtained in routine feed- 
ings. The engorged larvae in quantities of 10 cc. 
each are placed in glass cylinders i%" in di- 
ameter and 4" in length, their escape being pre- 
vented by pieces of muslin securely taped or tied 
over the ends (Fig. 55). After being held from 
4 to 5 weeks in thermal cabinets at 22 ° C. and 
relative humidity about 50%, the larvae will 
have molted to nymphs and the latter be ready to 
feed. 

Engorging of Nymphs. The procedure fol- 



A "tick picker" 
(Fig. 54) is used 
in recovering fed 
immature ticks 
from the bags in 
which they are 
contained after 
dropping from the 
animals. It is 
made of galva- 
nized sheet metal 
with dimensions 
as illustrated. 
The bags are re- 
moved from the 
cages, turned in- 
side out and 
shaken in the hop- 
per. Removable 
screen A, 6 mesh, 
and B, 14 mesh, 
retain the trash 
while the engorged 
larvae fall through 
and are caught in 





Fig. 55. — Diagram of 
pyrex cylinder in which 
fed larvae are held for 
molting. 



Ixodidae 253 

lowed in feeding nymphs is similar to that for feeding larvae. One of 
the i 3 //' x 4" cylinders is opened and, with the help of an assistant, the 
contents are equally distributed among 6 animals in infesting bags. 
Considerable experience and dexterity is required in completing the 
process without the escape of ticks. The rabbits are then treated as 
in larval feeding. Engorgement is accomplished in about 8 days and 
the tick picker is also used in collecting the fed ticks. The contents 
of the cage bags are shaken in the hopper and a stream of water from a 
hose is directed onto the canvas bottom of the bag to wash the remain- 
ing ticks and detritus into the picker. The soluble wastes are washed 
through while the ticks and non-soluble waste of the same size are 
retained on screen B. The coarse material is retained by screen A. 
Cup C is not used while the bags are being cleaned. After all the bags 
are cleaned, water is directed into the picker and the material in it 
thoroughly washed. Screen A is removed and its retained material 
discarded. Screen B is set on edge and the material on it washed down 
into the cup which is now in place. The cup is removed and the con- 
tents are slowly centrifuged in a screen container to remove the excess 
water. This is accomplished in a converted cream separator. Drying is 
then completed by placing the material in a current of warm air sup- 
plied by an electric hair dryer. Final cleaning is accomplished by 
sorting the remaining debris from the ticks, after which the latter are 
counted by measuring their volume in cubic centimeters, 35 ticks being 
the equivalent of 1 cc. An average of about 400 ticks per animal is 
ordinarily obtained. The nymphs are placed in cardboard pill boxes, 
2" in diameter and %" in depth, about 200 per box, and the lot number, 
date and other necessary data stamped on the covers. 

Molting of nymphs occurs within a rather wide range of temperature 
and humidity. Ordinarily nymphs are held at 22 C. and relative hu- 
midity of 40-80%. Under these conditions molting takes place in 
about 2 1 days. Transformation to the adult stage is accelerated by a 
low relative humidity, while a high humidity tends to retard the process. 

STORAGE OF TICKS 

In the course of routine or experimental rearing, situations occasionally 
arise when it becomes necessary to delay development or postpone the 
feeding of ticks. The conditions required for minimum mortality dur- 
ing storage are dependent on the stage of tick being dealt with, the 
principal factors being its ability to withstand desiccation and starvation. 
Adult ticks are better able to survive long periods of fasting than the 
immature stages, the nymphs being more resistant than larvae. None of 
the stages are markedly resistant to desiccation. 

Adults. Unfed adults of D. andersoni collected in nature or recently 




8 



254 Phylum Arthropoda 

molted from nymphs may be stored a year or more without excessive 
mortality by placing them in pyrex cylinders or "longevity tubes." These 
tubes are 8" in length by i%" in diameter. One end is plugged with well 
packed earth or with Plaster of Paris for a distance of 2", and a few 

coarse wood shavings or dried leaves are added 
so that the ticks are not forced into direct 
contact with the plug (Fig. 56). From 300 
to 400 ticks are placed in the tube and the 
latter is closed by a piece of muslin tied over 
the open end. The tubes are placed upright 
inside 4" galvanized metal cylinders set in the 
ground to their full length. The plugged end 
of the pyrex cylinder must be in close contact 
with moist soil. Moisture is supplied to the 
surrounding soil as necessary. The ticks are 
thus free to seek favorable humidity conditions. 
The "tick yard" in which the tubes are placed 
must be constructed or located so that the 
direct sun rays will not fall on the tubes. 
During the winter the tubes may either be left 
in the ground or stored indoors at 6° C. The 
ticks will survive outdoor conditions unless 
subjected to unusually abrupt transitions of 
temperature, particularly from cold to heat. 

For periods of at least 6 months adult ticks 
may be stored in pill boxes at 6° C. with the 
relative humidity maintained at about 80%. 

Engorged female ticks may be stored to delay 
oviposition for periods up to 4 months in card- 
board pill boxes at temperature and humidity 
conditions as stated above. 

Immature Stages. Unfed larvae may be 
stored for periods up to 2 months under the 
conditions just stated. 

Unfed nymphs may be kept successfully for 
periods up to 6 months in longevity tubes in the 
tick yard in the manner described for adult 
ticks. However, equally satisfactory results will ordinarily be obtained 
by storing the tubes upright on a tray of moist sand in a thermal cabinet 
operated at 6° C. 

Fed larvae and fed nymphs are susceptible to desiccation at the lower 
ranges of humidity and to molds at the higher ranges. Therefore, it is 
more desirable to store immature ticks as unfed nymphs. 



* \% » 

Fig. 56. — Diagram of 
pyrex "longevity tube," 
used for storing unfed 
adult or nymphal ticks 
for long periods. 



Ixodidae 



255 



REARING SMALL LOTS OF TICKS 

In some respects the rearing procedure and equipment described above, 
while adequate for the rearing of D. andersoni in large quantities, are 
not well adapted for the requirements of a small laboratory where a 
lesser number of ticks is required. 

Small numbers of ticks may be confined in cotton stoppered or muslin 
capped vials to permit ready observation. In the absence of thermal 



JF~l 



D D 

Fig. 57. — Section of tick feeding girdle for experimental purposes. A, threaded and 
flanged ring; B, cover; C, long adhesive band for girdle; D, short adhesive band 
covering toothed flange. 



cabinets ticks may be kept at room temperature in glass vials or card- 
board pill boxes almost in contact with moist sand. In cases where con- 
trolled humidity conditions are required the pill boxes or vials may be 
kept in ordinary desiccating jars containing solutions of salts that will 
provide the desired relative humidity. 

The tick feeding girdle shown in Figure 50, while simple and useful 
for routine tick feeding, is not adapted for experimental feeding of small 
groups of ticks where close observation and easy manipulation of the 
feeding ticks are 
desired. Once the 
adhesive tape is in 
place its partial 
removal in order to 
remove or replace 
ticks causes skin 
irritation and any 
active and un- 
attached ticks may 
escape. Therefore, 
a tin capsule made 
from the threaded 
end ring and cover 
used in cardboard 
mailing tubes is 
substituted for the 

brass gauze capsule used in mass feeding. A section diagram is shown 
in Figure 57. For complete details the reader is referred to Public 
Health Reports 1933, pp. 1081-82. 




Fig. 58. — Diagram of a cage designed for rearing ticks on 
small animals. 



256 Phylum Art hr op oda 

A diagram of a small cage suitable for rearing ticks on guinea pigs, 
white rats, mice, chipmunks and other small rodents, is shown in Figure 
58. The wire frame, for keeping the sack in which the cage is enclosed 
from being gnawed by the caged animal, is soldered to the cage. The 
cage is placed in a cloth bag, the animal introduced into the cage and the 
ticks to be fed are placed on the animal. The bag is closely tied and 
the unit placed over a tray of sawdust. No further attention, except for 
feeding the animal, is necessary until the ticks have fed and dropped. 
The fed ticks are recovered from the bag with forceps, and after having 
been immersed in water to remove the animal urine with which they are 
likely to have been in contact in the bag, are dried and put away in the 
usual manner for molting. 



Family hydrachnidae 



PARASITIC WATER MITES 

John H. Welsh, Harvard University 

MOST parasites may be maintained in the laboratory when it is 
possible to maintain their hosts, and the modifications which many 
of them exhibit never cease to interest students. Forms which show 
structural adaptations are numerous but forms which show clear-cut 
modifications in behavior are few. It would appear to be of interest to 
know that a form which is readily obtained and maintained in the 
laboratory does show a striking modification in behavior due to its para- 
sitic existence. 

Unionicola ypsilophora is a common parasitic water mite which is found 
in several species of Anodonta. It lives on the gills and all of the develop- 
mental stages may be found in or on the gills of a form such as Anodonta 
cataracta. A supply of these mussels may be kept for months in the 
laboratory in running water and the mites may therefore be available at 
any time, even though collecting conditions are unfavorable. The mites, 
after removal from the mussels, may also be maintained in finger bowls 
of water for several weeks and even months if the temperature is around 
40-50 F. They require little attention although they may be fed an 
occasional small bit of mussel gill. 

After these mites have been removed from their host they show a well- 
marked positive phototropism. This at first seems anomalous as it is 
difficult to understand why they are not attracted from the mussel, by 
way of the siphons, when a bright beam of light penetrates the mantle 
cavity as frequently happens at mid-day. The reason they do not leave 
the host is seen when the following experiment is performed. If water 
from the mantle cavity of the host or a filtered water extract of the gills 



Tetranychidae 257 

is added to water containing the mites they immediately show a negative 
response to light and maintain this state for a length of time depending 
somewhat on the concentration of host substance. 

It may be further shown that this reversal in phototropism is due to 
specific host material for if water from the mantle cavity or extract of 
gills of mussels on which this species of mite is not parasitic is used there 
is no effect on the phototropism of the mites and they remain positive to 
light. 

The parasitic Unionicola may be identified by reference to papers by 
Wolcott (1899, 1905) and Marshall (1933). 

Reference 
For the culture of Hydracarina see p. 136. 

Bibliography 

Marshall, Ruth. 1933. Preliminary list of the Hydracarina of Wisconsin, 

Part III. Trans. Wis. Acad. Sci. Arts and Letters. 28:37. 
Welsh, J. H. 1930. Reversal of phototropism in a parasitic water mite. Biol. 

Bull. 59:165. 
1931- Specific influence of the host on the light responses of parasitic water 

mites. Ibid. 61:497. 

1932. A laboratory experiment in animal behavior. Science 75:591. 



Wolcott, R. H. 1899. On the North American species of the genus Atax (Fabr.) 

Bruz. Trans. Amer. Micr. Soc. 20:193. 
1905. A review of the genera of the water mites. Ibid. 26:161. 



Family tetranychidae 

J 

BREEDING OF NEOTETRANYCHUS BUXI, A MITE ON 

BOXWOOD 

Donald T. Ries, Ithaca, New York 

THESE mites may be found on the underside of the leaves of Box- 
wood bushes in some localities. 
Small cuttings of boxwood about an inch in length were placed 
in small pots of earth. The pots of cuttings were then placed in 
large flats of peat moss in order to facilitate handling and also to keep 
them from drying out. Since the mites seemed unable to negotiate the 
distance between the pots over the moss there w r as no need of covering 
the individual plants. By this means mites were reared through nine 
generations. 

Each female was allowed to deposit one or two eggs on a plant and 
every 24 hours was removed to another plant by means of a fine camel's 
hair brush. In no case were more than two nymphs allowed to live on 
any one plant. 



258 Phylum Art hr op oda 

Newly hatched nymphs are active, moving about from one surface of 
the leaf to the other. From 1 to 3 days after hatching they become quies- 
cent and cast their skins. The second instar nymphs feed actively on 
the upper and lower surfaces of the leaves. The entire life cycle is 
completed in 18 to 21 days, averaging 18% during the summer months 
when temperatures are fairly high and humidity low. Females have lived 
as long as 6 weeks during the summer, while others have laid their full 
complement of eggs and died within 2 weeks after emerging. 

Copulation takes place soon after the female emerges from the third 
instar skin. In several instances the male was observed standing over 
a quiescent third instar mite even before she had begun exit from the 
skin. Later work showed that fertilization may take place during this 
time. Oviposition usually takes place % to % of an hour after fertiliza- 
tion. One egg is deposited at a time and from 1 to 5 may be laid during 
24 hours. 

Another method for rearing mites on plants having large leaves was 
to cut small doughnuts of felt. Each of these was fastened to a leaf 
with waterproof glue and covered with glass or cellophane which was 
held in place by a paper clip. Fiber circles, such as those used in making 
microscope slides, were also used with excellent results. A mite working 
on Rhododendron was reared through two seasons under these conditions. 

Class Myriapoda y Order diplopoda 

EURYURUS ERYTHROPYGUS* 

THIS millipede is abundant in the heartwood of much decayed logs, 
in moist and more decayed sapwood, and on the soil under decaying 
wood if rather moist conditions prevail. 

Specimens were collected and placed in glass receptacles approximately 
5% inches in diameter and about 3 inches deep. These were half-filled 
with small, broken pieces of moist and much decayed sapwood from 
a rotten log, together with a little humus. This material was examined 
carefully for contaminating forms, such as other millipedes, centipedes, 
mites, earthworms, eggs, insects, etc. A layer of vaseline was spread 
around the rims of the receptacles and glass covers placed over them, 
thus insuring very little if any evaporation. However, a few drops of 
water were added occasionally. Moisture and other conditions were kept 
as natural as possible. Some of the receptacles were opened for observa- 
tions every day and fresh air entered at these times, but the animals 
seemed to thrive in receptacles which were not opened so often. 

* Abstracted from an article in the Ohio J. Sci. 27:25, 1927, by Hugh H. Miley, Ohio 

State University. 



Thysanura 2 59 

From adults collected in the fall a number reared in the laboratory- 
survived during the following summer and most of the fall. Adult males 
and females, when observed copulating, and in some cases males and 
females not pairing, were isolated in separate receptacles. In most cases 
the females laid eggs in cavities made by themselves a short distance 
below the surface of the soil. These were permitted to hatch in the same 
receptacle with the adults. As soon as the larvae started emerging from 
the eggs, a number of them were placed in petri dishes in order to observe 
their habits more accurately. 

M. E. D. 

Order symphyla 

REARING OF SCUTIGERELLA IMMACULATA* 

THIS garden centipede will live on a wide range of vegetable matter 
and probably on decaying vegetable matter within the soil. 

Eggs are laid during the early part of the summer in the subsoil run- 
ways of the creatures, usually in clusters of 4 to 20. At room tempera- 
tures the eggs hatch in about 14 days. When the larvae first hatch they 
have 6 pairs of legs. One pair is added at the first molt, which occurs in 
1 to 4 days, and a pair is added at each successive molt, occurring 7 to 
14 days apart, until each individual has 12 pairs. 

Rearing of these creatures is done in petri dishes on a thin layer of 
soil and in stender dishes into which is poured a "muck-plate" made 
by mixing 10 parts plaster of Paris and 3 parts of finely ground muck. 
This mixture makes it easy to find the white eggs and to observe the 
whitish creatures as they move about. Lettuce leaves are placed on the 
surface of the plates for food and as a hiding place. Rearing records 
show that individuals may live n or 12 months. 

M. E. D. 

Class Insecta, Order thysanura 

REARING OF THYSANURA 

G.J. Spencer, University of British Columbia 

Campodeidae. These are difficult to rear because they are extremely 
sensitive to changes in moisture. A large volume of greenhouse potting 
soil or compost in a big tin box with a tight-fitting lid, will keep a colony 
going for a few weeks. 

Machilidae. These are also unsatisfactory to rear. The coastal species 

♦Abstracted from an article in /. Econ. Ent. 21:357, 1928, by George A. Filinger, 
Ohio Agricultural Experiment Station. 



260 Phylum Arthropoda 

in this province may be kept alive for some time in a mass of damp 
leaves from the forest floor; I have not succeeded in getting them to 
breed in captivity. Machilis maritima covers much territory on the rocky 
seashore; it dies quickly in confinement. Of the four (unnamed) species 
in the dry interior of British Columbia, only one may be kept in cages. 
It frequents deep moss from under timber in gullies. A square foot of 
this moss in a large tin will keep a small colony alive for several weeks. 

Lepismidae. Thermobia domestica is the easiest of all to rear. Any 
long box or laboratory drawer will do if it has a close-fitting lid and has 
one end against a hot radiator, to ensure a temperature, at one end, of 
90 to ioo° F., and a gradient dropping to 8o° F. at the other. Eggs are 
laid at about 80 ° to 85° F. 

I have kept one colony going for ten years in a large, 27-drawer incu- 
bator. The shell of the incubator is double- walled with celotex (corn 
stalk board), with a 4-inch air space between walls perforated at in- 
tervals with 1 -inch holes. The back has a baffle-plate of thin asbestos 
board heated with a battery of six large carbon lamps, two of which dip 
into a large pan of water. The ends of all drawers touch the asbestos 
board at the back, whence they get the necessary heat. The drawers are 
roughly 27 x 10 x 7 inches, open on top with a 2 -inch wide strip of cellu- 
loid cemented on the edge all round and overhanging inwards. 

Lepismids cannot walk on smooth surfaces, and so cannot escape from 
the open drawers. On the floor of each drawer is a dissecting pan or 
other shallow tin tray i-inch deep, full of sand which is kept always damp. 
At 2-inch intervals on the floor are flat 2 -inch squares of cotton-batting 
blackened with India ink and dried. 

Eggs are deposited in the cotton wool at various distances from the 
heated rear end, and the nymphs find adequate shelter under it until the 
third instar when they venture further afield. Food consists of whole 
wheat meal or plain flour; at intervals of 2 weeks I put in a teaspoonful of 
very lean beef or veal, thoroughly dried on a radiator and pulverized. 
This meat powder is a great attractant. All food must be dry. 

Watch out for overcrowding, or disease will wipe out the whole colony. 
Five hundred individuals can live in a drawer of the dimensions above ; 
two or three hundred is better. There is one brood per year. Breeding 
occurs at irregular intervals. 

Lepisma saccharina colonies have been kept in the incubator for eight 
years. They require less heat than T. domestica and more moisture. In 
addition to the tin tray of damp earth or sand, I use a heap of small 
squares of shingle separated from each other alternately, by a match. 
This gives narrow crevices in which the colony spends most of its time, 
especially the young nymphs. On top of the cedar shingles is a small 
saucer of raw earthenware which is filled with water every week, thus 



Lepismatidae 



261 



keeping all the shingles damp. Food, as for T. domestka. I have also 
used printers' starch at intervals. There is one brood per year. 

Ctenolcpisma quadriseriata was brought from Ontario and kept alive 
in Vancouver for only 1 % years. One brood was produced and then the 
whole colony died with the exception of a lone male which lived 2 years. 
This species can stand freezing, requires a much lower temperature than 
the others, and thrives best on slabs of alder bark. It will eat farinaceous 
foods but feeds in nature, I think, on algae on roots and on trees. 

The incubator and all rearing tins are kept in a small laboratory from 
which the heat goes off every night at 7:30, coming on again at 7:30 in 
the morning. In addition, the plug of the incubator is connected at 8 a.m. 
every day and pulled out at 6 p.m. By this means a change of tempera- 
ture is ensured, averaging week by week for seven winter months, a dif- 
ference of 30 F. between night and day. In summer the incubator is 
set so as to ensure a maximum mid-day temperature of ioo° F. 



Family 



LEPISMATIDAE 



METHODS OF REARING LEPISMATIDS 

J. Alfred Adams, Iowa State College 

The Firebrat, Thermobia Domestka 

THIS is the largest commonly available member of the Apterygota. 
Its possibilities as a laboratory animal have not been generally 
appreciated. 

Firebrats may be reared in great numbers in air-conditioned cabinets 
such as those described by Brindley and Richardson (1931). The con- 
ditions around the culture dishes are: 37 C, 75% relative humidity, 
gently moving air, and light of twilight intensity or darkness. 

The culture dishes (Fig. 59) 
are of glass, 20 cm. in diameter, 
with vertical sides, 8 cm. in 
height. They contain 20 or 
more paper strips 4 cm. in 
width and 30 cm. in length 
folded transversely, the folds 
occurring about every 2 cm. 
and alternating in direction so 
that the folded strip resembles the collapsible side of a bellows. These 
strips are stood on edge in the dish. Cotton batting receives the eggs. 

Firebrats thrive when they have continuous access to separate heaps of 
rolled oats, dried ground lean beef, cane sugar, dried brewer's yeast, and 
common salt. Rolled oats may be used as the sole food. If the tightly 



Folded 
Paper 

Cottor 




j Watering 
Tube 



Food 



Fig. 59.— Sketch of apparatus for general use 
in rearing the firebrat. The tubes are usually 
omitted. 



262 



Phylum Arthropoda 



Waterinq 
Tube 




otton 
lower Pots 



J-3 



-Food 



Fig. 60. — Cross section of rearing appara- 
tus for the firebrat for use in ovipositional 

studies. 



closed rearing cabinet has, under its fan, a large surface of brine con- 
taining an excess of common salt, the humidity will be held near the 
desired percentage and watering will be unnecessary. At humidities 
lower than 70% a slender test tube of water, tightly plugged with cotton 
batting, may be inverted on a piece of cardboard in the dish so that the 
insects may be able to rest and moisten themselves on the damp surface. 
Such a cage accommodates one to two hundred adult firebrats. A few 

hours' attention a month suffice 
for their care. 

Where a careful check on food, 
population, or oviposition is desired 
another type of apparatus is rec- 
ommended. It consists (Fig. 60) 
of a culture dish, similar to the 
above, into which are inverted 
three clay flower-pots graded in 
size so that the largest covers the 
medium-sized one, which in turn covers the smallest. In order that 
the insects may run under them the edges are supported on a thin wedge 
of wood. Cotton batting to receive the eggs is placed between the 
middle and outer pot. 

To segregate new generations the cotton, bearing eggs, may be trans- 
ferred to new rearing quarters. Watering is not advised. The tiny 
nymphs soon leave the cotton for the paper. At this temperature they 
become sexually mature in about 3 months and gravimetrically mature 
in about 6 months. 

Firebrats must not be grasped directly; they may be passed singly 
without injury from one glass dish to another. Two species of greg- 
arines, described by the author, are likely to appear in the cultures. 
To get gregarine-free cultures it is necessary to obtain freshly laid eggs, 
spore-free, and transfer them to sterilized, lidded, culture dishes. A ring 
of vaseline around the outside prevents the entry of book-lice. 

The Silverfish, Lepisma Sacchar'ma 

The author has not reared silverfish extensively. Sweetman (1934) 
states that 28 C. and 90% R. H. are satisfactory conditions for this 
species. 

It has been found convenient to keep silverfish in apparatus similar 
to that used for the firebrat but at a constant temperature near 28 C, 
a relative humidity near or above 75%, and with provision for the insects 
to have access to a small moist wick. The culture dish should be deeper 
than that for the firebrat or nearly closed and kept in semi-darkness. 
The papers should be more closely packed. 



Collembola 263 

References 

For the culture of Thermobia domestica see also p. 260. 
For the culture of Lepisma saccharina see also p. 260. 
For the culture of Ctenolcpisma quadriseriata see p. 261. 

Bibliography 

Adams, J. A. 1933. J. N. Y. Ent. Soc. 41:557- 

Brlndley, T. A., and Richardson, C. H. 1931. Iowa State College J. of Sci. 5:211. 

Spencer, G. J. 1930. Canad. Ent. 62:1. 

Sweetman, H. L. 1934. Bull. Brooklyn Ent. Soc. 29:156. 



Order collembola 

REARING OF COLLEMBOLA 

G. J. Spencer, University of British Columbia 

MANY species may be kept alive and breeding for long periods, 
in tightly lidded tobacco cans supplied with the humus or leaf 
mold on which they were collected. Rotting potato is a good medium for 
some species. I have kept a colony flourishing on this material for over 
two years. Another good container is a tin covered with a piece of glass, 
covered in turn with a piece of dark cardboard. This permits examina- 
tion of the colony without removing the glass. 

Two points are of importance in rearing Collembola: Keep the ma- 
terial on which they are feeding damp ; secondly, do not uncover them 
often or let wind currents disturb them and dry out their culture medium. 
Flat stones placed in the tins will serve for cover. 

REMARKS ON COLLEMBOLA* 

COLLEMBOLA are all extremely sensitive to any lack of humidity 
in their surroundings. The only way to keep them alive in captivity 
for any length of time is to put in the vial some source of moisture such 
as wet, rotten wood or damp filter paper. 

The white or yellow spherical eggs are laid singly or in masses under 
bark, among dead leaves, and in many other damp situations. Ovi- 
position apparently takes place only in the dark. Several species lay 
eggs freely in captivity. Incubation at room temperature takes from 
10 to 35 days, according to the species. In captivity, Achorutes socialis 
and some other species lay only in the spring, while A. humi and Neanura 
muscorum oviposit late in the fall. The eggs of the last named species 
require 35 days to hatch at an average temperature of 6o° F. This is a 

♦Abstracted from two articles in the Canad. Ent. 51:73, 1919, and 56:99, 1924, by 
Charles Macnamara, Arnprior, Ontario. 



264 Phylum Arthropoda 

remarkably long period compared with the 10 or 12 days required by 
the eggs of Achorutes socialis under the same conditions; and in the 
insect's natural environment incubation would doubtless have been even 
longer. 

Springtails feed largely on the vegetable molds and minute algae which 
flourish in such situations. Fungi are a favorite food of many species 
and both spores and pieces of mycelium are often found in their stomachs. 
Liquid food also attracts these chewers, and in the spring several species, 
particularly of the genus Achorutes, may be seen in large numbers feed- 
ing on the sweet sap exuding from freshly cut maple stumps. 

Species that live on the surface of pools and streams, such as Isotoma 
palustris and Sminthurus aquaticus, often pick up diatoms and desmids, 
and in the spring feed largely on the pollen which the conifers lavish 
upon the wind. 

Our pollen-eating species go direct to the flowers of various plants. 
In Switzerland, Handschin says, Sminthurus hortensis is always found in 
the blossoms of Ranunculus glacialis, and in this country the same 
species is common on dandelion blossoms. In America Achorutes 
armatus sometimes makes a nuisance of itself in beds of cultivated mush- 
rooms, and a few other species have some bad marks against them. 
Sminthurus hortensis feeds upon beans, beets, cabbage, cantaloupes, car- 
rots, clover, corn, cucumber, kale, lettuce, mangolds, onions, peas, po- 
tatoes, radishes, rye, spinach, squash, tobacco, tomatoes, turnips, violets, 
watermelons, wheat, wild cucumber. Its depredations make it the most 
widely known of the springtails. 

The species definitely known to be carnivorous are few, but further 
study of Collembolan life histories will probably reveal others. An un- 
doubted flesh-feeder is Anurida maritima, an inhabitant of the seashore. 
Folsom says the insect's principal food is the soft tissues of the mollusk, 
Littorina littorea, as well as dead fish cast up on the shore. Imms slightly 
extends the diet to include an occasional desmid or other green alga. 
Motter's courageous study of the fauna of the grave brought to light 
another carnivorous springtail in Isotoma sepulcralis, which was abun- 
dant with a large percentage of the 150 corpses examined. 

Anurida maritima and Isotoma sepulcralis feed on dead fish. Two 
other carnivorous species, Friesea sublimis and Isotoma macnamarai, 
are raptorial and devour living prey. The food habits of /. macnamarai 
were mostly observed in vials where the specimens were kept with small 
pieces of damp, rotten wood to provide the moisture so necessary to all 
these thin-skinned insects. Cannibalism in captivity was noted in these 
two last named species and the two species were found to eat each 
other. 

Arthropleona would not eat at all in captivity. m. e. d. 



Collcmbola 265 

A METHOD FOR REARING MUSHROOM INSECTS AND 

MITES* 

WHILE conducting studies on the biology and control of insects 
and mites affecting cultivated mushrooms, it was found necessary 
to rear large numbers of these pests. Various rearing methods were 
tried, including the use of soil in salve boxes, manure in vials, etc., but 
none was found more satisfactory than the following: 

The insects and mites, in as pure culture as possible, were introduced 
in small numbers into fresh 1 -quart bottles of commercial mushroom 
spawn, and allowed to breed and develop. This spawn is made of 
chopped straw and manure thoroughly mixed, sterilized in an autoclave, 
and later inoculated with mushroom mycelium, grown from spores. With 
incubation at room temperature, the mycelium penetrates to the bottom 
of the bottles and completely fills the interstices of the medium. This 
spawn is thus a pure culture and is uncontaminated with molds. Tight 
cotton plugs are used in the bottle mouths. 

The mushroom mycelium furnishes an excellent food for these mush- 
room pests, and they gradually eat it out, leaving the original straw- 
manure medium. Feeding begins at the top of the spawn, and as it 
progresses, the portion destroyed is sharply differentiated from the un- 
eaten part. Eggs are laid and the stages develop next to the glass, 
where they are easily observed with a binocular microscope. In studying 
the development of any particular eggs or groups of other stages, a 
circle is drawn around them on the glass with a wax pencil. They are 
thus readily referred to. 

It is very important that the flies and springtails to be reared should 
be free from mites, the hypopi of which are often carried on their bodies. 
Otherwise the mites may breed so rapidly as to destroy the mycelium 
and perhaps starve the insects. Contamination with molds should also 
be prevented as much as possible. 

These bottles of insect colonies should be kept in a somewhat humid 
atmosphere. After the cultures have been developing awhile the insect 
excreta, as well as bacteria entering with the insects, will usually make 
the medium moist enough so that further additions of moisture are un- 
necessary. Most of the rearing experiments were carried on at tempera- 
tures between 50 and 65 F. as these represent the usual temperature 
limits of the bearing mushroom houses. 

The following insects have been successfully reared in spawn bottles 
of this kind: 

Collcmbola. These tiny insects are sometimes rather difficult to rear 

*Reprinted, with slight changes, from an article in Ent. News 40:222, 1929, by C. A. 
Thomas, Pennsylvania State College. 



266 Phylum Art hr op oda 

under experimental conditions, because of their susceptibility to dessica- 
tion, but in these spawn bottles they thrive remarkably well, and large 
numbers of several species have been reared. 

Achorutes armatus thrives in the spawn bottles, but is quite susceptible 
to drying. Proisotoma (Isotoma) minuta, collected in the soil, usually 
breeds rapidly in spawn bottles. Lepidocyrtus cyaneus breeds very 
readily in spawn bottles. It can withstand somewhat dryer conditions 
than can some of the other springtails. L. albus breeds readily in spawn 
bottles. Sminthurus caecus breeds slowly in spawn bottles. 

Diptera. Many generations of Sciara coprophila and Neosciara pauci- 
seta, of the family Sciaridae, have been reared in these bottles. Three 
generations of a parasite of these flies, a species of the hymenopteran 
Calliceras (Ceraphron) near C. ampla Ashmead, were also reared. Small 
orange-colored flies of the family Cecidomyidae, the larvae of which 
were collected in mushroom caps and mushroom beds, have been reared 
successfully. 

Acarina. (Mites). There is usually no trouble in rearing tyrogly- 
phids and numerous other mites in the spawn bottles. In fact it is often 
difficult to obtain pure cultures of other mushroom insects because of 
infestation by these pests. The chief species feeding on mushrooms and 
mycelium are tyroglyphids, chiefly Tyroglyphus linteri, another Tyro- 
glyphus species, and sometimes a species of Histiostome which feeds on 
the decaying tissues of injured or diseased mushrooms. All of the 
above mites have been reared through many generations in the spawn 
bottles, and all stages, including the very interesting hypopi of the tyro- 
glyphids, have been found in the spawn. Abundance of moisture is no 
deterrent to the development of these mites as they may often be found 
partly submerged in the moist surface of the spawn medium. 

It is probable that other fungus insects could be reared in these 
bottles provided the moisture and other factors were regulated to suit 
the species. In order to make smaller cultures the spawn may be removed 
to smaller bottles or vials. However, it is necessary to avoid contamina- 
tion with molds and mites during this process. 

M. E. D. 

Order ephemeroptera 

REARING MAYFLIES FROM EGG TO ADULT 

Helen E. Murphy, Phoenix, New York 

SINCE no way has yet been found of inducing mayflies to mate in 
captivity, it is necessary to capture a female carrying her eggs. 
Late in the afternoon of a quiet, sunny day in late spring or in summer, 



Ephemeroptera 267 

it is usually possible to locate numbers of males and females dancing up 
and down in their mating flight near the bank of a pond, lake, or stream. 
One by one the females leave the throng and fly over the water, here and 
there dropping with their eggs to the surface. With a net capture one 
of these females before she reaches the water. Holding her gently by 
the wings wash the eggs into a culture dish and transfer them to the 
laboratory.* 

Mayflies of the genus Baetis do not carry their eggs in a protruding 
mass, but crawl into the water and lay them in a flat layer on a sub- 
merged stone. These may be removed with a scalpel as soon as laid and 
carried to the laboratory in water. 

With the aid of a lens and a scalpel, separate individual eggs and trans- 
fer them to culture dishes by means of a medicinal pipette. Covered 
culture dishes 75 mm. x 20 mm. prove very satisfactory. Fill each dish 
% full with tap water seasoned for 24 hours in an open container at room 
temperature. With a pipette change half of the water every three days. 
Avoid sudden temperature changes. 

Diatoms and desmids freshly scraped from stones from a stream 
furnish excellent food. One drop of thick culture every three days is 
sufficient for a very young nymph. Twice that amount is required later. 
Diet may be varied by the addition of a small fragment of Spirogyra. 

When longitudinal venation is clearly evident in the wing pads prepare 
for emergence. Cut a piece of No. 12 or No. 16 wire cloth 12 cm. x 30 cm. 
Remove the glass cover and roll the wire to fit tightly inside the glass 
rim. Pinch the top together and fold twice, making a closed seam. This 
cage allows the specimen to crawl from the water and to hang in the 
air waiting for the final molt. 

In Baetis vagans there are 27 instars and 6 months are required for 
the development of the summer brood. 

Bibliography 

Murphy, Helen E. 1922. Notes on the biology of some of our North American 

mayflies. Lloyd Library Bull. No. 22. 
Smith, O. R. 1935. Eggs and egg-laying habits of North American mayflies. In 

Needham, Traver, and Hsu: The Biology of Mayflies, p. 67. 

* Editor's Note: Artificial fertilization of certain mayflies (Hexagenia, Ephemera, etc. ) 
is easily effected by mixing freshly liberated eggs and sperms in a watch glass of lake water. 
A gravid female taken in her ovipositing flight will shed all her eggs with the greatest 
readiness on such stimulation as snipping off her head or subjecting her to tobacco fumes. 
Eggs of a subimago are sometimes mature and ready for fertilization. J. G. N. 



268 



Phylum Arthropoda 
Order odonata 



CULTURE METHODS FOR THE DAMSELFLY, ISCHNURA 

VERTICALIS 

Evelyn George Grieve, Cornell University 

REARING work was begun with the full-grown nymphs which were 
. collected early in the spring from a small, grassy fish pond. From 
the first generation, which were the stock adults, eggs were obtained for 
life history studies. As soon as the nymphs hatched they were isolated 
and reared in separate containers for the entire life span. 

.. ,... c ....,,,, JiV , n Cages. The type of cage 

..:■•-* "'. '~> ""'"" N " ; If** j (Fig. 61) used for breeding 
^ f ->^M' :: .$£': /-'M stock was a fairly large 

aquarium covered with a 
screen cage, fitting the 
aquarium closely at the 
sides, and allowing room 
above for the adults to fly, 
mate, and capture their 
prey. Aquatic plants, prin- 
cipally Eleocharis palustris, 
were kept growing in one 
corner of the aquarium; up 
these the nymphs could 
crawl to transform. When 
the females were ready to 
oviposit, one or two of the 
flexible stems of E. palustris 
were bent down into the 
water where they were held 
by the surface film. The 
females preferred to 
oviposit in these floating 
stems, and each day eggs 
could be removed and dated for the study of embryonic development. 

In order to keep records of all individuals, and also because the species 
is cannibalistic, separate containers were used for all reared specimens. 
The most satisfactory type for the young nymphs was a small boat 
(Fig. 62 ) , floating with others in a large pan of water. The frame of the 
boat is of balsa wood, which is very light and buoyant, and the "hold" of 
the vessel is made of silk. Although the illustration of the boat is about 
natural size, the silk was of finer mesh than that shown, namely about 




Breeding cage for Ischnura verticalis. 



Odonata 



269 




Fig. 62. — "Boat" for rearing 
nymphs of Ischnura verticalis. 
(About natural size.) 



144 threads to the inch, which is fine enough to retain the smallest nymph 
and its food organisms, and yet permits circulation of water. The silk 
was fastened to the frame with paraffin, 
and the frame was covered with a film of 
paraffin to reduce "water-logging." The 
little "sail" was a paper tag, attached by 
a special pin, and was used to record dates 
of molting and other data. 

The advantage of such containers, over 
Syracuse watch glasses for instance, is that 
the nymph has all the benefits of a large 
vessel of water, which does not stagnate 
so quickly, and is less subject to tem- 
perature fluctuations. 

When the nymphs were nearly full grown 
(i.e., in the last two or three instars) and 
were inclined to crawl out of the boats, 
they were transferred to glass tumblers, 
provided with a strip of wire screen to 
serve as a perch. Previous to transformation, the glasses were covered 
with cheesecloth fastened down with an elastic band, thus making con- 
venient emergence chambers in which to observe molting. 

The young imagos were then transferred to individual aquarium-cages, 
similar to, but smaller than, the stock cages (Fig. 61 ) , one pair to a cage, 
so that data might be kept on mating, oviposition, color changes, etc. 

Feeding. Both nymphs and adults are strictly carnivorous, and for any 
large scale rearing it is necessary to maintain colonies of food organisms. 
The adults have a preference for dipterous insect prey. One method 
of supply was to stock the breeding tank with quantities of full grown 
blood worms and mosquito larvae and pupae, and to allow the damsel- 
flies to feed on the emerging midges and mosquitos. Later in the season 
when this supply failed it was necessary to collect midges in the woods 
with a net, morning and evening, and liberate them in the cages. 

For the very young nymphs, Paramecium cultures were tried and 
abandoned. Very minute chironomid larvae were fed to the next group 
of damselfly nymphs that hatched, with good results, and their use was 
continued for all nymphs up to the 5th or 6th instar. The method was 
to collect chironomid egg masses each morning from out-door troughs or 
ponds, and to keep them in small pans of water until they hatched ; then 
to transfer the infant blood worms with a pipette to the boats with the 
nymphs. The nymphs seemed to thrive on them. But the blood worms 
have a nasty habit of making little dwelling tubes out of anything they 
can fasten down, and occasionally they would weave in the cast skin of 



270 Phylum Arthropoda 

a nymph, thereby destroying the evidence of the length of instars. 

From the 6th or 7th to the 9th or 10th instars, the nymphs were fed 
Ceriodaphnia, and Daphnia pulex; and after the 9th or 10th, Daphnia 
magna. The latter were also used to feed the full grown nymphs which 
developed the breeding stock. Cultures of these Daphnia had to be main- 
tained in the laboratory throughout the season. [For culture see pp. 207 
to 220.] 

Diseases. This species is susceptible to a disease which may occur 
under laboratory conditions (as well as in nature), if pond water is used. 
The invading organism is a green protozoan which lives in the lumen 
of the rectum, and is fatal to a large percentage of infected nymphs. A 
heavy infection is detectable, even with the naked eye, for the rectum 
appears green and may be seen in any but heavily pigmented nymphs. 
Methods of preventing infection were not worked out, but it might be 
effective to boil the water in which the insects are to live. 

The species is also subject, under natural conditions at least, to two 
species of trematode parasites, which may or may not be fatal; to a 
Gregarine infection ; and to external infestation by an aquatic mite.* 

Reference 
For the rearing Sympetrum vicinum see p. 272. 

METHODS OF REARING ODONATA 

P. P. Calvert, University of Pennsylvania 

THE following methods were successful in rearing larvae of Nanno- 
themis bella and Anax Junius from egg to adult, f 
Eggs of N. bella were obtained in the field in July, 1925, by dipping 
the abdomen of a female, caught in the act of oviposition, in a vial of 
water. As the eggs hatched the resulting 1st instar larvae were removed 
to various small dishes. After they made their first larval (nymphal) $ 
molt they were isolated, each one being placed in a glass salt cellar 
having a capacity of about 5 cc, covered with a sheet of glass, and num- 
bered. In each dish a small quantity of an aquatic plant (Elodea in 

♦Editor's Note: Others have reared Odonata from egg to adult both in Europe and 
America, but the preceding account appears to be the only one of maintenance of any spe- 
cies through successive generations. W. H. Krull (Ann. Ent. Soc. Amer. 22:651, 1929) 
reared Sympetrum obtrusum, finding 9 (in one case 10) nymphal instars. He fed the 
very young nymphs on Paramecium and Tubifex. (For culture see pp. 119 and 142.) 

The larger nymphs will eat almost any small living animals that come in their way. 
It is easy to rear them in, pillow cages of the sort shown in figure 42, and if the size of 
wire mesh be just small enough to retain them and large enough to admit their prey, they 
will feed themselves in any natural pond, protected by the cage from predatory enemies. 

J. G. N. 

t Abridged by the author from a paper in Proc. Amer. Philos. Soc. 68:227-274, 1929. 

+ Editor's Note: Equivalent terms; nymph is used in preference elsewhere in this book 
to denote this type of larva. J. G. N. 



Odonata 271 

earlier months, Lemna later) was kept growing. Through the nearly 
three years during which the rearing of N. bella was carried on, the dishes 
containing the larvae were kept on the inner sill of a window facing 
north. The temperature of the room, as indicated by a self-recording 
maximum and minimum thermometer, ranged from 89 to 50 F. (31. 6° 
to io° C). 

In December, 1925, each of the four living larvae, then somewhat over 
2 mm. long, was transferred from the salt cellar to a glass "caster cup 1 ' 
having a capacity of 15 cc, with the water and other contents in which 
it had been living previously. In June, 1926, each was again transferred 
from the caster cup to a finger bowl. Each caster cup and finger bowl 
was kept constantly covered with a piece of glass. 

In June, 1927, when the first of the N. bella larvae gave indication, by 
the whitening of its eyes, that transformation was approaching, its 
finger bowl was placed uncovered in a cylindrical glass battery jar. In 
the finger bowl was put a stick of wood so placed that it extended below 
the surface of the water; its upper end rested against a strip of wire 
netting attached to a framed window screen in front of a closed window 
facing south. Thus was provided a place for transformation. 

One larva, from the same lot of eggs, lived for another year, until June 
1928, before it transformed. 

When the larvae were about one week old drops of a culture of 
Paramecium were placed in the vessels in which they were living. [For 
culture see pp. 1 12-128.] Soon after, other Infusoria, small copepods, 
ostracods, rotifers, larvae of Anopheles, and other organisms smaller 
than N. bella were added. For nearly a year from Sept. 19, 1925, the 
food supply was chiefly small Crustacea, some of which bred in the dishes 
containing N. bella. In August, 1926, chironomid and ephemerid larvae, 
in September and October 1926, corixid and culicid larvae were given in 
addition. In late February, 1927, mayfly larvae were more frequently 
furnished; although these were taken from a swiftly flowing stream they 
lived in the absolutely still water of the dish for at least eight days and 
possibly longer. Similar observations were made in connection with other 
mayfly larvae, those of the genus Heptagenia being much more able to 
survive in still water than those of Baetis, for example. 

Special attention was paid each day that the N. bella larvae were 
examined to noting whether any possible living food material had sur- 
vived from the previous week and in only three cases was an entire lack 
of such found. The slow rate of development of the larvae may not, 
therefore, be ascribed to absolute starvation, although it is of course 
possible that the optimum food was not in the dishes. The hairs on the 
body of the larvae are fairly dense, and to them vegetable debris usually 
adhered to such an extent as to hide the body surface. This mass which 



272 Phylum Art hr op oda 

the slowly moving N. belia carried constantly served as a part of the 
food supply of the small Crustacea or of the still smaller animals (In- 
fusoria, rotifers) on which the Crustacea fed. 

Almost identical methods were employed in rearing a larva of Anax 
Junius* up to the time when it reached a length of 28 mm. It, the water in 
which it had been living, Lemna and other plants, were then transferred 
from the finger bowl to a battery jar, 145 mm. in diameter and 160 mm. 
high, and covered with a sheet of glass as before. A stick of wood was 
leaned against the side of the jar to give the larva an object on which to 
climb. It was there that its final exuviae was found. 

For the earlier stages small Crustacea were the chief food; when 5 mm. 
long, Culex and Anopheles larvae, small corixids, Daphnia, and mayfly 
larvae were used. During the final months the last named, especially 
those of the genus Heptagenia, were almost the only food given. A . Junius 
seemed to pay no attention to Asellus. 

Nevin f reared Sympetrum vicinum from egg to adult in stender 
dishes. During the first instar specimens from a protozoan culture were 
placed in the dishes. This culture proved to be mainly of Paramecium 
aurelia which were so small that the larvae did not seem to notice them 
and continued to die, probably from starvation. When Paramecium 
caudatum were fed the larvae ate them greedily. Another mixed culture 
used at this time contained many Stylonychiae and a few Vorticellae, 
although the larvae were never seen to eat the latter except with Daphnia 
to which Vorticellae were attached. Several ostracods of the family 
Cyprididae were eaten before the first molt. Later larger ostracods and 
copepods, including Cyclops and Diaptomus, were eaten. The copepods 
proved more proficient in capturing Paramecia than the dragonfly larvae 
and had to be removed until the larvae were larger. They also destroyed 
the exuviae of 5. vicinum or parts of them if the latter were left too long 
after the individuals had molted. During the last few instars Daphnia, 
Hyalella, and small mayfly and stonefly larvae were fed. Daphnias were 
not eaten from choice, but formed an important part of the food, espe- 
cially when mayfly larvae were not at hand. All larvae were fed and 
cared for three times a week. 

Miss Laura Lamb,$ in rearing Pantala flavescens, removed the larvae 
soon after hatching from the dish in which the eggs were laid and placed 
them in small stender dishes which were kept in a shaded part of the 

♦Editor's Note: Freshly laid eggs of Anax Junius may easily be obtained by setting a 
stem of cat-tail (Typha) in a place to attract ovipositing females. It should be set aslant 
in the surface of the open water several yards out from, the pond margin. It will then 
have preference over stems at the margin (where, presumably, enemies may lurk). If 
a fresh stem be used each day, the eggs may be dated. J. G. N. 

t Trans. Amer. Ent. Soc. 55:79, 80, 1929. 

X Trans. Amer. Ent. Soc. 50:289, 1924. 



Plecoptcra 2 73 

laboratory. During the first instar no food was given ; during the second 
and third stages Paramecium and small mosquito larvae formed the food. 
The remaining stages fed upon mosquito larvae and small crustaceans 
until the tenth instar, when pieces of earthworms and mayfly larvae were 
supplied. In the eleventh instar small fish were eaten. The individuals 
were kept in separate dishes after the third instar on account of cannibal- 
ism. 

Order plecoptera 

REARING THE STONEFLY, NEMOURA VALLICULARIA* 

THIS herbivorous stonefly is an inhabitant of clear-flowing cold spring 
brooks in the eastern United States. Adults appear on the wing in 
the latitude of Ithaca, N. Y., about the middle of March and disappear 
about the middle of April. They live among the vegetation of the brook- 
side where they run about actively and make short flights when the sun 
shines warmly about noon. They eat sparingly of the young leaves of 
wild Touch-me-not (Impatiens). They are able to survive without food 
for several days. 

About a week after transformation they reach sexual maturity and 
mate about midday. After mating, the females live for about a week, 
hiding in shaded places, and depositing gelatinous clumps of whitish eggs, 
numbering about 150 to 200 each. 

To obtain eggs for rearing, adults of both sexes were confined in a 
small box with porous bottom, so placed that it rested on the surface 
of the brook. This insured high humidity. Leaves of wild Touch-me- 
not were added for food, and blocks of decayed wood were placed inside, 
supported on slender twigs. The eggs were deposited on the under side 
of these blocks. 

Newly hatched nymphs were isolated in numbered shell vials, closed 
at the mouth with silk bolting cloth. These vials, assembled in an 
enameled tray, were immersed in the bed of the brook. They were ex- 
amined each day for cast skins. For food the nymphs were supplied 
with bits of decaying elm leaves. The vials were cleaned and the food 
was renewed once a week. 

Twenty-two instars were recorded during the nine months of develop- 
ment from hatching on the 2nd of July until emergence on the 29th of 
the following March. 

j. G. N. 
r 

♦Abstracted from an article in Lloyd Library Bull. 23, 1923 (Ent. Ser. No. 3) by 

Chen-fu Francis Wu, of Yen Ching University, Peiping, China. 



274 Phylum Art hr op oda 

REARING FALL AND WINTER PLECOPTERA* 

T)ECAUSE of their habit of congregating in places exposed to the 
O warming influences of the sun's rays, most of the fall and winter 
stonefly adults are easy to capture. During the warmer days of late 
fall and early winter they are likely to be found crawling about on ex- 
posed tree trunks, fence posts, or rocks located close to a stream inhabited 
by the nymphs, particularly if these objects are covered with an algal 
growth. 

In spite of the general belief that most adults of stoneflies do not feed, 
all of the fall and winter stoneflies of the Oakwood, Illinois, region have 
been found to feed. On many occasions the adults of Taeniopteryx 
nivalis, Allocapnia recta, A. vivipara, A. mystica, and A. granulata were 
observed feeding upon blue-green algae (Protococcus vulgaris) growing 
on tree trunks, fence posts, and stones near the habitat of the nymphs. 
A single specimen of T. parvula was also seen feeding on algae on a tree 

trunk. 

Eggs have been obtained by the simple expedient of catching females 
with egg masses and submerging them in water. The egg mass soon 
falls from the female and the individual eggs separate and settle to the 
bottom of the dish. 

Simple methods have sufficed for rearing fall and winter stoneflies. 
Fullgrown nymphs are collected and kept alive in small tins containing 
moist leaves until the emergence of the adults. In order to observe closely 
feeding, mating, and egg-laying habits, the adults may be kept alive and 
healthy in small hermitically sealed aquarium jars containing a layer of 
moist sand, a supply of bark bearing a good growth of green algae, and 
old leaves and stems on which the adults may run around or in which 
they may hide or rest. The eggs will hatch in glass tubes covered at both 
ends with fine bolting and submerged in unpolluted streams. No doubt, 
if supplied with the proper quality and quantity of food, the nymphs 
will develop under the same conditions.** 

The nymphs of Taeniopteryx and Allocapnia, both young and nearly 
grown, are herbivorous. No doubt a few protozoans are occasionally 

♦Abstracted from an article in III. Nat. Hist. Bull. 18: 345. 1929, by Theodore H. 
Frison, Illinois Natural History Survey. 

** Editor's Note: Lucy Wright Smith, of Cornell University, reported in Ann. Ent. Soc. 
Amer. 6:203, 1913, the use of essentially these same methods for Perla immarginata. 
Adults were kept in small wire cages and mating and egg laying occurred readily in 
captivity. These nymphs are carnivorous, however. Black fly larvae and mayfly nymphs 
proved satisfactory as food. . . 

By these same methods adults of Pteronarcys have been obtained and kept in captivity 
where mating takes place readily. Eggs obtained from these or from wild adults have 
been taken in June and have hatched in running water the following February. Due 
undoubtedly to a lack of the proper food, however, these newly hatched nymphs were 
not reared. Older nymphs eat well decayed leaves. M.E.D. 



Is opt era 275 

taken into the alimentary tract along with the diatoms, but yellow fresh- 
water diatoms and decaying vegetation constitute their main food. Old 
leaves which have fallen into the water apparently supply the bulk of 
decaying vegetable matter used by the nymphs for food. 

M. E. D. 

Order isoptera 

LABORATORY COLONIES OF TERMITES* 

Esther C. Hendee, Limestone College, Gaffney, South Carolina 

DAMP-WOOD TERMITES 

THE damp-wood termites, Zootermopsis angusticollis and Z. nevaden- 
sis, because of their comparatively large size and the ease with which 
they may be maintained in laboratory colonies, are satisfactory termites 
for experimental work. These species are confined to western North 
America. Although they are sometimes found in sound wood, they more 
frequently inhabit rotten wood. Rotten logs which lie near a stream are 
favorable places in which to look for damp-wood termites. Since the 
termites seal up the entrance to their burrows, it is usually necessary 
to chop into the log in order to detect their presence. 

The termites are brought to the laboratory in large pieces of the wood 
in which they have been found. If more than a few hours are to elapse 
before they reach the laboratory, the wood is wrapped in damp paper to 
prevent drying. 

Upon reaching the laboratory the termites are removed from the wood 
by splitting it cautiously so as to expose the termite galleries and then 
gently knocking the ends of the pieces. The termites thus dislodged are 
allowed to fall into a large container or, as an extra precaution against 
injury, onto a sheet of cloth. At best, a few termites will be injured. It is 
therefore advisable to wait two or three days before using the termites 
for any experimental purpose. During that time weak or injured indi- 
viduals may be detected and removed from the group. 

If it is desired to transport termites which have already been removed 
from the wood in which they were found in the field, the following 
method of packing is used. Numerous holes are bored in some blocks of 
wood by means of an auger. The termites are placed in these holes. The 
blocks are then wrapped in many layers of damp paper and finally en- 
closed in heavy dry paper or a box. 

Termites from different natural colonies are, if possible, kept separate 

* Further information concerning the biology of termites and a biblography are given 
in "Termites and Termite Control" which is edited by Kofoid, Light, Horner, Randall, 
Herms, and Bowe (1934). 



2 76 Phylum Arthropoda 

in the laboratory. More nearly similar genetic constitution within the 
group, a condition often desirable in experimental work, is thereby as- 
sured. Furthermore, the excessive cannibalism which is likely to follow 
the mixing of colonies is avoided. If colonies must be mixed, as is some- 
times necessary in order to get the numbers of termites desired for a 
given experiment, about a week is allowed for cannibalism to subside 
before the mixed group is used in any experiment. 

Individual termites are most safely handled by lifting them on a 
camel's hair brush or a tapering piece of stiff paper. Large numbers of 
termites may be transferred from one container to another by pouring 
them through a trough or funnel made of cellophane. 

Petri dishes or shallow stender dishes are suitable containers for 
colonies of ioo or fewer termites. For larger colonies refrigerator dishes 
or moist chamber dishes have been found satisfactory. The lids of the 
containers, while not air tight, should fit closely enough and be heavy 
enough to prevent the escape of the termites. Anyone who has observed 
the crowded condition in termite galleries in nature will realize that 
there is little danger of getting too many termites into one container so 
long as there is opportunity for each termite to get at the food provided. 
In fact, large colonies thrive much better than small cononies. 

The dishes containing the termites are kept in a dark room or cabinet. 
Large, open dishes of water placed nearby aid in maintaining a desirable 
humidity. Damp-wood termites live and reproduce at ordinary room 
temperatures (20 to 23 ° C). They will survive throughout a con- 
siderably greater temperature range, provided the change is gradual. 
They should be protected from sudden changes in temperature. 

Rotten, fungus-containing wood constitutes a satisfactory diet for 
damp-wood termites (Hendee, 1934)- The pieces of wood should be 
sufficiently heavy and so placed that they may not be moved by the 
weight of the termites. 

Cook and Scott (1933) describe an artificial diet for termites which 
consists of sucrose, casein, "Crisco", salts, cod liver oil, and rice polish- 
ings, all incorporated in an agar gel. Filter paper, while it fails to com- 
prise a complete diet for termites (Cook and Scott, 1933 ; Hendee, 1934) , 
affords a convenient source of the carbohydrate portion of the diet. 

The food should be kept slightly damp. If the containers are left 
undisturbed, the termites will partially seal the lids from the inside with 
fecal matter. In this way they partially control the humidity within the 
container and prevent evaporation of water from their food. If the 
containers are opened frequently, however, a few drops of water should 
be added to the food every two or three days. 

Fungi have been shown to supply an essential part of the natural diet 
of termites (Hendee, 1934). At the same time they constitute a poten- 



Isoptera 277 

tial hazard to the termite colony. A large colony will keep the fungi 
eaten down, but a small colony is often unable to do so and the termites 
themselves are attacked by the fungi. Precautions which may be taken 
to prevent this are: (1) choice of wood for food which does not contain 
excessive amounts of fungus mycelium, (2) frequent changes of the 
food when an artificial diet is used, (3) avoidance of excessive moisture 
in the food and of temperatures much above 20 C, (4) constant watch 
for excessive fungous growth, and (5) prompt removal of diseased 
termites. 

Colonies of damp-wood termites, as found in nature, are composed of 
nymphs and adults. The adults are of two distinct castes, soldiers and 
reproductives. There are two types of reproductives: primary repro- 
ductives and supplementary reproductives. In some colonies a pair 
of primary reproductives, founders of the colony, may still be present. 
They may be distinguished by their brown bodies. In colonies from 
which the primary pair has disappeared, and in groups of nymphs which 
have become separated from the parent colony, some of the nymphs 
develop into supplementary reproductives. The latter are caramel 
colored. 

Either type of reproductive, if included in a laboratory colony, will 
provide for the increase of the colony. Reproduction is carried on much 
more rapidly, however, by the supplementary reproductives. In a 
laboratory colony set up with nymphs alone, supplementary repro- 
ductives will develop and become functional within 4 to 7 weeks after 
the establishing of the colony. 

Primary reproductives, before they swarm, have wings. When these 
alate forms develop in a laboratory colony, they are either removed 
from the colony or allowed to swarm. If it is desired to start colonies 
with primary pairs, the alates, as soon as they become fully pigmented, 
are allowed to swarm in the laboratory. The covers are removed from 
the containers, and the alates, which are positively phototropic, fly 
toward a window or other source of light. After a few minutes they drop 
to the floor and shed their wings. They are then picked up and males 
mated with females for the founding of primary colonies. Males may be 
distinguished from females by the smaller size of the posterior three or 
four sternites. The growth of primary colonies, however, is very slow. 
So for most purposes it is preferable to use nymphs alone or nymphs 
and supplementary reproductives for establishing laboratory colonies. 

So far as I know, no one has succeeded in rearing termites from eggs 
without the presence of adult termites or older nymphs to care for the 
eggs and to groom and feed the young nymphs. I have found nymphs of 
the 4th instar satisfactory for experiments such as those on nutritional 
requirements. At that stage in development they are able to care for 



278 Phylum Ar thro poda 

themselves; their growth rate is rapid; and they are far enough from 
the adult stage to allow time for experiments of several months' dura- 
tion. Individuals of different instars may be distinguished by their 
relative head widths and the number of antennal segments (Heath, 
1927). 

DRY-WOOD TERMITES 

The dry-wood termites which belong to the subgenus Kalotermes are 
found in Mexico and in the southern part of the United States. They 
live typically in the dry, sound wood of buildings and other wooden 
structures. 

In choosing sound wood for feeding laboratory colonies of these 
termites the pieces are looked over carefully and all pieces which show 
evidence of a high content of resin or volatile oils are rejected. 

Examination of the wood upon which Kalotermes minor feeds in 
nature has shown it to contain fungus mycelium, even when it shows no 
macroscopic evidence of decay (Hendee, 1933). Therefore, if wood 
other than that in which the termites have previously been living is 
supplied to laboratory colonies, it is first infected with fungi so that it 
will be similar to the natural diet of the termites. This is done by 
dampening the wood, scattering crushed termite pellets over it, and 
allowing five or six days to elapse before it is fed to the termites. By 
that time fungus spores which were contained in the pellets will have 
given rise to a growth of fungus mycelium in the wood. 

While Kalotermes does not require as much moisture as Zootermopsis, 
the wood upon which it is fed should not be allowed to dry out com- 
pletely. Light ( 1934) reports that many species of Kalotermes require a 
minimum of 10% moisture in their food. 

Dry-wood termites will survive at ordinary room temperatures. For 
normal growth and reproduction, however, they require a minimum of 
25 C. For swarming, Harvey (1934) reports a temperature ranging 
from 8o° to ioo° F. (27 to 38 C.) to be the optimum for Kalotermes 
minor. 

Kalotermes minor is recommended by Kofoid and Bowe (1934) as 
the best species for use in testing wood and other materials for termite 
resistivity. They give a detailed account of the method of making the 
tests. 

SUBTERRANEAN TERMITES 

Subterranean termites of the genus Reticulitermes are found in nearly 
all parts of the United States. They demand a constant supply of 
moisture and are found in wood so situated that they may maintain 
runways into the soil. 



Corrodentia 2 79 

Dr. A. L. Pickens (unpublished communication) has devised what is 
probably the most successful method of keeping laboratory colonies of 
Reticulitermes. A thin piece of wood, preferably decayed, is carved 
with grooves along the grain and occasional connecting grooves across 
the grain. The grooves are made wide enough to allow termites to pass 
each other. The piece of wood is dampened slightly and placed with 
grooved side down on the bottom of a lidded glass vessel. A mixture of 
damp soil and sand is then packed in above it. By means of a wire a 
tunnel is sunk to connect with the end of one of the grooves in the wood. 
The termites are introduced through this tunnel and soon establish them- 
selves in the wood below. A few drops of water are added as needed to 
keep the soil damp. The colonies are kept at ordinary room tempera- 
ture. 

Bibliography 

Cook, S. F., and Scott, K.G. 1933. The nutritional requirements of Zootermopsis 

(Termopsis) angusticollis . J. Cell. Comp. Physiol., 4:95. 
Harvey, P. A. 1934. The distribution and biology of the common dry-wood 

termite Kalotermes minor. II. Life history of Kalotermes minor in Kofoid, 

Light, Horner, Randall, Herms, and Bowe, Termites and termite control (2nd edit. 

[2] ; Univ. Calif. Press), pp. 217-233, figs. 72-75. 
Heath, H. 1927. Caste formation in the termite genus Termopsis. J. Morph. and 

Physiol., 43:387- 
Hendee, E. C. 1933. The association of the termites, Kalotermes minor, 

Reticulitermes hesperus, and Zootermopsis angusticollis with fungi. Univ. Calif. 

Publ. Zool., 39:111. 

1934. The role of fungi in the diet of termites. Science 80:316. 

Kofoid, C. A., and Bowe, E. E. 1934. A standard biological method of testing 

the termite resistivity of cellulose-containing materials in Kofoid, Light, Horner, 

Randall, Herms, and Bowe, Termites and termite control (2nd edit. [2]; 

Univ. Calif. Press), pp. 517-553. figs. 131A-131C. 
Light, S. F. 1934. Habitat and habit types of termites and their economic 

significance in Kofoid, Light, Horner, Randall, Herms, and Bowe, Termites and 

termite control (2nd edit., Univ. Calif. Press), pp. 136-149. figs- 33-38. 
Snyder, Thomas E. 1935. Our enemy, the termite. 8°, pp. xiv + 196. Comstock 

Publishing Co. Ithaca, N. Y. 



Order corrodentia 

TROCTES DIVINATORIA* 

THE book-louse, a more or less cosmopolitan, parthenogenetic insect, 
is excellent as a source of material for life history studies by classes 
in entomology. Each student is given a mature specimen from which 
to obtain eggs, and the following equipment must be available: 

♦Abstracted from an article in Ann. Ent. Roc. Amer. 23:192, 1930, by O. W. Rosewaix, 
Louisiana State University. 



280 Phylum Arthropoda 

A large supply of drop-culture slides (those of matte finish with 
bottom of cell smooth but not polished are the best). 

Trays to hold slides which are of a size easy to handle. 

Cover slips, size 22 mm. 

Vaseline 

Corn meal (yellow meal seems to be the most satisfactory as food). 

The book-louse is placed in the cell of a drop-culture slide with a few 
grains of corn meal. Too much corn meal will make it impossible for a 
beginner to find the eggs. The cell is then covered with a 22 mm. cover- 
slip held in position by a trace of vaseline. As the eggs appear they 
are placed in separate slides and properly labeled. The eggs usually 
adhere to the point of a needle when touched, so it is an easy matter to 
transfer them. The work is best handled indoors in a room of fairly 
constant temperature. 

M. E. D. 

Order mallophaga 

LIPEURUS HETEROGRAPHUS* 

IN 1930 the writer undertook the study of Mallophaga under con- 
trolled laboratory conditions. A standard incubator which had a vol- 
ume of 29,970 cc. was used. Ventholes allowed for proper ventilation. 
Glass dishes exposing a total water surface of about 100 sq. cms. were 
placed on the bottom of the incubator to supply the air with moisture. 

Feathers were obtained from the neck region of a white fowl, and 
were cut into two parts. The fluffier basal portion was fastened in the 
center of a Syracuse watch glass by means of paste applied to the quill. 
The barbs of the feathers were trimmed so that they did not come in 
contact with the edges of the watch glass. 

A male and a female Lipeurus heterographus of unknown age were 
placed on the feather in each watch glass. The watch glass was num- 
bered and put in the incubator. The satisfactory temperature was 
found to be 33°-34° C When an egg was laid, the male and female 
were placed on a fresh feather in a new dish. 

Breeding was carried on during the summer of 1930. Specimens kept 
in the incubator until December bred actively during the entire period. 
The life cycle, from egg to adult, may take 32-36 days. There are 
three nymphal instars. 

It was found by experiments that lice reared in the incubator would 
mate and produce fertile eggs. The males and females of Lipeurus 
heterographus copulate readily in captivity. For study purposes it was 

♦Abstracted from an article in J. Paras. 5:304, 1934. by F. H. Wilson, Tulane Uni- 
versity. 



Embiidina 281 

more satisfactory to keep the males and females in separate dishes. 
After isolation, when placed together on feathers they would copulate 
in a very short time and their activities could be observed. 

In the life history studies the lice were given only the fluffier parts of 
the feather as food. They fed readily, particularly if they were removed 
from the feather for a time and then replaced. Experiments with 
feathers from the Little Green Heron (Butorides virescens virescens) 
showed that Lipeurus heterographus of the hen will feed on these 
feathers. 

Experiments on feeding with pulverized dried blood from a fowl show 
that this species relies on feathers for its essential food supply but 
supplements this with blood when it is obtainable. The maximum time 
for which 1st instar nymphs and adults could be kept alive on dried 
blood alone was three days. This is, however, not true of all biting lice 
since the writer failed to rear Menopon gallinae and M. stramineum, 
or even to induce them to lay eggs, under the same conditions under 
which Lipeurus heterographus thrived. Quite different food habits are 
indicated for Menopon stramineum (Wilson, 1933). 

Bibliography 

Wilson, F. H. 1933. A louse feeding on the blood of its host. Science 77:49°- 

M. E. D. 

Order embiidina 

OLIGOTOMA TEX AN A* 

DURING the spring and early summer a number of Embiids belong- 
ing to the species Oligotoma texana were captured alive and kept in 
captivity in a vial for three months. They were killed at the end of 
that time only because it was impossible to care for them longer. No 
difficulty was experienced in keeping them alive; in fact they seemed 
quite hardy. 

The individuals were kept alive in a loosely corked vial on a piece of 
rotting wood which they immediately covered with their webs. After a 
few days a cake crumb was dropped into the vial, and within thirty 
seconds embiid heads appeared at the openings in the webs, followed by 
their slender brown bodies as they made their way toward the food. 
By the next morning the crumb was completely hidden in a maze of 
silken tunnels which led from the rotting wood to it. Thereafter they 
were fed a diet of bread crumbs, and they always elongated their tunnels 
to include the food. 

M. E. D. 

* Abstracted from an article in Ann. Ent. Soc. Amer. 25:648, 1932. by Harlow B. Mills, 
Montana State College. 



282 Phylum Ar thro p oda 

Order dermaptera 

DERMAPTERA 

B. B. Fulton, North Carolina State College 

EARWIGS may be reared in any jar or cage provided with moist 
sand, but a large petri dish makes a very convenient cage. One 
side of the dish is filled to the top with moist sand and the other side 
left vacant for placing food materials. If the dish is shallow enough the 
nest will be made against the glass if kept darkened. If the dish is too 
deep, it may be partly filled with plaster or other material that the ear- 
wig can not dig into. Earwigs feed on a great variety of food but most 
species prefer food of animal origin. They require very little ventilation 
and little care. I have had a petri dish with some Anisolabis annulipes 
living in it for about a year. Sometimes I have forgotten to feed or water 
them for several weeks at a time but have always found a few still alive. 

Order orthoptera, Family grylloblattidae 

GRYLLOBLATTA 

Norma Ford, University of Toronto 

IN KEEPING Grylloblatta in the laboratory an attempt has been 
made to provide natural conditions. Found in their native habitat in 
the Rocky Mountains, they live in cold, damp places, where the tem- 
perature ranges from o°-s'° C., and the rotten logs or mosses are almost 
dripping with moisture. 

In the laboratory each insect is kept in a separate jar because of 
cannibalistic tendencies. The pieces of moss or decayed wood in the 
jar are always kept wet. In fact, a quarter to a half inch of water may 
be left in the bottom of the container. The jars are kept packed in ice 
in a large, insulated tub. Covering the tub and jars is a fairly loose 
packing of cotton batting. This allows for a certain variation in tem- 
perature. 

The insects are usually fed on mealworms, cut in small pieces, although 
pupae of ants, dipterous larvae, or adult flies give variety to the diet. 
The insects are fed about once a month and care is taken to remove 
from the jar any food which is left and has become moldy. 

Under these conditions the grylloblattas have lived for three and four 
years, slowly reaching maturity and depositing eggs. 



Blattidae 



283 



Family 



BLATTIDAE 



CARE AND REARING OF BLATTELLA GERMANICA 

C. M. McCay and R. M. Melampy, Cornell University 
Blatella germanka is a useful insect for physiological studies because 
of its quick rate of growth, rapid reproduction, and 
omnivorous feeding habits. An ordinary fish aquarium 
or museum jar covered tightly with cheesecloth may be 
used as a cage for a stock colony. Water may be supplied 
by an ordinary baby chick waterer. Absorbent cotton 
should be placed in the pan of the fountain as it prevents 
drowning of the insects. A stock diet of ground whole 
wheat 50',, dried skim milk 45%, and dried bakers' 
yeast 5% is adequate for growth and reproduction. 

For experimental work with individual insects or 
small groups, ordinary half-pint milk bottles may be 
used as cages ( Fig. 63 ) . The milk cap is perforated with 
numerous pin-holes to allow air to enter the container. 
The water is supplied by a vial containing damp cotton 
which is mounted on the cap by a cork. The diet to be 
studied is placed in a small paper cup or similar container. 




COCKROACHES* 



Fig. 63. — 
Rearing bot- 
tle for Blat- 
ella germanica. 



Cockroaches may be kept in wide-mouthed gallon 
glass jars, each containing a layer of sawdust on the bottom and a 
small pan of water. Over the top of each jar is stretched a piece of 
cheesecloth, held in place by a rubber band. A thin line of vaseline 
is placed around the inside shoulder of the jar, and the cockroaches do 
not attempt to pass this line.** The jars are kept in a rather dark place 
where the temperature averages 70 F. 

* Reprinted, with slight changes, from Turtox News 7:Xo. n, 1929, by John M. Kelley, 
General Biological Supply House. 

** Editor's Note: J. Franklin Yeager, of Iowa State College, has described to us the 
cage he uses for keeping large numbers of Periplaneta. It consists of a wooden framework 
on legs, with a bottom of pressed board, sides of glass below for observation and copper 
wire above for ventilation, and a top of pressed board with a hinged door for easy access. 
The legs supporting the cage are set in cups of water to keep out ants. 

A covered hole in the bottom with a metal shaft leading downward serves for removal 
of numbers of the roaches. They are swept down this shaft into a beaker edged at the 
top with vaseline. A rim of vaseline is also kept around the top of the glass portion of 
the sides of the cage. 

"Ootheca dropped by the females may be removed to other containers for hatching 
purposes. When the cage is kept clean of ootheca, molted exoskeletons, and dead in- 
dividuals, it is suitable for retaining large numbers of roaches over considerable periods of 
time. The cage may also be used with certain other species of insects." J. G. N. 



284 Phylum Arthropoda 

The food of these insects is very variable, but they exhibit marked 
preferences. We have found that a mixture of bread, cornstarch, and 
water, with added bits of lettuce or other green material, serves as a 
fine food. Sour milk and library paste are also relished. A great amount 
of water is required by the cockroaches. 

M. E. D. 

Family mantidae 

TWO SPECIES OF PRAYING MANTIS* 

TWO species, the common Stagmomantis Carolina and a big Chinese 
species, Paratenodera sinensis, have been reared in the laboratory 
and carried through several successive generations in as many successive 
years. 

Eggs taken from twigs out-of-doors, or laid in the laboratory, were 
placed outside all winter. When they began to hatch, in May or June, 
individuals were isolated in homeopathic vials, tightly stoppered. These 
vials were handled in wooden racks holding about a dozen. In each vial 
a strip of filter paper furnished a support to which the baby mantis could 
cling. About the third molt the insects became rather too big for the 
vials and were transferred to 4-ounce wide-mouthed bottles with cork 
stoppers and a strip of cardboard to stand upon, individuals still being 
kept separate. Before the last molt they were given still larger accommo- 
dations, either quart specimen jars or 6-inch stender dishes. 

As is well known, the praying mantids are preying insects and are 
classed as beneficial because they eat plant lice, caterpillars, and various 
other enemies of vegetation. They are, furthermore, very cannibalistic. 
When hungry they ate readily almost every insect species that came 
their way, the only invariable requirement being that the food be served 
"alive and kicking." Tiny leafhoppers, Drosophila, Meromyza, minute 
"looping" caterpillars, etc., collected in a sweep net and distributed to 
each vial, furnished most acceptable food.** Bigger leafhoppers, larger 
flies and caterpillars, and young grasshoppers became suitable food as 
the mantids increased in size. After the third molt, they could capture 
houseflies, and never seemed to tire of the diet. Quantities of these were 

♦Abstracted from an article in Ent. News 37:169, 1926, by Mary L. Didlake, Uni- 
versity of Kentucky. 

** Editor's Note: R. A. Roberts, of Iowa State College, reports in Canad. Ent. 60:209, 
1928, the breeding of Stagmomantis Carolina in glass lantern globes with gummed labels 
stuck on the inner surface to provide footholds. In each cage a twig with several leaves 
was held upright in a glass vial filled with water and plugged with cotton. Sufficient 
moisture was important for the young mantids and it was found necessary to sprinkle 
these leaves daily. For the first few weeks the mantids were fed entirely on aphids by 
sticking aphid-infested twigs in the vials of water. M. E. D. 



Tettigoniidae 285 

caught in wire traps placed outside a laboratory window, baited with 
banana. 

Full-grown adults, if hungry, ate almost any living thing: spiders, 
hairy caterpillars (Datana, Apatela), furry moths, bad-smelling stink 
bugs, hard-bodied wasps (Vespa), huge cockroaches, and grasshoppers 
as large as themselves. Some individuals which survived late in the 
season when other insects grew scarce, relished fat chestnut worms, meal 
worms, etc. 

M. E. D. 
Reference 

For the feeding of mantids see also p. 242. 



Family tettigoniidae 

CEUTHOPHILUS* 

THE eggs of the majority of the species of these camel crickets are 
deposited in the ground at a depth determined by the length of the 
ovipositor. However, eggs have been found in rotten wood. Many of 
the more strictly hypogeic species probably oviposit in their burrows. 
Eggs of C. latibuli laid at night in cages in the laboratory hatched in 
between two and four weeks. On the other hand it is highly probable 
that many species normally overwinter in the egg stage, wholly or in part. 

Post-embryonic development varies in rate, depending on the amount 
and nature of the food supply and presumably on temperature and 
moisture conditions. Specimens of C. virgatipes reared with abundance 
of food but under otherwise normal conditions reached maturity long 
before adults were taken in the open and are far larger and differently 
proportioned than any feral specimens seen. 

Observations on caged individuals of several species of Ceuthophilus 
show that the members of this genus are immobilized by strong light, 
are inactive by day but extremely energetic by night, are unaffected by 
sound stimuli, but highly sensitive to air movements and other mechan- 
ical stimuli, and to odors. 

All Rhaphidophorinae appear to be practically omnivorous. Blatch- 
ley** states that caged Ceuthophilus fed upon meat as well as upon pieces 
of fruit and vegetables, appearing to prefer the latter. My own observa- 
tions accord with these; caged individuals of C. virgatirpes, C. latibuli, 
C. peninsularis, and C. pallidipes ate with avidity cheese, butter, jam, 
sweet fruits, fresh and dried meat, sugar, dead insects, and other items of 

♦Abstracted from Univ. oj Fla. Biol. Ser. 2, No. 1, 1936, by T. H. Hubbell, Uni- 
versity of Florida. 
** Blatchley, W. S. The Locustidae of Indiana. Proc. Ind. Acad. Set. for 1892, p. 141. 



286 Phylum A rthropoda 

food. They were especially fond of peanut butter, neglecting other 
favorite foods when it was available. A few individuals of C. latibuli 
were reared to maturity on a diet consisting solely of peanut butter and 
sugar. Grass and other green vegetation were rejected, as were bread, 
flour, and other starchy substances unless no other food was supplied, 
when they were eaten sparingly. In colonies of Ceuthophilus there is 
heavy mortality when ecdysis occurs for the soft, helpless teneral insects 
are eaten by their cannibalistic mates. Only a small proportion of the 
individuals in a crowded cage survive to reach maturity. 

The need for shelter and protection from low humidity are dominating 
factors in the lives of these insects. Caged individuals of C. latibuli 
continually undermined dishes of food and water placed on the sand. 
Though many spent the day clustered together in the shadowy upper 
corners of the cage, others hid themselves more or less completely in 
burrows. 

M. E. D. 

Reference 

For the feeding of meadow grasshoppers see p. 242. 



Family gryllidae 



ON REARING GRYLLIDAE 

B. B. Fulton, North Carolina State College 

GROUND crickets, Nemobius and Gryllus, may easily be reared in 
large battery jars of one gallon or more capacity. Single pairs do 
fairly well in jars as small as one quart. There should be about one 
inch of sand in the bottom, kept slightly moist. If the jars are at least 
8 inches tall it is not necessary to cover them. Mold develops more 
rapidly on the food materials if the jars are covered. A watch glass or 
small dish of water may be kept in the jar with larger crickets but the 
very young crickets may drown in it. This is not necessary if the sand 
is kept moistened. The jars should be kept out of direct sunlight in the 
summer. The crickets may be fed on a great variety of food materials.* 
Those things that do not mold too quickly are satisfactory. I have used 
lettuce, grass, and fruits, but the least troublesome food is one that I 
make from rolled oats. The rolled oats are ground in a mortar with a 
little sugar and skim milk powder and enough water to make a stiff 
paste. This is spread thinly on heavy wrapping paper with a spatula 

♦Editor's Note: Norman Criddle reported in Canad. Ent. 57:79, 1925, that Gryllus 
assimilis pennsylvanicus and G. a. luctuosus consume animal matter with relish and that 
he has reared 1st instar nymphs on tabanid flies alone. Moistened bran was also used. 
M. E. D. 



Acrididae 287 

and allowed to dry. This food will keep indefinitely. About 1 square 
inch every three days, or oftener in damp weather, will feed several 
crickets. I have kept cultures going on this food for over a year. I 
have not experimented to find out whether the sugar and milk are 
necessary. The crickets will eat dry rolled oats readily. 

Burrowing crickets such as Anurogryllus and mole crickets may be 
reared in jars supplied with several inches of sand. If observations are 
to be made on the underground habits a special type of cage is necessary. 
For this I have confined the sand between two vertical pieces of glass, 
separated by not more than the width of a normal burrow. The food 
and water may be introduced by having the top of the frame removable. 
Burrowing crickets that forage above ground need a small attached cage 
at the top. They may also be kept in a cylindrical jar containing a 
shorter and narrower jar or tin. The space between the two should be 
only the width of a burrow and filled with sand. This type, or the glass 
plate cage, should have an outer removable cover to keep the burrows 
dark except when under observation. 

Tree crickets and bush crickets need a screen or cloth-sided, partly 
glassed, cage supplied with potted plants or cut plants in water. The 
plants should be sprinkled with water every day. A known host plant 
should be used if possible. Most adult crickets may be kept alive a 
long time by placing a few drops of sugar water or pieces of sweet fruits 
on the foliage. For life history work it is necessary to have the proper 
kind of plant material for oviposition, for many species are inclined to 
be exacting in their requirements. If these are not known it is best to 
supply at the same time a variety of kinds and sizes of plant stems. 

Reference 
For the feeding of crickets see also p. 242. 

Family acrididae 

CULTURE METHODS FOR GRASSHOPPERS 

E. Eleanor Carothers, State University of Iowa 

WITH foresight and a little equipment, these insects may be used 
as a convenient source of live material which a laboratory may 
have on hand in any desired stage at all times. The rearing of hardy 
species, like many other things, is simple if one knows how, but faulty 
methods may result in the loss of a stock in the midst of an experiment. 
Aside from studies growing out of economic problems and the recognized 
value of short-horned grasshoppers as cytological material, they are well 
adapted for physiological, embryological, and genetical studies. 



288 Phylum Ar thro p oda 

I. Equipment. One of the chief essentials is a sunny, warm room 
with good ventilation. Exposure to direct sunlight without the inter- 
vention of glass is desirable and may be achieved usually during the 
spring and summer. Constant temperature rooms or an incubator where 
one can obtain temperatures of from 25 to 30 C. at will and an electric 
refrigerator for the storage of hibernating eggs are necessary for a con- 
tinuous supply of the various stages. 

Two types of cage are desirable. A large size for stock or mass- 
cultures and a smaller size for special experiments. The following descrip- 
tion applies to cages which have proven satisfactory. They may be 
made in any ordinary workshop. The frame and bottom of each are 
made of %-inch cypress, since this wood does not warp, split, or rot 
easily. The ends, top, and back of each type are of copper gauze with 
18 meshes per inch. All parts of the cages must be tight enough to pre- 
vent the escape of the newly hatched insects and the sifting out of sand 
or soil. The bottom in both cages is built up with the cypress to form 
a box about 2% inches deep. 

The larger cage is 25 inches long, 15 inches wide, and 18 inches high. 
A sliding tray for the floor which may be pulled out without otherwise 
opening the cage, while not essential, greatly facilitates cleaning the 
cage. The front is a glass plate fitted into grooves above and below so 
that it slides in from one end. It may be cut vertically at the center for 
greater ease in handling. Such a cage will accommodate several hundred 
newly hatched grasshoppers and twenty-five to fifty adults, depending 
upon their size and hardiness. Crystallizing dishes 2 or 3 inches deep 
or even cigar boxes packed firmly with damp sand are placed in these 
cages during the egg-laying period. If eggs of known age are desired the 
sand may be removed and gently sifted daily. 

The smaller cage is 7 inches long, 6 inches wide, and 8 inches high. 
The front is a glass plate fitted vertically into grooves. It is cut about 
3 inches from the bottom so that the top part may be slipped up to open 
the cage, thus the bottom of the cage will hold sand to the depth of 3 
inches. Twelve to twenty-five young individuals may be kept in such a 
cage. The amount of care necessary varies greatly with the hardiness 
of the species. For the immature insects a thin layer of clean, dry sand 
is kept on the floor and changed as often as cleanliness demands. Mold 
must be avoided. When it is time for eggs to be deposited, 3 inches of 
sand is placed in the bottom and the cage dipped in water every week 
or ten days. The egg pods may be obtained by passing the sand through 
a sieve. After ten days to two weeks of development the eggs of most 
species reach the diapause and may be stored in moist sand in a re- 
frigerator. After a few weeks they may be removed to an incubator or 
a laboratory room in batches as desired and will resume development. 



Acrididae 289 

II. Choice oj a Species. The following points should be considered 
in selecting a species for laboratory culture if the problem does not 
necessitate the use of a particular species: (1) hardiness, (2) availabil- 
ity, (3) availability of natural food plants, (4) number of generations 
per year and size of broods. Let us briefly consider these points. 

1) Hardiness and 2) availability. Usually indigenous species are 
preferable since change of climate and altitude are thus eliminated. The 
shift from freedom to captivity is severe under the best conditions. 
Correlated with the above advantages are natural conditions for their 
food plants. I have no doubt that hardy species may be found in every 
locality. In general, tryxalines are the most delicate, oedipodines 
intermediate, and acridines the most vigorous, but as a group the latter 
are the most restricted as to their food plants. However, many species 
of Melanoplus in addition to being extremely hardy are almost om- 
nivorous so far as plants are concerned. M. dijjerentialis in particular 
on account of its wide distribution and large size is very favorable for 
general laboratory purposes. M. bivitatta in its range would be equally 
satisfactory. Romalea microptera, a large, almost flightless acridine 
from the south thrives in captivity and may be purchased from collectors. 
Owing to its size the provision of food and space becomes a problem if it 
is necessary to keep large numbers on hand. Dactylotum pictum is 
another acridine which does well in captivity, perhaps because of its 
wingless condition. 

Most oedipodines are strong flyers and perhaps for that reason do not 
thrive in captivity. Notable exceptions exist, however. Encoptolophus 
subgracilis taken from Tuscon, Ariz., to Philadelphia, proved to be the 
most satisfactory grasshopper I have ever raised so far as ease of culture 
is concerned. It is hardy and produces a generation in six weeks. Chor- 
tophaga australior, also a southern form, is nearly as favorable. More 
northern species of these genera normally hibernate over the winter, 
E. sordidus in the egg and C. viridijasciata in the nymph stage. Both 
will complete their development without a pronounced pause if kept at 
2 2°-25° C. Trimerotropis maritima and T. vinculata are satisfactory 
but will produce normally only one generation per year. 

The tryxaline, Chloealtis conspersa, is a notable exception to the rule. 
It is very hardy and conveniently lays its eggs in old, somewhat rotten 
wood, preferably fallen branches of trees. The eggs are laid in August 
and September and normally hatch in the spring. 

All of the species mentioned are known to be sufficiently hardy to 
make good laboratory stock. But no one should be deterred from trying 
out other species, especially if they are indigenous so that the stock may 
be replenished from nature. 

3) Availability of natural food plants. As has been stated, many 



290 Phylum Arthr op oda 

acridines are limited to certain weeds as food plants. These plants in 
turn are restricted to given soil and climatic conditions. Alkalinity or 
acidity of the soil is often a limiting factor for the plants. In case it 
is desirable to use a restricted species away from its normal habitat, 
not only seed or young specimens of its food plant should be take but 
also soil should be shipped or at least samples taken for analysis so that 
the necessary constituents may be added to the soil in the new locality. 

4) Number of generations per year and size of broods. Some species, 
like Hesperotettix viridis, lay only 5 to 8 eggs per pod and one female 
does not make more than 6 pods, so that, if there is normally only one 
generation per year, a given female will not produce more than 40 off- 
spring. Most oedipodines lay from 15 to 24 eggs in a pod and produce 
3 to 4 pods, so that one female may have as many as 75 offspring. 
Furthermore, some southern species, already mentioned, may produce 
6 to 8 generations per year in the laboratory, although they probably do 
not produce so many in nature. 

Melanoplus differentialis, M. bivitatta, and Romalea microptera fe- 
males on the other hand lay as many as 150 eggs in the first pod and a 
given individual may make 3 pods each with progressively fewer eggs. 
Such females will produce from 200 to 300 offspring. Fortunately 
they are restricted normally to one generation per year. 

III. Life Cycle. Three well defined stages occur: 1)— Embryonic, 
extending from the first cleavage of the egg until hatching. Some eggs, 
as noted by Nabours for tettigids and Slifer and King or M. differen- 
tialis, develop parthenogenetically. Most northern species hibernate 
over winter in the eggs. 2) — Nymphal, from hatching to the last molt, 
marked by 5 instars during which progressive development of pre- 
existing structures takes place. Certain species hibernate for the winter 
in the 2nd and 3rd instars. 3)— Adult, extending from the last ecdysis 
until death. Usually an individual does not live more than 3 months 
after becoming an adult. 

IV. General Care. In order that experimental results may be valid 
the stock must be healthy. The following suggestions may help to 
attain that end. 

1 ) Cleanliness. Debris should be removed and clean dry sand should 
be scattered on the floor of the cages at least once each week. Except 
during egg-laying the cages should be kept dry. Warmth and sunlight 
are essential. 

2) Handling. Shell vials are convenient for capturing the young 
insects when it is necessary to study or move them about. Older ones 
may be caught gently by the thorax or wings with the fingers. Never 
catch them by the jumping legs since these are readily detachable. The 
blood clot which then forms, as well as the inability of the crippled insect 



Acrididae 291 

to properly suspend itself, causes trouble at ecdysis. The loss of a leg is 
not so serious for an adult as for a nymph. 

3) Food. An abundance of suitable food should be present in the 
cages at all times, either growing in soil or kept fresh in water. If water 
is used, care is necessary to prevent the insects from drowning. Tryxa- 
lines and oedipodines feed mostly on grasses. Various species of Poa 
and Andropogon are suitable. Wheat and millet are satisfactory and 
readily grown in small flower pots in the laboratory. Oats are totally 
unsuitable, as are some grasses. Dandelion, plantain, and clover are 
good for giving variety. Lettuce and apple will tide many species over 
a period of food scarcity. For some species lettuce will serve as the chief 
food. These foods are adequate for the previously mentioned acridines 
also. But many of this group are highly specialized in regard to their 
food plants. Hypochlora alba is restricted largely to Artemesia jrigida, 
Hesperotettix speciosus to sunflowers, H. viridis to Grindelis and certain 
species of Solidago, H. pratensis to a different Solidago. In fact, the 
experienced collector in this group looks for the food plant of the species 
sought rather than the animals themselves when starting to collect in 
a new region. 

4) Diseases and Parasites. The chief dangers to laboratory cultures 
are diarrhoea caused by an Amoeba (recently described by R. L. King) 
which may wipe out cultures during cold damp periods, and molds which 
thrive in damp places in the absence of sunlight. Hyphae of the fungi 
grow throughout the body cavity. Gregarines also cause trouble, espe- 
cially when cages with damp sand and much debris are kept at a con- 
stant high temperature as in an incubator. Nematodes, too, sometimes 
become troublesome, especially if the cultures are fed on grass brought in 
from a cool, damp place and put into the cages without a thorough 
washing. And, finally, a small parasitic wasp may attack the eggs. 
It thrives chiefly on dead eggs and I am not sure that it ever attacks 
those in a healthy condition. It is more apt to be present when soil is 
used in place of clean sand for the eggs. 

None of the above troubles is a serious menace to cultures which are 
given proper care. In case the insects start to die transfer them daily 
to cages which have been washed and sterilized in very hot water, dis- 
card all individuals which are obviously sick, and correct faulty cultural 
conditions. 



292 Phylum Arthr op oda 

THE GROUSE LOCUSTS* 
Robert K. Nabours, Kansas State College 

THE grouse locusts, so-called probably because of a fanciful re- 
semblance of some of them to the grouse (Tetraoninae), are among 
the smaller Orthoptera. They show conspicuous dimorphism as to 
length of wings and pronotum, with occasional intermediates. There 
are extraordinary variations in the conspicuous color patterns on the 
pronota, the legs, and other parts of the body. The stripes along the 
median pronota vary in color and width, and there are many kinds of 
spots, specks, mottlings, and all-over colors. One species, as Paratettix 
texanus, may exhibit nearly all the colors and patterns. Food, light, and 
other features of the environment do not appear to condition the colors 
and patterns in any special way. (Hancock, 1902 ; Nabours, 1925 and 
1929.) 

The subfamily Tetriginae is widely distributed over the tropics and 
temperate zones. Hancock (1902) estimated that there were more 
than 100 species in North America alone. They mainly inhabit moist 
areas, along the margins of ponds and streams, and in forests, though 
they may live, temporarily at least, in quite dry places. Some, as 
P. texanus and A. eurycephalus in southern Texas and Mexico, are 
found more abundantly along the flat, algae-covered margins of ponds 
and streams, in the absence of larger vegetation and much exposed to 
the sun. Others, as Tettigidea lateralis, are found farther back in the 
grasses, or higher vegetation, where there is more shade. 

The northern grouse locusts (roughly from the line of the Ohio, 
lower Missouri and Kansas rivers in the U. S. A.) probably produce 
one, or an average of about one and one-half generations a year. The 
cold weather coming on in October, or November, finds both adults and 
nymphs, and they all go into hibernation for the winter. They stop in 
tufts of grass, under stones, pieces of wood, etc., but receive little real 
protection from the weather. They have been observed to endure and 
survive a temperature lower than o° F. However, there is usually a 
high mortality, due probably as much to desiccation as to cold. There 
is no regularity about their going into, or emerging from, hibernation. 
They do not become inactive till the cold weather actually arrives, and 
they become active during any very warm periods. During aberrantly 
early warm weather they emerge from hibernation, to be driven back 
later in the spring if there is more cold weather. 

When warm weather arrives in the spring the adults mate and soon 

♦Pending a much needed, comprehensive revision of this subfamily by taxonomists, 
I have undertaken to follow Hancock's classification of the few species used in the 
studies of inheritance. However, I now propose to follow A. N. Caudell's (Smith. Inst.) 
suggestion, in a letter, 1932, of subfamily Tetriginae; family Acrididac. 



Acrididae 



293 



lay eggs during a period of several weeks, in March, April and May, 
depending on the latitude and season. The nymphs which have hiber- 
nated become adults, mate, and lay eggs some weeks later. 

The southern species give about 
four generations a year in the green- 
house. They probably do not have 
a definite hibernating period in 
their natural habitats. 

Paratettix texanus, Apotettix 
eurycephalus, Tettigidea lateralis, 
Telmatettix aztecus, and Acrydium 
arenosum have been bred in the 
greenhouse at the Kansas Ag- 
ricultural Experiment Station. 
The first two mentioned have been 
bred extensively. A. eurycephalus, 
due to factors which have not been 
ascertained, breeds better than any 
of the others used. All, except A. 
arenosum and some of the T. 
lateralis, were from stocks secured 
in southern Louisiana and Texas, 
and the region of Tampico, Mexico. 
They are bred best in a green- 
house laboratory, with the tem- 
perature ranging around 8o° F. 
A variety of cages may be used, 
but 8" x 12" glass cylinders, with 
lids of 16-20 mesh wire, set in steam 
sterilized loam in bulb pots, serve 
very well. Sterilized sand is placed 
in the lower part of the pot, and 
around the cylinder. A smaller 
empty pot is placed upside down 
over the hole in the bottom of the 
bulb pot. It is supposed to aid in 
aerating the soil, and the food is 




Fig. 64. — Cage for rearing grouse locusts 
and apparatus for transferring progeny. 
(Transfer apparatus developed by Edgar 
Millenbruch.) The glass cylinder is 8" 
in diameter and 12" high, a, suction con- 
nected with water air pump or sweeper; 
b, cheesecloth; c, glass chamber into which 
progeny are sucked; d, the glass cage 
(covered with 16- to 20-mesh wire screen 
lid); e, tape; f, corks; g, sand; h, bulb 
pot; i, loam; k, inverted pot; m. air 
chamber; n, algae. 



placed on its extension above the soil (Fig. 64). A pair of adults is 
placed in a cage. The eggs are laid in the soil, or in masses of algae. A 
few hours or a day or so after hatching the offspring are picked up by a 
suction tube and transferred in batches of 20-25 to newly made-up 
cages. At the 3rd-4th instar records may be made of the color patterns, 
which do not materially change at any time, and the males and females 
separated. 



294 Phylum Ar thro poda 

The extraordinary and diagram-like color patterns, the good size of 
their chromosomes and, perhaps, a few other features justify the extreme 
effort necessary in breeding the grouse locusts for genetics studies. I 
believe that it would be easier and more economical to breed 100,000,000 
Drosophila than 1,000,000 of these Orthoptera. 

As already stated they eat mainly algae. In a humid climate, and 
where they are available, the scrapings from the pots in greenhouses 
serve very well. In the climate of Manhattan, Kansas, the filamentous 
algae only are available, Hydro diet yon sp. being the most satisfactory. 
Much difficulty is exerienced from the invasion of black or blue-green 
algae, fungi, and masses of decayed algae, and the acidity and other 
chemical conditions of the soil which render it unavailable. 

It has been ascertained that an extra, winter generation of the Man- 
hattan, Kansas. Acrydium arenosum may be secured by exposing parents 
and then the offspring to continuous lights, either the ordinary white 
light from Mazda bulbs or mercury vapor radiation through ordinary 
glass, the latter being somewhat more effective (Sabrosky, Larson, and 
Nabours, 1933). It is believed that these insects offer very fine op- 
portunity for irradiation work of various sorts. 

The females of P. texanus, A. eurycephalus, and others reproduce 
also parthenogenetically, the unfertilized eggs, with rare exception, pro- 
ducing females. The females are generally about three times as pro- 
lific when mated as when unmated. A mated female may have part of 
her ova fertilized, and also produce from unfertilized ova, by partheno- 
genesis, an additional number of offspring which are nearly always 
females. 

Bibliography 

Hancock, J. L. 1902. The Tettigidae of North America 

Nabours, R. K. 1925. The grouse locust Apotettix eurycephalus. Kan. Tech. 

Bull. 17. 

1929. The genetics of the Tettigidae. Bibliographia Genetica 5:27. 

Nabours, R. K., and Robertson, W.R.B. 1933. An X-ray induced translocation in 

Apotettix eurycephalus, Hancock (Grouse Locusts). Proc. Nat. Acad. Sci. 19:234. 
Robertson, W. R. B. 1935. Demonstration of effects of X-rays on male germ 

cells in Apotettix eurycephalus; Amer. Nat. 69: 
Sabrosky, Larson, and Nabours. 1933. Experiments with light upon reproduction, 

growth, and diapause in grouse locusts (Acrididae, Tetriginae) ; Trans. Kan. 

Acad. Sci. Vol. 36. 



Thysanoplera 295 

Order thysanoptera 

REARING THRIPS TABACI* 

THE thrips were reared on successive plantings of onions in a green- 
house. The thrips were transferred from the plants to vials by 
sucking them into a glass tube which was about 4 inches long and about 
Y 2 inch diameter. [See Fig. 41.] A cork was placed in one end of this 
tube, through which a 2 -inch length of glass tubing % inch in diameter 
was inserted. A 12 -inch piece of rubber tubing was inserted in the other 
end of the large glass tube. A piece of silk placed over the end of the 
rubber tube prevented the passage of thrips into it when they were being 
sucked into the glass tube and also furnished a tight fit between the rub- 
ber and the glass, thus preventing the escape of the thrips. The rubber 
made the apparatus flexible so that it was easy to reach the thrips on 
any part of the plant. When the desired number was secured, the cork 
was removed and the thrips shaken into vials, from which they might 
later be transferred to the feeding cages. 

The cage finally evolved for controlled feeding experiments was a 
Syracuse watch glass and its cover. The inside of the bottom of the 
glass was covered with white blotting paper, the edges of which were 
sealed to the glass by means of hot paraffin to prevent the thrips being 
caught between the paper and the glass. When the cage was inverted 
the blotting paper absorbed the excess moisture, preventing con- 
densation within the cage and the possible drowning of the thrips. 
It also aided in maintaining a high humidity which seemed de- 
sirable. Three discs of blotting paper the size of the coverslip, placed 
one on top of the other, were used as feeders. The feeder was placed 
on the cover of the watch glass, covered with a coverslip to keep the 
thrips from sticking to the feeder, and then saturated with the substance 
to be fed. The saturated feeder adhered fairly well to the cover of 
the cage. After the thrips were shaken into the cage, the cover was 
sealed to the watch glass by means of hot paraffin. This prevented the 
escape of the thrips and aided in maintaining the humidity. The cage 
was then placed cover downwards in the oven where a constant tempera- 
ture of 21 C. was maintained. Even though the cage was placed in 
an oven, the relative humidity remained high throughout the experiment 
for condensation occurred between the blotting paper and the bottom 
of the cage which it lined. The feeders were also damp when the cages 
were opened at the end of the experiment. There seemed to be sufficient 

♦Abstracted from "The utilization of carbohydrates and proteins by onion thrips, 
Thrips t abaci Lindeman," a thesis submitted to the faculty of Cornell University, Sep- 
tember 1932, by Burl Alva Slocum, University of Nanking. 



296 Phylum Ar thro p oda 

air within the cage to meet the needs of the insects for some of them lived 
as long as 25 days thus enclosed.* 

Order anoplura 

REARING HOG LICE ON MAN 

Laura Florence, N. Y. Homeopathic Medical College and Flower Hospital 

^"pHE hog louse, Haematopinus suis, is the largest of the lice affecting 
JL domestic animals. It is suitable for experimental work, because 
it is easily obtained and feeds readily on man. Its size and its habit of 
taking hold of any slender object placed in front of it lessen the difficulty 
of keeping it in confinement. 

On infested hogs, lice are readily found in the folds of the skin on the 
neck and jowl, within and at the base of the ears, on the under side of 
the legs, on the flanks, and in smaller numbers on the back, where they 
crawl under the scales in order to come in contact with the new skin. 
From these regions they may be collected with small forceps or with the 
fingers and placed in any easily handled receptacle to be taken to the 
laboratory. Without undue delay they should be transferred to small 
vials, approximately 5 cm. x 1 cm., containing some hog bristles and 
threads of gauze. Four to six lice should be placed in each vial. The 
mouth of the vial should be closed by tying over it two layers of gauze 
with a very thin layer of absorbent cotton between. These vials must 
be worn continuously under the clothing, so that the lice may be kept 
as near body temperature as possible. 

The captive lice are fed on the forearm, and should not be handled un- 
necessarily. In the vial they will be found attached to the threads and 
bristles. These should be withdrawn from the vial and placed on the 
arm. The lice will then move to the skin and may feed at once or move 
about more or less rapidly. The peculiar structure of the feet enables 
them to grasp the hairs on the arm. After the insertion of the stylets the 
insect holds itself in a more or less straight line and at an angle of 40 

♦Editor's Note: C. O. Eddy and W. H. Clarke (/. Econ. Ent. 23:704, 1930) report 
life history studies carried on with the onion thrips by the use of i-gram homeopathic 
vials, ?4 by 6 inch test tubes, absorbent cotton, insect-free seedling cotton leaves, and 
water. The females were confined separately in the homeopathic vials and a section of 
a fresh seedling cotton leaf placed with each of them every 24 hours. The used sections 
of leaves were removed from the vials, wrapped individually in moist absorbent cotton, 
and each placed in a sterile test tube. The open ends of both vials and test tubes were 
closed with absorbent cotton plugs. The leaves in the test tubes were removed daily and 
observed under a low power binocular microscope for emerged larvae. When a larva was 
found, it was removed from the leaf with a small brush and transferred to a fresh leaf 
in a small homeopathic vial where development was observed. Fresh leaves were supplied 
to all larvae when needed. Vials and test tubes containing the thrips were inserted 
slightly in the soil of soil tables in an out-door insectary. M.E.D. 



Anoplura 297 

to 45 with the arm. As the feeding progresses the body is gradually 
lowered, until it rests on the arm. Blood is seen passing into the ali- 
mentary canal in which a continuous peristalsis is evident. Feeding is 
continued until all feces and a drop of fresh blood have been ejected from 
the canal. The average length of a meal is from 8 to 12 minutes, but it 
may last from 20 to 30 minutes. At its close the mouthparts are with- 
drawn, apparently by a short jerk of the head. The lice should then 
be carefully removed from the arm with a small forceps and returned to 
their vial. They should be held by the legs and not by the body in order 
to avoid rupturing the distended alimentary canal. Newly hatched 
lice will feed readily and must be given at least four opportunities to 
feed in 24 hours until they reach maturity. Mature lice should be given 
two, and if possible three, opportunities to feed in 24 hours, since those 
exhausted by too long fasting will not feed on man. 

The unfed louse is of a grayish color and much wrinkled, while the 
fed louse has a highly refractive, smooth tegument, showing very clearly 
the areas of sclerotization. 

At every feeding the threads and bristles from each vial should be 
examined carefully for eggs. These, if present, are found attached to a 
thread or bristle which should be removed to a second vial. If worn 
continuously at body temperature and if fertile, the eggs will hatch in 
13 to 15 days. In the course of their development hog lice undergo 
three molts. Rearing in captivity has proved the cycle from egg to egg 
to occupy from 29 to 33 days. The life history, as we have observed it, 
may be summarized as follows: 

Time from laying to hatching of eggs 13 to 15 days 

First molt occurred after 5 to 6 days 

Second molt occurred after 4 days 

Third molt occurred after 4 to 5 days 

Sexual maturity occurred after 3 days 

Time of development from first stage larva to 

mature adult 16 to 18 days 

Temperature 35 C-> 

(continually next to body in vials) 

Number of feedings in 24 hours. 1 to 4 

Duration of cycle from egg to egg 29 to 33 days 

This method of keeping lice in captivity has proved satisfactory in 
investigations carried over a period of years, and large numbers of lice 
have been fed on the forearm without any harmful results. Egg-laying 
and molting have been observed and lice have been reared from the egg 
to maturity. All our attempts to rear a second generation of captive lice 
have failed. Eggs laid by reared females, kept in separate vials with 
males, have all quickly changed color and become shriveled, although 
the insects were seen in position for copulation a number of times. 



298 Phylum Arthropoda 

In the course of this study hog lice have been found to be the normal 
host of a symbiont, living in enlarged epithelial cells of the mid-intestine 
and passing from generation to generation through the egg. In artificially 
reared lice these symbionts tend to disappear, and the possibility of an 
intimate relationship between the symbiont and the blood ingested by the 
insect host may be the explanation of the impossibility of rearing hog 
lice elsewhere than on their natural host. 

Bibliography 

Florence, Laura, 192 i. The hog louse, Haematopinus suis Linne: its biology, 
anatomy, and histology. Cornell Univ. Agric. Exper. Sta. Mem. 51. 

1924- An intracellular symbiont of the hog louse. Amer. J. Trop. Med. 

4:397- 

Order hemiptera, Family scutelleridae 

NOTES ON REARING A SCUTELLERID 

H. M. Harris, Iowa State College 

THE geographical distribution of Acantholoma denticulata is limited 
by the range of its host plants, Ceanothus pubescens and the related 
C. ovatus. The species hibernates in the adult stage and usually may be 
taken at any season by sifting leaf mold from around the host plants. 
However, the bugs congregate in the corymbs at the time of seed-pod 
formation and may be collected in numbers most easily at this time by 
quietly approaching the plant and cupping the hands beneath the flower- 
heads. Adults mate and oviposit readily in captivity. Eggs normally 
are placed in the vegetable mold, but when deprived of this (as is desir- 
able in rearing studies) the females will oviposit on leaves or other 
surfaces. Almost any type of cage with provision for ventilation will 
suffice for the adults when fresh seed-heads of the host plant are avail- 
able for food. For the eggs and nymphs, however, small stender dishes 
serve best. It is essential to preserve proper moisture relations in the 
cage if success in rearing is to be achieved. A disc of absorbent paper 
(white is preferable) tightly fitted in the bottom of the dish and slightly 
moistened once or twice a day is all that is necessary. The paper must 
be renewed and the dishes cleaned regularly to hinder the growth of 
molds. Food for the young consists of seeds of Ceanothus. In nature 
the seeds must remain in the duff layer of soil for varying periods of 
time before the very durable outer coats are softened sufficiently to allow 
penetration by the feeding stylets of the young bugs. For use in feeding 
caged individuals it is sufficient to soak the seeds in water to soften 
them or to crush them mechanically. The insects readily feed on these 
soaked or crushed seeds. 



Pentatomidae 299 

It is worth pointing out that related bugs, particularly the Cydnidae 
and the Corimelaenidae, in many cases have habits similar to those of 
Acanthaloma denticulata and no doubt might be reared successfully by 
collection and use of the proper seeds as food. Likewise, in the Ly- 
gaeidae (Andre, 1935), cultures of almost any of the so-called milkweed 
bugs (Lygaeus and Oncopeltus) may be kept going easily and almost 
indefinitely simply by providing the caged insects with dried seeds or 
ripened seedpods of milkweeds for food, water for drink, and cotton in 
which they will readily oviposit. 

Bibliography 

Harris, H. M., and Andre, Floyd. 1934. Notes on the Biology of Acantholoma 
denticulata Stal (Hemiptera, Scutelleridae) . Ann. Ent. Soc. Amer., 27:5. 

Andre, Floyd. 1935. Notes on the Biology of Oncopeltus fasciatus (Dallas). 
Iowa State College J. of Sci. 9:73. 



Family pentatomidae 



A METHOD OF REARING FOUR SPECIES OF PLANT BUGS 

F. G. Mundlnger, New York State Agricultural Experiment Station 

THE species concerned here are Acrosternum hilar e, Euschistus 
euschistoides, E. variolarius, and E. tristigmiis. These species are 
common in central New York State from early spring until late fall and 
may be captured by sweeping grass or beating the bushes with a strong 
insect net. Some of the natural food plants are Viburnum aceri folium, 
Cornus racemosa, Vaccinium stamineum, and a species of Smilax, prob- 
ably 5. herbacea. Cucumber serves as an excellent supplementary food 
but only the firmer portions of this or any other fruit should be used 
where small nymphs are concerned. Screen-topped, glass jars, quart-size 
or smaller, partially filled with sterile, moist sand packed down to sup- 
port a small twig of the food plant, make good breeding and rearing jars. 
Daily inspection of these should be made in warm weather and wilted 
food material replaced by fresh. The same technique may be followed 
for the four species. 

I have not reared successive generations of these species. In this 
region these plant bugs appear to be single-brooded. I have caged reared 
specimens for some time but failed to secure any eggs. Each year I 
have begun my experiments with material captured in the field. 



300 Phylum Arthropoda 

PERILLUS BIOCULATUS* 

THE adults of Perillus come forth from hibernation as soon as the 
ground thaws out in the spring. By the time potato plants show 
above ground and the first potato beetles appear, Perillus may be found 
in the potato fields. Their first meal in the spring is sap from the potato 
plant, but after that their food is almost exclusively the body fluids of 
potato beetles, eggs, and larvae. 

For rearing Perillus the ordinary type of jelly glass was found to be 
a very convenient cage. One pair of bugs in a glass will take very kindly 
to this arrangement, and, when fed daily, will produce eggs quite as 
freely as in the field. After the bugs have been confined in the jars for 
two or three weeks they become very tame, rarely trying to fly when 
handled. The female bug will lay her eggs on the potato leaves when 
these are provided, but in the absence of these will lay eggs readily on 
the sides of the jar or on cheesecloth when it is supplied. As fast as the 
eggs are laid they may be removed to new jars for rearing. After the 
nymphs attain the 3rd instar it is best not to keep more than 6 or 8 in 
one jar. Unless plenty of food is available at all times the bugs may 
develop cannibalism. However, when the bugs are not overcrowded this 
difficulty rarely occurs. It was found necessary to clean the breeding 
jars frequently, especially when rearing nymphs on larvae of the potato 
beetle. 

The writer reared Perillus from egg to adult on nothing but mature 
beetles, but the beetles were always rendered helpless for the benefit of 
the nymphs in the 2nd and 3rd instars. The bugs should be fed once or 
twice a day, although they will get along if neglected for a day. More 
labor is necessary in rearing Perillus on adult beetles alone than when 
grubs are available, yet it has been done in order to rear a fall genera- 
tion of bugs after the potato beetle grubs have disappeared. By this 
method of rearing Perillus might no doubt be kept active and breeding 
during the winter months if proper greenhouse facilities were available. 

M. E. D. 

Family coreidar 

J 

CORIZUS HYALINUS AND C. SIDAE** 

THE writer has reared Corizus hyalinus through the whole course 
of its life history on a wild lettuce, Lactuca scariola, and has observed 
numerous adults, nymphs, and eggs on this plant in the field. Eggs of 

♦Abstracted from an article in the igth Rept. State Ent. of Minn. p. 50, 1922, by 
Harry H. Knight, Iowa State College. 

** Abstracted from an article in Ann. Ent. Soc. Amer. 21:189, 1928, by Philip A. 
Readio, Cornell University. 



Lygaeidae 301 

this species have also been found on a seed pod of velvet leaf, Abutilon 
theophrasti, and the young have been reared through to the 5th instar 
on this plant. Two successive generations have been reared in the 
summer. The insect has been reared from adult to adult in 17 days. 
The adult life is comparatively long, one adult female having been kept 
in confinement for 50 days. 

The writer has had difficulty in getting Corizus sidae to thrive on 
Sida, though it has been seen to feed on this plant when confined. It 
has been found as nymph and adult on the seed pods of velvet leaf, 
Abutilon theophrasti. Furthermore, the eggs have been laid on this 
plant in the laboratory and the nymphs reared through two instars before 
the cold weather put an end to the food supply. C. sidae has not been 
reared through its complete life cycle. Eggs were obtained from con- 
fined adults kept in a warm room during late October and early Novem- 
ber. These eggs hatched in from 10 to n days, but probably would 
have hatched in a shorter time out-of-doors in the summer. 

These rearings were conducted at Lawrence, Kansas, during late 
August and early September when the temperature was high, reaching 
the high eighties and nineties during the middle of the day. The insects 
were reared in an outdoor insectary, confined in glass stender dishes, 
and fed daily with pieces of the food plant.* 

M. E. D. 



Family 



LYGAEIDAE 



LYGAEIDAE 

F. M. Wadley, U. S. Bureau of Entomology and Plant Quarantine 

THE chinch bug, Blissus leucopterus, has given considerable difficulty 
in rearing. Cages must be tight because of the insect's habit of 
crowding itself into any possible crack. Humidity seems important 
with younger stages; very dry conditions are unfavorable, and free 
water causes bugs to stick to the cage. The species thrives at ordinary 
summer temperatures, but not under cool conditions. The chinch bug 
may be reared on bits of fresh food-plant put in daily, growing plants 
not being absolutely necessary. Seedlings of corn, wheat, and sorghum 
have been used with success ; corn has a greater tendency than the others 
to cause free water in the cages. Bits of crab-grass stalks have also been 
used, but are less desirable. 

♦Editor's Note: J. C. Hambleton, U. S. Bureau of Entomology and Plant Quarantine. 
reported in Ann. Ent. Soc. Amer. 2:272, 1909, that several broods of Corizus lateralis 
were reared to maturity on blossoms and young seed of Polygonum persicaria and on P. 
pennsylvanicum. Eggs were deposited on the latter. The adult forms fed freely on these 
plants in captivity. M. E. D. 



302 Phylum Arthropoda 

To obtain eggs, adults were confined in flat-bottomed vials about 
4 inches long and i inch in diameter, stoppered with cotton. A little 
ground litter such as occurs in the field, consisting of bits of soil and 
plant material, was put in, and pieces of fresh food-plant were supplied 
daily. This method gave fair results. 

Nymphs when confined with food in ordinary vials did not thrive; 
few matured, and growth periods seemed to be much lengthened as com- 
pared with those in the field. After trying several devices, one was hit 
upon which gave more nearly normal development. The i" x 4" vials 
mentioned were prepared with plaster casts about %" thick in the bottom. 
The eggs were placed in these vials in a little ground litter. The vials 
were kept upright and stoppered with cotton. The nymphs on hatching 
were kept in these vials during the first two instars, several in a vial, 
with fresh food supplied daily. With practice the 1st and 2nd instars 
could be readily distinguished; and by daily inspection those in the 2nd 
instar were removed to other vials soon after molting. After the 2nd 
instar, nymphs did not do so well in these vials, and were reared through 
later instars in little, individual, chimney cages on growing sorghum 
seedlings. These were made of 1" x 4" vials open at both ends. The 
vial was placed around the seedling, pushed well into the soil, and the 
soil inside carefully smoothed and tamped with a pencil. The top was 
closed with cotton. Several such cages may be placed on one flower-pot. 
Sorghum seedlings are favorable in cages because of their slow growth. 

The writer assisted Mr. F. B. Milliken in rearing the false chinch- 
bug, Nysius erkae* This work also presented some difficulties. The 
species feeds on a number of plants; small crucifers such as shepherd's- 
purse and pepper-grass are important early in the season, and a spurge, 
Euphorbia, of procumbent habit, is used in late summer. Adults were 
confined for egg-laying in small vials on potted food-plants. The vials 
were stoppered with cotton, which pinched a branch of the plant pro- 
jecting into the vial. Eggs were deposited in the cotton. Mr. Milliken 
had best success rearing nymphs individually in small muslin bags tied 
on the food plants. The bag was made of two pieces of muslin, perhaps 
2" wide and 3" long, stitched together with fine stitches around 3 sides. 
By careful examination molts could readily be found in these bags. 

Reference 
For the rearing of Lygaeus and Oncopeltus see p. 299. 
*/. Agric. Res. 13:571-578, 1918. 



Lygaeidae 303 

REARING METHODS FOR CHINCH BUGS, BLISSUS HIRTUS 

Kenneth E. Maxwell, Cornell University 

FOR a study of the length of nymphal instars, it was desirable to 
isolate individual nymphs and observe their moltings. A satisfactory 
cage for this work was a %" x 2%" round-bottom flint shell vial, of the 
same shape as biological laboratory test tubes. The tubes were stoppered 
with cotton plugs and supplied with the stems of grass plants for food. 
The food used was creeping bent grass, a variety in which the bugs breed 
very readily and which is commonly used for lawns and putting greens 
in the Northern United States. 

It was found necessary to replenish the food supply nearly every 
day because of the drying out of the grass. The grass furnished both 
food supply and moisture. Grass stems were cut at the base and im- 
mediately immersed in water, where they were kept until ready for 
use. Frequent moistening of the cotton plugs aided in maintaining a 
high humidity. Attempts to conserve the moisture content of the grass 
by the use of cork stoppers were not successful, due to the fact that 
evaporation from the food material saturated the air, and moisture con- 
densed on the glass. The insects, particularly the younger stages, fre- 
quently stuck to the wet surface, and suffered a high mortality. 

The nymphs undergo five molts, with considerable variation in the 
time required for each instar under different conditions. The shortest 
time required from egg to adult, under field laboratory conditions, was 
44 days, in contrast to the longest, which was 81 days. As many as 
9 chinch bugs have been successfully reared through all stages in a 
single tube, and a larger number may be used, depending on the food 
supply. 

Oviposition. For fecundity studies of individual females, single pairs 
were confined in the same type of cage as that used for life history 
studies. Copulation occurred frequently during the oviposition period, 
and some females oviposited daily for several consecutive days. There 
is considerable variation in the fecundity of individual females, and ovi- 
position fluctuated with the temperature and with the quantity of food 
material and moisture present. Oviposition decreased with low tempera- 
tures, and almost ceased when the prevailing temperature remained below 
70 F. It was desirable to maintain a high humidity, keeping the air 
as nearly saturated as possible without obtaining condensation on the 
sides of the cages. No difficulty was experienced with fungus killing the 
insects in the laboratory studies. 

When a female was ready to oviposit, she inserted her ovipositor be- 
neath the leaf sheath, and deposited the eggs, sometimes singly, but 
usually in groups. When the leaf sheath was pulled back, the eggs 



304 Phylum Arthropoda 

were found to lie side by side, frequently stuck together in a compact 
row. In such cases they could be removed in a group. 

Incubation. When first laid the eggs are opaque, pearly white or 
with an amber tint. They soon darken, take on a pink coloration, and 
become deep red prior to hatching. The incubation chambers consisted 
of glass tubes similar to those used for rearing, into the bottom of which 
had been poured a small quantity of plaster of Paris, and closed with 
cork or moist cotton stoppers. In these observation of hatching was as 
possible without removal of the stoppers. 

The length of the egg stage was largely dependent on prevailing tem- 
peratures, and the eggs hatched under a surprising range of moisture 
conditions. Eggs placed in dry containers with cotton plugs became 
shriveled in a few hours, and remained pale yellow for a number of 
days. A large percentage eventually hatched, requiring, however, a 
somewhat longer time to do so than those which were kept moist. Eggs 
which were kept for long periods in closed containers with moisture 
present, likewise remained viable until hatching. The emerging nymphs, 
however, floundered in the water film present, and soon perished. 

Bibliography 

Janes, Melvin J. 1935. Oviposition studies on the chinch bug, Blissus leucopterus 

Say. Ann. Ent. Soc. Amer. 28:109. 
Luginbill, Philip. 1922. Bionomics of the chinch bug. U. S. D. A. Bull. 1016. 
Shelford, V. E. 1932. An experimental and observational study of the chinch bug 

in relation to climate and weather. ///. Nat. Hist. Surv. Bull. 19:487. 



Family reduviidae 

J 

REDUVIUS PERSONATUS* 

A DULTS of Reduvius personatus, collected at a light trap and paired, 
l\ were confined in small cartons and were fed daily on houseflies 
which had been caught in a net and disabled before being introduced into 
the cages. Eggs were removed daily and placed in a salve box, % of an 
inch high by 1% inches in diameter. It was found desirable to line the 
box with a tightly fitting wad of heavy, unglazed paper, which served to 
give the bugs a foothold and also to absorb excess moisture from food and 
excrement. Later, in work involving humidity, % inch holes were 
punched in the lids, and these openings covered with coarse silk bolting 
cloth, so that the atmosphere could come into equilibrium readily with 
that of the rearing cabinet. 

The nymphs also were fed houseflies since there was a large supply 

♦Abstracted from an article in Ann. Ent. Soc. Amer. 24:19, 1931, by Philip A. Readio, 
Cornell University. 



Mesovcliidae 305 

available. However, later a change was made to the larvae of Tribolium 
confiisum, the confused flour beetle. This insect proved to be a very 
satisfactory food supply for Reduvius, since it could be reared in any 
numbers desired, and in a short time, on common flour. The greatest 
objection to its use was that occasionally a larva would crawl upon 
and kill or cripple a molting Reduvius. 

M. E. D. 



Family mesoveliidae 

HOW TO REAR MESOVELIA 

C. H. Hoffmann, U. S. Bureau of Entomology and Plant Quarantine 

A DULTS were collected and divided according to species, after which 
l\ they were isolated in finger bowls containing lake water and several 
pieces of decayed cat-tail leaf available for oviposition. The water was 
changed daily. Fruit flies or houseflies served as food. When a large 
number of eggs were laid in the stem provided, it was then removed to 
a petri dish and given a number that corresponded with that on a card. 
Permanent data were, of course, kept on cards. An eye dropper was 
used to change the water in the egg containers, and a dropper or two 
full of water was sufficient for the daily change. 

The little nymphs that hatched out were isolated in small stender 
dishes 1 inch deep and 2 inches in diameter and in some other stenders 
slightly larger. In the case of individual rearings, each stender was 
supplied with a single dropper full of water, a small piece of white card, 
and two adult fruit flies for food. The card served as a support for 
the small nymphs, and, being white, did not interfere in the search for 
molted skins. The bugs flourished with a daily change of water and a 
fresh supply of food every other day, the remains of the previous 
feeding being removed at this time. 

It is often difficult to find the newly hatched bugs, because they cling 
to the surface of the cat-tail stem. In changing the water of the egg 
cages daily, therefore, the fresh water was squirted directly on the 
surface of the stem and the young washed off into the clear water. These 
were then easily transferred to stenders by tilting the petri dish and 
placing the tip of a pair of curved forceps under the nymph, thus lifting 
it out together with a drop of water. Mass rearing in finger bowls was 
tried and found to be satisfactory. All rearings were carried on in a 
room which maintained a temperature of approximately 24 C. 

Houseflies served as good food but scarcity during the winter neces- 
sitated the finding of a food supply plentiful enough to care for many 



306 Phylum Arthropoda 

rearings. It was found that adult fruit flies filled the need, and of 
course they were easy to rear in quantities. The technique used might 
well be related, however, for it affords a simple way of capturing the 
adults to be used as food for winter rearings. 

A lamp chimney covered at one end with a piece of cheesecloth, fas- 
tened by means of a rubber band, was placed over a pint jar in which 
a banana and the fruit flies were placed. In a short time, the chimney 
was swarming with the adults. To capture a hundred or more specimens, 
a 4-inch vial was slipped through a piece of cardboard, in which a suffi- 
ciently large hole had been cut, and placed over the top of the chimney 
as soon as the cheesecloth was removed. If an electric light was placed 
above the vial, the desired number of flies was readily secured. When 
this number was obtained, the vial was quickly corked and a dark cover 
put over the chimney until the flies were scattered again. The chimney 
was then re-covered with cheesecloth. 

The adults in the vial could be killed by placing it over a hot radiator 
for a few minutes, or by applying the flame of a match to the bottom of 
the vial. In the latter case, the fruit flies could be forced to the bottom 
of the vial by sudden taps against something that would not break it. 
Heat kills the flies quickly and does not injure them for feeding pur- 
poses. 

Successive generations of two species of Mesovelia have been raised 
in the laboratory. The reared adults readily paired and deposited fertile 
eggs. 



Family nabidae 



A METHOD OF REARING TWO SPECIES OF NABIDAE 

F. G. Mundinger, N. Y. State Agricultural Experiment Station 

THE species here concerned are Nabis roseipennts and N . rufusculus. 
These insects are fairly common throughout New York State during 
the summer and may be captured by sweeping grass with an insect net. 
Screen-topped lantern globes placed over potted grass stalks or small 
raspberry plants make suitable breeding cages. Aphids are excellent 
food for the nabids. A petri dish containing a small green leaf and a 
drop or two of water serves well as a rearing cage for one or two nymphs. 
These cages should be cleaned daily, fresh leaves and drops of water sup- 
plied, and a few live aphids dropped in for the nymphs to feed upon. 
Since the Nabidae are predacious it is not advisable to place more than 
one or two in the same cage. 

I have not reared successive generations of these species. In this region 
the nabids appear to be single-brooded. 



Cimicidae 307 

Bibliography 
Mundinger, F. G. 1922. N. Y. State College of Forestry Tech. Publ. 16:149. 

Family cimicidae 

BREEDING AND REARING CIMEX LECTULARIUS 

R. M. Jones, Liquid Carbonic Corporation, Chicago, Illinois 

THE experiments were conducted under constant conditions of tem- 
perature and relative humidity. Incubating ovens were used to obtain 
the desired temperatures and relative humidities were kept constant by 
using saturated solutions of certain inorganic salts [see footnote on 
p. 307 ] . The eggs were obtained from bedbugs kept in small stender 
dishes in a glass battery jar under a constant condition of 27 C. and 
75% relative humidity. These bugs were fed every six days by being 
placed in wide-mouthed glass tubes and held against the forearm.* The 
females deposited their eggs on small circular pieces of paper toweling 
placed in the dishes for that purpose. At least once a day, or oftener 
when an experiment required an accurate record of the time the eggs 
were laid, the papers were taken out and the eggs removed with a 
camel's hair brush. They were then put in other jars under the same con- 
ditions and used for experimental work as soon as they were hatched. 

The following method was employed in rearing the bugs. Short pieces 
of 8 mm. by 40 mm. glass tubing were ground to a roughened surface on 
one end by applying to an emery wheel. On this end was then glued a 
small circular piece of 60 mesh bolting cloth, the other end being closed 
with the cap of a No. 000 gelatin capsule in which holes were punched to 
allow free circulation of air. One egg was placed in each tube and this 
furnished the permanent home for the bedbugs. The cages containing 
the eggs were kept under the conditions outlined above. After hatching, 
the nymphs were permitted to feed by holding the tubes against the 
wrist, no difficulty being experienced by the nymph in inserting the 
rostrum between the meshes of the bolting cloth. By using this method 
it was not necessary to remove the bugs from the cages until after they 
had reached the adult stage. The jars were aerated each day by fanning 
in fresh air with a piece of cardboard. 

In determining the length of time required for incubation the eggs 
were placed in 10 mm. by 50 mm. shell vials. These were then placed 
in 20 mm. by 80 mm. vials containing a saturated solution of the salt 

♦Editor's Note: Ezekiel Rivnay, in Ann. Ent. Soc. Amer. 23:758, 1930, gives a list 
of recorded hosts for bedbugs and on which they presumably may be fed for experimental 
purposes. This list includes: bat, cat, calf, dog, guinea pig, hare, mouse, rat, monkey, 
rabbit, duck, goose, hen, pigeon, sparrow, starling, and swallow. M. E. D. 



308 Phylum Arthropoda 

giving the desired relative humidity and tightly corked. The vials were 
also aerated once a day. 

To determine the length of life of the ist instar nymphs without 
food the nymphs were placed in No. ooo gelatin capsules in which many 
holes had previously been punched to allow a free circulation of air. 
The capsules were put in a small screen cage which was suspended in 
a pint Mason fruit jar containing a saturated solution of the salt giving 
the desired relative humidity. Aeration of the jars was performed daily 
as in the previous experiments. 

Bibliography 
Jones, R. M. 1930. Ann. Ent. Soc. Amer. 23:105. 

Family hydrometridae 

HYDROMETRA* 

"Hydrometras are easily kept in captivity and breed in aquaria, thriv- 
ing on a diet of flies and other small, soft-bodied insects. They are, 
therefore, ideal for observation." 



Family 



GERRIDAE 



THREE SPECIES OF GERRIDAE** 

GERRIDS are not easy to rear for the reason that they do not lend 
themselves readily to life in captivity. If good-sized containers are 
used and placed well back on the laboratory table, the specimens are 
less likely to injure themselves by dashing against the sides. If placed 
near the edge of the rearing table the striders will be disturbed each time 
someone passes. At feeding and observation time the bugs make frantic 
efforts to escape and in so doing continually butt against the sides of the 
breeding jar. Observations may not be made if the rearings are carried 
on in a large container, unless one has access to a movable-arm binocular. 
A recently captured female Trcpobates pictus laid a small mass of eggs 
on the underside of a small willow twig that was placed in the aquarium 
jar. Individuals reared in captivity and depositing eggs every day will 
usually fail to deposit eggs the first day after they are placed in a new 
breeding jar. 

* From an article in Ent. Amer. 7:87, 1926, by J. R. De La Torre Bueno, Tucson, 
Arizona. 

** Abstracted from an article in Ann. Ent. Soc. Amer. 17:419, 1924, by William E. 
Hoffmann, Lingnan University. 



Veliidae 309 

Gerris buenoi laid eggs in the laboratory on sticks, stems, bark, and 
other floating material that was supplied. With aquaria large enough to 
accommodate growing plants more satisfactory studies could be made. 

Limnoporus rufoscutellatus, like the other members of the family 
Gerridae, is predacious, feeding upon insects to be found on the surface 
of the water. The adults prefer to cling to vegetation or other support 
most of the time. The nymphs seem to be more independent of sup- 
ports, as they were reared in jelly glasses with no supporting surface 
afforded. 

The rearing methods were essentially the same for the three species. 
Jelly glasses were used for the most part as containers, while the food 
consisted of flies, leafhoppers, and other soft-bodied insects. The jelly 
glasses were half filled with water. Sand was not used in the containers. 
Care had to be taken not to disturb the nymphs. When disturbed they 
would make frantic efforts to escape and would often become water- 
logged. When the body pile becomes thoroughly wet, the bugs will drown 
unless removed to an aquarium containing only wet sand. After the 
body is again clean and dry and the bug has recovered from the weaken- 
ing effects of its struggles, it may be returned to the aquarium containing 
water. Gerris remigis and G. marginatum were kept under the same con- 
ditions with about the same degree of success.* 

M. E. D. 

Family veliidae 

WINTER FOOD FOR WATERBUGS IN AQUARIA** 

FLIES may be collected within buildings in limited numbers all 
winter. Bruchus and Tribolium adults may be secured in quantities 
from places where there are heavy infestations, kept in a large container 
with their respective foods, and used as needed. Tenebrio molitor larvae 
and Drosophila are satisfactory. 

However, the most successful food in the experience of the writer has 
been cockroach nymphs. They are easy to secure, easy to handle, and 

♦Editor's Note: C. F. Curtis Riley, of the University of Manitoba, reported in Ent. 
News 33:86, 1922, on the rearing of Gerris remigis and G. marginatus. Both will feed on 
a variety of insect food, such as the pupae and adults of Culex, small and large species 
of Tipulid flies, Syrphid flies, Musca domestica, Chironomus, Tabanus, and Drosophila. 
G. remigis is a more vigorous and daring feeder than is G. marginatus and has been ob- 
served to feed on Notonecta undulata, Chrysopa, Calopteryx maculata, Hetaerina ameri- 
cana, and Arctocorixa. Both species have at times been noticed feeding on the soft parts 
of banana fruit and on the inner soft parts of the skin in the absence of other food. 
During confinement both species will suck the juices of freshly killed snails, Physa and 
Planorbis. and also small pieces of fresh beef. M. E. D. 

** Abstracted from an article in Bull. Brooklyn Ent. Soc. 19:149, 1924- by William E. 
Hoffmann, Ungnan University. 



310 Phylum Arthropoda 

they produce healthy bugs. Five species of Microvelia and two species 
of Velia have been successfully reared on a straight diet of cockroaches. 
Several specimens of Curicta have been carried from the 3rd instar to 
the adult on this diet while adult Nepa, Curicta, Ranatra, Velia, and 
Microvelia have been kept through the winter. 

Immediately upon hatching the nymphs are large enough to make a 
meal for Microvelia or Velia, while those a week or two old serve nicely 
for the larger waterbugs. If one has access to a place infested with cock- 
roaches it is a simple matter to catch nymphs. A space on the floor is 
cleared, a few bits of food placed there and covered with a piece of 
cardboard or beaverboard. After the lights have been turned off a few 
minutes they may be turned on, the cardboard lifted, and dozens of the 
nymphs killed or crippled with a fly swatter. The clean floor makes 
them readily visible and they are easily picked up with a forceps. Often 
they will come in numbers while the lights are on and even during the 
day. The scattering of food particles or even sprinkling of water on the 
floor will attract them. They may also be trapped by placing a heavy 
paper funnel in a deep bottle, but specimens caught in this manner are 
unsatisfactory for they get wet and for that reason sink through the 
surface film of the water. Microvelia and Velia will catch living organ- 
isms beneath the surface film, but they do not care for flies, cockroaches, 
or other similar food that does not rest upon the surface. Because of this 
it is preferable to kill the nymphs just before feeding time. If placed on 
their backs they are not likely to sink and in this position the parts easiest 
to pierce are uppermost. 

To insure a continuous supply of food, adult cockroaches bearing egg- 
cases may be trapped and the cases removed to containers with damp 
blotting paper or other damp material on the bottom. Corked bottles 
or glass containers with rather tight-fitting lids will serve the purpose. 
Upon hatching the nymphs may be reared by giving them a piece of 
apple every few days. 

j. g. N. 

VELIA WATSONI* 

FIFTH-STAGE nymphs and adults of Velia watsoni, taken among the 
roots of smartweed and other weeds and grasses on the moist banks 
of pools in the bed of a small stream, were placed in tin boxes with damp 
vegetation. Little trouble was experienced in keeping the bugs in cap- 
tivity and in rearing them. 

Mating in captivity has taken place during every month of the year, 
but has been more frequent during the warmer part of the season. 

♦Abstracted from an article in Canad. Ent. 57:107, 1925, by William E. Hoffmann, 
Lingnan University. 



Saldidae 3 11 

In the aquarium they deposit eggs on the side of the glass container 
and on rocks, as well as on floating objects. Eggs have been deposited 
in the laboratory during every month but December, with the peak of 
production occurring during August and September. 

This species is predacious and cannibalistic. The adults will kill the 
nymphs and the nymphs will kill each other. Consequently it is necessary 
to remove the adults to new containers as fast as the eggs are ready 
to hatch. Velia brachialis was found to differ in this respect, for a num- 
ber of nymphs, or even a number of both nymphs and adults, have been 
kept in the same aquarium with little danger of cannibalism developing. 

Velias, like Microvelias, feed upon the very small animals swimming 
beneath the surface film as well as upon food particles on the surface. 
The Velias are very leisurely in their feeding activities. They feed on 
living, crippled, or dead insects placed on the surface of the water. The 
adults do well on any insect food, while the nymphs thrive better on 
cockroach nymphs than upon any other food. Many nymphs were reared 
from hatching to maturity on a straight diet of cockroaches. The adults 
have been fed on flies, nymphs and adults of cicadellids, cercopids and 
mirids, grasshopper nymphs, adult Tribolium, Bruchus, Dermestes, and 
larval forms such as Tenebrio, Mediterranean flour moth, and various 
caterpillars. 

M. E. D. 

Family saldidae 

SALDULA MAJOR AND S. PALLIPES* 

NYMPHS of Saldula major, captured in June, transformed into 
adults and mated by June 18. Newly laid eggs were found at the 
base of a blade of grass on June 21. These hatched on July 3. The 
nymphs liked to stay hidden most of the time, but would come out 
readily to feed. 

Nymphs of Saldula pallipes (?), captured on June 1, became adults 
and laid eggs before June 23. These were thrust in the stems and blades 
of grass growing in the jars in which the saldids were confined. 

In all the rearings dead flies were used as food as well as other soft- 
bodied insects, chiefly mirids and cicadellids, which were usually easy 
to obtain in large numbers either by sweeping or at light at night. 

M. E. D. 

♦Abstracted from an article in Kan. Univ. Sci. Bull. 14:301, 1922, by Grace Olive 
Wiley. 



312 Phylum Arthropoda 

Family notonectidae 

REARING NOTONECTIDAE* 

Buenoa mar gar it aec a, B. scimitra, B. elegans, Plea striola, and No- 
tonecta undulata, all common species in Kansas, have been reared from 
egg to adult. Successive generations have been raised in the case of several 
of these species. 

Since about 1895 various attempts have been made to rear the noto- 
nectids, but usually they have been only partially successful. The diffi- 
culties have been in establishing conditions in the aquarium duplicating 
those of the ponds or natural habitat of these insects. At least three 
important factors have been at fault: the oxygen content of the water, 
the condition of the surface film, and the food supply. 

In attempting to rear the early stages of Buenoa it was found that few 
nymphs ever reached the 2nd instar in the stender dishes used. It even 
took the eggs longer to hatch there than out in the ponds. Frequently 
changing the water in the stender dishes did little good. When it was not 
so changed, a scum formed over the surface after one or two days. The 
bugs died more quickly. The daily removal of this scum helped the 
situation but did not remedy it. It occurred that the trouble probably 
was due to oxygen deficiency and the greater difficulty in breaking the 
surface film when it was covered with scum. Air forced through the water 
with a large pipette almost immediately revived the dying bugs, but the 
relief was only temporary. Then a device first described in 19 10 ** for 
continuously aerating an aquarium was used. It was a success, and 
must be credited for the life histories of the genus Buenoa. 

Another difficulty was the food supply. Ostracods and such minute 
organisms as the Buenoae and Pleae usually fed upon were inconvenient 
to collect. Other workers had reported mosquito wrigglers as good food 
for notonectids. So during most of the season mosquito eggs were col- 
lected each morning from one or two tubs of rainwater and kept in a 
smaller vessel until next day when they hatched and served well as 
food for all species. A barrel of rainwater near the insectary supplied 
larger mosquito larvae for the more mature nymphs and adults of 
Notonecta and Buenoa. But even tiny Plea will not hesitate at times to 
attack almost full grown mosquito wrigglers. 

Other factors than those just mentioned should be considered. The 
laboratory in which the rearing work was done was in a basement which 
affected the temperatures and lighting, and it was artificially heated 
during the latter part of the season. All aquarium jars except a few kept 

♦Abstracted from an article in Ann. Ent. Soc. Amer. 19:93, 1926, by Clarence O. 
Bare, Sanford, Florida. 

** Schaeffer, A. A. 1910. Science 31:955. 



Nepidae 3 T 3 

as controls were constantly aerated, and each was supplied with sprigs 
of Ceratophyllum. All aquaria were filled with pond water. Careful 
regulation of the air was necessary to prevent too vigorous bubbling. 
With strict attention to details Notonecta and Plea were easily reared, 
but several attempts were necessary to get the full life story for the 
Buenoae. The mortality due to molting was considerable, as must also 
be true in the ponds; and nematodes seemed to cause the death of many. 

M. E. D. 

Family naucoridae 

PELOCORIS CAROLINENSIS* 

A LARGE number of rearings were started in April when eggs were 
laid, attached to a sprig of Nitella. The incubation period varied 
from 32 to 45 days with the majority requiring 39 to 40 days. By the 
time the eggs were ready to hatch the plant sprigs to which they were 
attached were dead and in some cases in a state of disintegration. 

Since Pelocoris caroUnensis is fiercely predacious it was necessary to 
isolate each newly hatched nymph. Each specimen was placed in a tall 
stender dish or jelly glass half full of water and supplied with a sprig 
of Nitella. Mosquito wrigglers, Chironomus larvae, corixids, and Ento- 
mostraca were given as food and the water was replaced by fresh pond 
water at frequent intervals. There was something grievously wrong with 
the rearing technique for only nine specimens were reared to the adult 
stage out of 134 isolations. Several females mated and laid eggs. 

M. E. D. 

Family nepidae 

CURICTA DRAKEI** 

SEVERAL pairs of adult Curicta drakei were collected and the pairs 
placed in separate glasses. Small glass containers were used with 
gauze tied over the tops. Sand and a few pieces of water plants were 
placed in the bottoms and made rather wet. 

Like Nepa, Curicta is a mud-loving bug. When the adults were given 
mud, rotten wood, decayed vegetation, and live water plants, the mud 
was always chosen for the deposition of the eggs. Both nymphs and 
adults are fond of getting out of the water and lying close to the ground, 
where they are hardly discernible. 

* Abstracted from an article in Bull. Brooklyn Ent. Soc. 22:77, 1927. by H. B. Hunger- 
ford, University of Kansas. 

** Abstracted from an article in Kan. Univ. Sci. Bull. 14:507, 1922, and one in Ent. 
News 35:324, 1924, by Grace Olive Wiley. 



314 Phylum Ar thro poda 

Only occasionally are these bugs cannibalistic. This may happen 
when no other food is available. Sometimes when hard pressed for food 
both young and adults have been observed to feed upon the eggs that 
were found in the water. They like small notonectids, corixids, small 
carabids, freshwater shrimps, and such. They refused small minnows, 
however. It is not uncommon to see three feeding quietly on one shrimp 
or two feeding on one small beetle. In the rearing work all sorts of 
insects were used as food, such as grasshoppers; stink-bugs; various 
species of beetles including blister beetles, flies, mealworms, and small 
snout beetles; membracids; and mosquito larvae, of which they are 
very fond. It was a problem to procure food for these insects during 
the winter, as they were kept in a warm room and were more or less 
active. Many times when the bugs were hungry and no insect food was 
available they were fed on small bits of raw beef. 

Adults readily mated and laid eggs in captivity. Curicta adults were 
kept over the winter and the following summer a number of these insects 
laid eggs from which young were again reared. 

M. E. D. 
References 
For the feeding of Nepa, Ranatra, and Curicta see p. 310. 

Family gelastocoridae 

GELASTOCORIS OCULATUS* 

TALL stenders, or staining jars, of glass, about the size of jelly 
glasses, were used as containers. In each of these was placed an 
inch of sand or soil that had been sterilized by heat. The paired adults 
were confined in low stenders of various sizes, and the sand searched 
every day for eggs. The young were isolated in the tall stenders as soon 
as they hatched, for they are cannibalistically inclined. The sand was 
moistened each day, and the jars were covered with ground glass covers. 

The insects were fed houseflies, oscinid flies, cicadellids, and many 
other small insects taken in sweeping the grass. Each day the dead car- 
casses were cleaned out of the rearing jars and freshly killed insects in- 
serted. Nymphs and adults of Gelastocoris pounce upon their prey, 
which appears to consist of almost any sort of insect they can capture, 
from a grouse locust to a lacebug. 

Mortality in captivity was very high and indicates that some essential 
factor of their natural habitat was lacking. Mortality was greatest in 

♦Abstracted from an article in Kan. Univ. Sri. Bull. 14:145, 1922, by H. B. 
Hungerford, University of Kansas. 



Cercopidae 315 

the 1 st stage and the nymphs usually succumbed on the date when molt- 
ing might be expected to occur. 



M. E. D. 



Order homoptera, Family cercopidae 

REARING CERCOPIDAE 

Kathleen C. Doerlng, University of Kansas 

SPITTLE insects, like all sucking insects which are plant feeders, are 
difficult to rear. It is very necessary that they have healthy, suc- 
culent plant tissue upon which to feed in order to get sufficient nourish- 
ment for their proper development. In addition, spittle bugs need vast 
quantities of plant juice in order to form their spittle masses. Normally, 
in the field, spittle nymphs perhaps do not move around to any great 
extent. In the laboratory, however, they are apt to be exceedingly rest- 
less, especially the tiny first instars. The restlessness of the nymphs is 
usually caused by disturbance of them in their spittle mass, or by lack 
of juice in the plant. 

In bringing spittle insects into the laboratory great care must be taken 
not to lose them enroute. The most satisfactory method for the author 
was to cut the entire plant stalk without disturbing the insects in the 
spittle mass and then to wrap the stem tightly in a newspaper. At the 
laboratory they should be transferred to growing young plants by means 
of a camel's hair paint brush. They ramble aimlessly over the plant 
for some time but eventually settle down when they find a favorable 
feeding spot. They should be watched carefully during this wandering 
period for very frequently they fall off the plant onto the dirt beneath 
where they sometimes are not able to regain their feet. Spittle bugs dry 
up quickly when not living in their spittle mass. When once settled 
upon the plant, and if the plant is growing, they give little trouble. 

Certain precautions should be noted in regard to the care of the host 
plants. In the first place if more than one host plant is found for the 
insect the plant chosen for rearing should be the one that lends itself best 
to transplanting and for which it is easiest to get small, seedling, or tender 
plants. For Lepyronia quadrangularis which has some sixty-two host 
plants, small plants of the common ragweed, Ambrosia artemisijolia, 
proved most satisfactory. Secondly it was found that cuttings of plants 
placed in water are unsatisfactory apparently because there is not enough 
plant juice present to supply the amount of fluid needed to make the 
spittle masses. Small, tender plants should be planted in flower pots. 
The nymphs are then confined on the plants under lamp chimneys. Over 



316 



Phylum Arthropoda 



the top of the lamp chimney is placed a covering of cheesecloth. For 
the tiny ist instars it was even necessary to double the cloth to keep 
the nymphs from straying away from the plant. It is necessary to use 
lamp chimneys for the first instar or two because they are so small at 
this time and so restless that they become lost easily in any larger 
space. For later stages wire cages would be better because the plants 
would do better in less cramped quarters and no difficulty is likely to be 
experienced in keeping the larger nymphs under observation. 

ARTIFICIAL FEEDING OF LEAFHOPPERS* 

A METHOD has been devised whereby a reasonably large number of 
leafhoppers may be caged and fed simultaneously upon a nutrient 
solution. It consists (Fig. 65) of a shallow "saucer" to which had been 

sealed a vertical L tube. This 
vessel is capped with a mesentery 
membrane of the type recom- 
mended by Carter.** The solution 
is added to the feeding apparatus 
until the liquid is in contact with 
the entire surface of the membrane 
by means of the side arm, which is 
corked to prevent contamination. 
This feeding vessel may be washed 
and sterilized in alcohol without 
removing the membrane or impair- 
ing its efficiency. 

For use with this feeding dish, 
a cage made of a 3-inch cylinder of 
1 % -inch glass tubing, capped at 
both ends with a fine open-mesh cloth such as georgette or scrim, is 
employed. The upper end of the cloth capping has a small opening for 
admission of the test insects, which is at other times closed by a cotton 
plug. The cage containing the test insects is placed upright upon the 
membrane surface over the solution. 

"With this arrangement, feeding will begin almost at once and freely 
continue as long as the insect lives. No evidence of unwillingness or in- 
ability of the leafhopper to locate the solution or to feed upon it was 
found. Furthermore with this arrangement it was possible to transfer 
feeding insects from one solution to another without handling them, 

♦Abstracted from an article in Science 79:346, 1934, by R. A. Fulton and J. C. 
Chamberlin, U. S. Bureau of Entomology and Plant Quarantine. 

** Animal mesentery sold under the name of "fish skin" at a drug store. /. Agric 
Res. 34:449, 1927, and Phytopathology 18:246, 192S. 




Fig. 65. — -Apparatus devised for simul- 
taneously feeding large numbers of homop- 
terous insects on a nutrient solution. 



Cicadellidae 317 

merely by lifting the cage from one saucer to another. From 25 to 50 
test beet leafhoppers may be confined in this cage without apparent over- 
crowding." 

j. G. N. 

Family cicadellidae 

THE BEET LEAFHOPPER, EUTETTIX TENELLUS* 

THE adults were confined in a cylindrical cage (12 by 8 inches) with 
top and sides covered with lawn, except for a glass plate (10 by 5 
inches) through which observations were made. The bottom of each 
cage was covered with denim fastened with a loop of copper wire. The 
leaves of a sugar beet with the base of the petioles wrapped in cotton 
projected through two central intersecting incisions in the denim. The 
denim rested against 2 inches of dry sand covering the surface of the soil 
in a 1 o-inch pot. The hoppers might be transferred rapidly to another 
potted sugar beet by blowing a breath of air through the sides and by 
jarring the cage, causing the insects to change their resting place from 
the foliage to the cloth; the cage was then lifted so that the leaves pulled 
through the incisions, leaving the bugs in captivity. This removal of the 
cage from the potted beet was performed in a dark chamber provided 
with a glass plate, outside of which was a 50-watt electric lamp covered 
with a shade, so that any specimens which perchance remained on the 
plant, resting between the petioles, were attracted to the light after the 
cage was removed. The glass of each cage faced to the north in the 
field. Each pot was placed in a saucer and the saucers were watered 
daily during hot weather. To prevent ants from entering the cage, the 
sides of the saucer were smeared with tanglefoot. 

M. E. D. 



Family chermidae 



CULTURE METHODS FOR THE POTATO PSYLLID 

George F. Knowlton, Utah Agricultural Experiment Station 

THE potato psyllid, Paratrioza cockerelli, yields readily to domestica- 
tion in the laboratory, when suitable cultural methods are used. 
Collecting Methods. A colony of potato psyllids may be readily estab- 
lished by collecting large nymphs upon potato or matrimony vine leaves, 
and then transferring them to vigorous young potato plants in the labora- 

*Abstracted from an article in Univ. of Calif. Pub. in Ent. 5:37. 1930, by Henry 
H. P. Severin, University of California. See original publication for illustrations of 
equipment. 



318 Phylum Arthropoda 

tory. Another method of establishing a colony is to collect the adult 
psyllids by "sweeping" infested plants with an insect net equipped with 
a heavy gauze or sheeting bag. For removal to the laboratory the adult 
psyllids may be placed in a temporary gauze cage, together with portions 
of a succulent plant, or inside of celluloid cages placed over young potato 
plants. In the laboratory, the adults may be transferred from the field 
cages to the rearing cages by means of a common aspirator. [See p. 46.] 
Care should be used to prevent aphids from entering the rearing cages 
with the psyllids or upon host plants. Plants upon which potato psyllid 
eggs have been laid may be potted and moved to the laboratory, if desired. 
A high nymphal mortality usually occurs when the leaves are picked off 
and become dry before the eggs hatch or when the egg stipes are shaved 
from the leaves and the eggs placed upon moist blotting paper inside of 
petri dishes. 

Cages. Several types of cages have been found to be suitable for the 
rearing and handling of potato psyllids: 

1. Large gauze cages, built to the width of laboratory windows, and about 20 inches 

deep, were found to be excellent for rearing potato psyllids. West windows 
were best in the winter and shaded east basement windows during the hot 
summer weather. 

2. Medium to large gauze cages with a wooden bottom and frame work. Cages 12 

to 18 inches wide, 16 to 24 inches long, and 16 to 20 inches deep were well 
adapted to rearing the psyllids. The front end consisted of a celluloid obser- 
vation sheet which covered approximately the upper Ys to V2 of the frame; 
to this was cemented (with acetone) a gauze flap extending approximately 2 
inches beyond the margins of the end. This gauze flap serves as a door, being 
fastened snugly with thumb tacks, except when potted plants are to enter 
or leave the cage. Small slits, which may be kept plugged with cotton, 
admit the 6 to 8 mm. aspirator, for capturing adult psyllids, and the glass 
tube, used with a funnel to water the host plants. 

3. Other suitable breeding cages consisted of round or square gauze cages, with a 

glass or celluloid panel and with a round opening in the bottom to fit upon a 
flower pot containing a host plant. 

4. Celluloid cages of various sizes to fit 6- to 8-inch pots. Cages fitted with a top 

and two side ventilators of gauze usually were satisfactory, but moisture 
would occasionally collect, resulting in the drowning of adult psyllids. 

5. Small clip cages, made of 4-dram homeopathic vials, with the closed end cut off, 

flanged, and covered with gauze, made satisfactory cages for one to several 
psyllids. These cages were held in place by means of a piano wire spring, 
a flat metal disc covering the top. To prevent the death of contained portions 
of the leaf, it was necessary to move such cages daily. 

6. A similar cylindrical cage, made of fine screen wire or celluloid and set into a 

small, thin, padded board, with a matching, padded board to fit on the 
opposite side of the leaf, avoided this difficulty. Such cages were held together, 
without excessive pressure, by means of rubber bands around the ends of the 
small boards. (Cages 5 and 6 are similar to those used on beet leaf hopper 
investigations.) 

7. Individual nymphs may be reared, when under constant observation, by placing 



Chermidae 319 

but one to a leaf and only two or three upon a small plant. If disturbed the 
nymphs may move around. 

Nymphal mortality was somewhat higher in the small than in the 
larger cages. A high mortality often occurred at the first and second 
molts, where conditions were for any reason unfavorable. 

Optimum conditions for egg-laying and nymphal development seem to 
occur between 70 ° and 75 ° F., with slight temperature fluctuations. The 
eggs, which are at the extreme end of a short stipe, are laid principally 
upon the younger, apical potato leaves. Females have a rather long 
oviposition period, and nymphs and adults of all stages may be had 
by adding a new potted potato plant to the breeding cage about once or 
twice each week. Where caged in pairs, it may be necessary to replace 
males that die in order to maintain the fertility of the eggs. Adults 
reared in cages appear to live longer in captivity, and to be less excitable 
than do wild adults collected out of doors. The mortality of freshly 
captured wild adults is sometimes heavy, especially if they are placed 
in small cages during hot weather. 

When the nymphs become excessively abundant, it may be necessary 
to kill or to remove part of the insects to save the plants. As many as 
2,000 to 3,000 nymphs, in addition to adults, have occasionally been 
found upon medium sized potato plants under both field and laboratory 
conditions. 

Greenhouse rearings are usually successful during the fall, winter, and 
spring months, but the psyllids have difficulty surviving during the sum- 
mer if temperatures become excessively high. Nymphs and adults have 
been observed to survive when the potato plants upon which they were 
feeding were destroyed by frost. 

In order to have vigorous potato plants at all seasons of the year for 
rearing potato psyllids, it sometimes becomes necessary, in order to induce 
growth, to use chemicals to break the rest period upon potato tubers to 
be used for seed. [See footnote on p. 328.] Tomato plants, and some 
other solanaceous hosts, may be used to raise psyllids in smaller numbers. 

Biological Control. Potato psyllid colonies may be lost by allowing 
heavy aphid infestations to develop upon and destroy the host plants. 
Adult and larval ladybird beetles, predacious Hemiptera such as damsel 
bugs and big-eyed bugs, lacewing fly larvae, syrphid larvae, spiders, and 
other predators may attack laboratory colonies of psyllids, unless the 
cages are kept free from them. No insect parasites have been noted in 
the area under observation, but when kept under humid conditions a few 
adults have been found which appear to have been destroyed by fungus. 



320 Phylum Arthropoda 

Family aphididae 

A USEFUL CAGE FOR REARING SMALL INSECTS ON 

GROWING PLANTS* 

IN THE rearing of aphids and their parasites and scale insects for the 
past two years, the writer has found a specially designed cage of sheet 
celluloid very satisfactory. Any cage for rearing these insects must allow 
for a free circulation of air, permit the entrance of light, and not cause 
a concentration of heat or moisture. Still another feature bears consider- 
able importance when dealing with small insects, and that is the absence 
of any cracks or niches in which the insect may hide or escape. 

The simplest and most suitable form of cage is the cylindrical type 
made by bending together the edges of a rectangular piece of celluloid and 
sealing them with 95% alcohol. Ventilation holes of any size and posi- 
tion on the sheet, may be cut in before bending. After the edges are 
sealed together and the cylinder is formed, the ventilation openings and 
one end opening are covered with fine cheesecloth or voile shellacked on 
the celluloid around the edges of the openings. The cage is now ready 
to be placed over the plant, usually a small one in a pot. In order to 
hold the cage securely in position and provide a smooth white surface on 
the bottom of the enclosure about the plant, melted paraffin is poured on 
the soil around the plant, and the open end of the cage set down into 
it after the edges have first been given a thin coat of vaseline to prevent 
the adherence of the paraffin. Thus treated the cage may be lifted 
free when the paraffin cools, leaving a smooth tight groove into which 
the edges just fit and prevent the escape of any insect when the cage 
is in place. Entry for the introduction or removal of material is easily 
made by tearing back a corner of the cloth ventilator, or lifting up the 
cage from its paraffin base. 

Sheet celluloid is a very satisfactory material with which to construct 
small cages for a portion of a plant like a small twig, or a part of the sur- 
face of a leaf or fruit on which it is desirable to confine small sedentary 
insects such as scales and aphids. In these small cages, ventilation may 
be secured by punching small holes in the celluloid with a fine needle. 
Shellac was found useful in joining together the sharp edges of the cage 
and sticking it to the plant surface. This material soon drys, with a 
hard surface so that it will not entangle the insects, and yet remains 
soft enough to prevent cracking and breaking apart. 

Cages constructed from sheet celluloid are as transparent as glass, and 
do not "sweat" like the glass cages. They may be made any size 

♦Abstracted from an article in Ohio J. of Set. 23:201, 1923, by E. A. Hartley, 
N. Y. State College oj Forestry. 



Aphididae 321 

and shape desired. They may be ventilated by cutting any number and 
size of openings in the side. They are very neat and smooth within, 
making it easy to observe specimens at all times. 

J. G. N. 

THE NASTURTIUM APHID, APHIS RUMICIS 

H. H. Shepard, University of Minnesota 

KPHIS rumicis is used more as a test insect for determining the relative 
l\ effectiveness of contact insecticides than for any other purpose. It is 
the common black aphid, or plant louse, infesting nasturtium (Tro- 
paeolum ma jus) and many other plants, and is known as the bean, dock, 
or nasturtium aphid. Although this aphid is reported from many woody 
and herbaceous plants, some of these do not seem to furnish suitable 
food, for after a few generations the aphid leaves them for more favor- 
able locations. In laboratory work in this country it has been reared 
upon nasturtium plants, whereas in the work of Davidson and of Tatters- 
field in England, the broad bean (Vicia jaba) was employed as its food. 
There is some confusion regarding the number of species of aphids in- 
cluded under the name A. rumicis (Franssen, 1927, and others). Hors- 
fall has pointed out that there are two distinct types of life cycle; one 
with woody shrubs as primary food plants on which the eggs are laid and 
the first generations develop in the spring, the secondary food plants 
being herbaceous; the other with primary food plants such as Cheno- 
podium album and Rumex, and these and other herbaceous plants as 
secondary food plants. 

There are several reasons why A. rumicis is to be preferred for the 
purpose mentioned. The individuals are black and show up well on a 
light background, resulting in less eye strain after counting large num- 
bers of them than if a green species were employed. A. rumicis is short 
legged and less likely to be injured in handling than the other species 
which are long legged and more awkward. It moves more slowly when 
disturbed by irritating chemicals, and hence is easier to keep within 
artificial barriers. 

This species is easily reared in the greenhouse, reproduction being con- 
tinuous throughout the year if favorable conditions are maintained. 
Colonies may be started easily on young nasturtium plants by trans- 
ferring to them a few aphids. The most important consideration is the 
growth and condition of the nasturtium plants. The seeds of the dwarf 
nasturtium, soaked in tepid water for an hour or so, should be planted 
in rich black loam. For convenience in handling it is customary to plant 
several seeds in a 3- or 4-inch flower pot and set the pots closely on the 
surface of the soil in the greenhouse bench. However, after the plants 
become of medium size the roots extend into the soil of the bench and are 



322 Phylum Arthropoda 

broken if the pots are moved, the plants wilting as a result. In order 
that the soil be kept fairly moist without becoming soggy; watering should 
be more frequent in sunny or warm weather and less so in cool, damp 
weather. When young and well watered, the plants have a very succulent 
growth upon which aphids do well. On the other hand very young 
plants of but two or three leaves may be injured by too heavy an infesta- 
tion. It is well to plant about twice a week so the aphid population may 
continually expand to plants of suitable size. It is also well to eliminate 
the older, heavily infested plants gradually so the population will not 
become too great and overrun the young plants excessively. The older 
plants are less succulent and less attractive to migrating aphids ; colonies 
developing upon them produce increased numbers of winged migratory 
females. Furthermore, the older colonies are likely to become so para- 
sitized by hymenopterous parasites that the entire aphid culture is in 
danger of being affected. 

Other factors than moisture may influence the physiological activity of 
the plant, thereby affecting the nature of the cell sap upon which the 
aphids feed. In the heat of summer the greenhouse should be kept as 
cool as possible and the plants shaded with cheesecloth or white-wash on 
the glass overhead. In midwinter in the north it is difficult to keep aphids 
in health because of the reduction of available daylight. 

In his tests of the toxicity of contact insecticides, Richardson used 
undisturbed colonies of Aphis rumicis by cutting the leaves and plants 
bearing them and inserting the stems through two-hole rubber stoppers in 
small bottles of water. These bottles were then set on white paper and 
surrounded by tanglefoot bands. Tattersfield used only adult wingless 
parthenogenetic females, descended from a single female. The successive 
generations were reared upon broad bean plants in pots. In order to 
separate the desired individuals the plants were cut and allowed to wilt 
slightly to make the aphids remove their stylets from the leaf tissue and 
wander about. Then the insects were easily and safely handled with a 
camel's hair brush. If the mouthparts were to remain inserted in the 
leaf they might be injured when the aphids were brushed off. 

Bibliography 

Davidson, J. 1926. The sexual and parthenogenetic generations in the life-cycle 

of Aphis rumicis, L. Verh. Ill Internat. Ent. Kongr. Zurich, 1925, 2:452. 
Franssen, C. J. H. 1927. Aphis fabae, Scop., en aanverwante soorten in Nederland. 

90 pp. Wageningen. (Abst. in Rev. Appl. Ent. 15:464, i9 2 7)- 
Horsfall, J. L. 1925. The life history and bionomics of Aphis rumicis. Univ. 

Iowa Studies Nat. Hist., 11, No. 2, 57 pp. 
Richardson, C. H., and Smith, C. R. 1923- Studies on contact insecticides. U.S. 

Dept. Agric, Dept. Bull. 11 60. 
Tattersfield, F., and Morris, H. M. 1924. An apparatus for testing the toxic 

values of contact insecticides under controlled conditions. Bull. Ent. Res. 14:223. 



Aphididae 3 2 3 

APHIS MAIDI-RADICIS* 

THE complete life history of this corn-root aphis from the egg stage in 
spring to the eggs in autumn has been obtained. The vivaria used for 
the rearing and observation of the aphis consisted of 8- or io-dram vials, 
each containing a ball of moist cotton in the bottom and plugged at the 
top with a piece of cotton. In this cage a sprouting corn plant was 
placed, a reserve supply of these food plants being constantly kept for use. 
The first young and the last young of each generation were placed on corn 
roots in separate vials. These vials were kept in closed boxes to exclude 
light, thus giving conditions probably most favorable to the optimum 
development of the aphis. As soon as the plant began to wilt it was 
replaced by a fresh one, the aphids being transferred by means of a 
camel's hair brush. 

During the life cycle of this aphid there appear five different forms, 
namely: winged viviparous females, wingless viviparous females, ovip- 
arous females, males, and eggs. Eggs were collected originally in the 
nests of the common brown ant (Lasius niger L. var. americanus Emery) 
in April. Taking the first young of the first young all through the series, 
22 generations were obtained. 

M. E. D. 
Reference 
For the rearing of Aphis maidis see p. 397. 

REARING METHODS FOR APHIDIDAE 

F. M. Wadley, U. S. Bureau of Entomology and Plant Quarantine 

CONSIDERABLE work has been done in rearing the green bug 
(Toxoptera graminum) and the apple-grain aphid (Rhopalosiphum 
prunijoliae) ; some rearing has been done with Macrosiphum granarium, 
the pea aphid, the melon aphid, the corn-leaf aphid, and several other 
aphid species. Aphids have been reared almost entirely on growing 
plants, though they will live for a time on cuttings in water. It seems 
inadvisable to try rearing them on detached bits of food plant, as feed- 
ing appears to be almost continuous. Many aphids seem to thrive under 
a wide range of humidity and light conditions if on a favorable and 
thriving food plant. They also develop under a fairly wide temperature 
range, but it is difficult to carry some species through a hot summer. 
In warm weather a shady, moist, well ventilated place should be sought 
for cages for such species. Some species are quite susceptible to fungous 
disease, and precautions must be taken against its entry. 

Cages must permit food plants to thrive, but otherwise have not 

* From an article in U. S. D. A. Tech. Ser. Xo. i2:Pt. 8, 1909, by J. J. Davis, Purdue 
University. 



324 Phylum Arthropoda 

presented a difficult problem. Most aphids develop well in confinement, 
and do not often try to leave a favorable plant. The old device of a 
cloth topped lantern globe on a flower pot containing the food plant 
answers the purpose fairly well. Several modifications have been used. 
Glass lamp chimneys with the top closed with cloth or stoppered with 
cotton have been successful ; light weight mica chimneys have been un- 
satisfactory outdoors because they blow off easily. If plants in the flower 
pot are small, observations may be made easily, and special precautions 
may be taken to make the cage tight and to keep the soil smooth and 
bare. If it is desired to find molts and dead insects without fail, paper 
may be placed on the soil, or melted paraffin may be poured over the soil 
to form a temporary floor. 

Plants should be of a favorable species; aphids will not thrive on all the 
plants they are known to infest. They should also be in a thrifty growing 
condition; new and vigorous seedlings are especially favorable. Trans- 
ferring has been done with a small camel's hair brush; in transferring, the 
worker should be sure the aphid has a good foothold on the plant before 
leaving it. Some aphids, especially small nymphs, will fail to get back 
on the plant if they fall on the soil. 

References 

For the culture of Toxoptera graminum see p. 397. 
For the culture of Macrosiphum cornelli see p. 499. 

CULTURE OF APHIDS 

A. Franklin Shull, University of Michigan 

THE breeder of aphids needs to remember that many species will 
feed only on certain plants. They wander from any others on which 
they may be placed and starve rather than accept them. No artificial 
method of feeding them has yet proved feasible. There are, however, 
a number of cosmopolitan species which feed on many different plants. 

If a phenomenon is to be studied which is exemplified in only one or 
a few species, the investigator's first question is whether he can supply 
the food plant for the duration of the study. Aphids feeding only on 
a deciduous tree or shrub are not usually suitable for such studies. If the 
problem is one of general physiology or of a wide spread anatomical 
feature, the cosmopolitan species are available. One who is not well 
acquainted with the group would do well to consult a taxonomist, to learn 
the accepted food plants, before essaying a breeding problem with any 
given species. Once the suitable plant or plants are known, the possibility 
of providing them for the length of time required will be easily ascer- 
tained. 

Some of the common vegetables and garden flowers may be grown 



Aphididae 325 

from seed at any time of year. Among these are beans, peas, cabbage, 
radishes, tomatoes, nasturtiums, calendulas, and asters. Wheat and oats 
are also useful. Each of these is accepted by some of the cosmopolitan 
species, and some of them are the favored food of certain species. The 
most convenient plants for experiments are those which grow rather 
slowly. If a group of aphids is to be left on one plant during its entire 
reproductive period of several weeks, the food plant should not become 
unmanageable in that time. Calendulas and cabbage are among the best 
of the plants named, in this regard, but are not always the most favorable 
to aphid growth. If more rapidly growing plants are used, the aphids 
may need to be transferred to fresh plants at intervals. 

Seeds should be planted, not in the pots with which the experiments are 
to be conducted, but in seed plats. The young seedlings should be trans- 
planted to very small pots, and, when they are satisfactorily established 
in these and begin to grow actively, should be removed, soil and all, 
without disturbing their roots, to the larger pots to be used in the experi- 
ments. 

The common potato is suitable for cosmopolitan species, and is like- 
wise the host of species peculiar to it. The plants may be furnished in 
quantity the year round, but one must look ahead. New potatoes re- 
quire 6 or 8 weeks of rest before they will sprout. Through the winter, 
any potatoes harvested in the fall may be counted on to sprout fairly 
promptly. Old potatoes planted later than May, however, especially 
indoors, are likely to produce a proportion of unhealthy plants. Hence it 
is better to begin to obtain new potatoes (shipped in at first from warmer 
regions) at least as early as March, and periodically thereafter through 
the summer and fall, so that a supply capable of sprouting is always on 
hand. There are ways of hastening the sprouting of potatoes, [See foot- 
note on p. 328.] but none of them is superior to the method of allowing 
time for rest, which requires only foresight. The soil used for any plants 
should be renewed occasionally, and pots should be cleaned. 

Aphids which in nature ordinarily alternate between two host plants 
may often, perhaps usually, be induced to live indefinitely on one of 
them. The most extensive experiments yet done with aphids concerned a 
species that alternates irregularly between rose and potato. It would 
be difficult to use the rose in experiments, but potato plants were found 
to suffice. 

In general, aphids should be placed on a plant while it is still very 
small, and be allowed to accumulate while the plant grows. Since most 
aphids wander on slight provocation, they must be confined. Lantern 
globes closed at the top with voile or cheese cloth are suitable. The pots 
for the plants should be large enough to let the lantern globe rest on the 
soil, thus making an aphid-tight seal of the enclosure. Water for the 



326 Phylum Arthropoda 

plant may be introduced at the bottom from saucers, or more conven- 
iently and with little harm through the voile covers of the lantern globes. 
If the aphids are to be kept for long periods of time at constant tempera- 
tures, light must be supplied for the plants. This may be done by having 
one wall of the temperature chamber made of glass and placing an elec- 
tric lamp outside. If the offspring of certain parents are to be separated 
into successive groups in the order of their birth, it is better to transfer 
the parents from one plant to another, leaving the offspring on the plant 
where they were born. 

The insects are best handled with a moist camel's hair brush. For 
examination alive under a microscope, the aphids may safely be etherized. 
They may also, without serious injury, be immersed for a short time in 
water on a hollow-ground slide to bring out details of structure. 



Family 



COCCIDAE 



METHOD FOR REARING MEALYBUGS, PSEUDOCOCCUS SP.* 

Stanley E. Flanders, University of California 

EARLY investigations by Professor Harry S. Smith in connection with 
the study of the life history of certain parasites of the Baker mealy- 
bug indicated that the potato sprout was the most suitable host plant for 
the latter. While numerous other host plants have been tried, none has 
yet been found which has the year-round availability of seed potatoes, 
the adaptability to simple laboratory methods, and the ability to stand 
the continued abuse of laboratory practices. 

While the propagation of the potato sprouts is a comparatively simple 
problem, success is dependent upon close attention to several details, the 
neglect of any one of which may result in failure. 

The first and a very important point to be considered is the selection 
of seed tubers. In this respect many varieties which possess a reputation 
for heavy top growth have been tried. The variety known as British 
Queeri produces an excellent long, succulent, sturdy sprout but seems to 
retard the development of certain mealybugs with which it is infested. 
California Burbank also produces an excellent sprout, but not all mealy- 
bugs will feed on it satisfactorily. Red Rivers and Bliss Triumphs pro- 
duce an abnormally rapid growth of numerous, large, succulent sprouts, 
but have exhibited at times a tendency to break down very rapidly under 
heavy infestation and high temperatures. Idaho Rurals produce very 
usable sprouts but have too few eyes. 

The Idaho Russet to date has met best all of the requirements of 
laboratory use. It is readily available on the open market from October 1 

♦Extracted from paper by Harry S. Smith and H. M. Armitage. Univ. Calif. Agri. 
Exper. Sta. Bull. 509, 193*- 



Coccidae 327 

to July 1. It comes from an area free from tuber moth. This fact is 
very important, because this potato pest develops rapidly under in- 
sectary conditions, often destroying valuable host material before it may 
be put to its proper use. The Idaho Russet variety produces an abun- 
dance of long, slender, succulent sprouts which are susceptible to easy 
infestation, and it withstands to a marked degree heavy infestation and 
other necessary insectary abuses. 

Well matured, small to medium sized tubers, averaging 4 to 8 ounces 
in weight, possessing an abundance of well formed eyes, are selected for 
planting. Freedom from cuts and bruises is desirable, since the condi- 
tions under which the tubers are placed in the insectary makes them very 
susceptible to destructive rots and molds which gain entrance through 
such injuries. Lots of tubers which show any appreciable percentage 
of Fusarium wilt should be rejected, and as far as possible those affected 
with scab and particularly Rhizoctonia should be avoided. 

Though small seed pieces, split tubers, and seed end pieces have all 
been tried repeatedly in the interest of economy, careful experimentation 
has demonstrated that whole tubers produce better and more hardy 
sprouts under insectary conditions. Sprouts developed from whole tubers 
seem less dependent on the medium in which they are being grown and 
consequently are less seriously affected by the fluctuating moisture con- 
tent of the trays, which may occur under the most careful handling. 

The whole tubers are planted in a prepared soil medium composed of 4 
parts of light sandy loam to 1 part of screened dairy manure. The top 
is then covered with % inch of coarse sand, which serves as a mulch, 
minimizing the danger of drying out and baking under the heated room 
conditions. Fifteen to eighteen tubers, depending on size, weighing 
approximately 6 pounds, are required to plant one tray. 

When planted the trays are placed in the production rooms, usually 
on well constructed racks. These racks should be in the form of evenly 
spaced shelves not less than 12 inches apart. The weight of each filled 
tray is approximately 38 pounds, and vertical supports must be placed 
between the shelves for every 3 trays, allowance being made for sufficient 
side clearance between trays so that they may easily be installed or re- 
moved. The first tier of trays is never placed directly on the floor but 
is supported on i-inch floor strips. 

By stacking the trays checker-board fashion, racks may be dispensed 
with, but permanent cleat supports on the walls at both ends of the room 
are needed. A specially constructed tray is used for this purpose. It has 
a depth of 6 inches on the ends while the sides are the standard 4 inches 
in depth. By stacking the trays on the wider ends a 2 -inch opening is 
provided laterally between trays, permitting cross ventilation throughout 
the entire stack and continued lateral growth of the sprouts from one tray 



328 Phylum Arthropoda 

to another. The trays are usually staggered 12 high, leaving 2 feet 
between the top trays and the 8-foot ceiling. 

Each method of handling has its advantages and either may be used 
with satisfaction. Where racks' are used the trays are more easily placed 
in position ; poor trays may be easily removed and new ones added ; and 
less labor is required to keep the sprouts out of the aisles and within the 
confines of the racks. Where the stacking method is used all of the host 
plant or host insect material in the entire lot of trays is directly accessible 
to the mealybugs. 

The average period required for developing the potato sprouts to the 
point where they are ready for infesting with the mealybug is 60 days. 
However, this period is considerably lengthened during the winter months 
when the "new crop" seed tubers are more or less immature, and reduced 
almost to 30 days in midsummer when fully matured seed from cold 
storage is used. 

The sprouts are allowed to develop in subdued light in order to 
promote longitudinal growth and to limit the formation of chlorophyl, 
which tends to inhibit the settling down of the mealybugs during the 
period of infestation. A temperature averaging 65 ° F. and a humidity of 
approximately 70% is maintained during the growing period. The 
moisture requirements of the growing sprouts necessitate that the trays 
be watered at 10-day intervals. Top trays and others directly exposed to 
ventilators dry out rapidly, and must be watered every 5 days. 

Materials, such as sphagnum moss, wood shavings, sawdust, and coarse 
sand have been used as growing media but with less satisfactory results 
than with prepared soil mixtures. 

Whole tubers in open trays with no growing medium may be used. 
Twice the number of tubers are then required, but a proportionate in- 
crease in production is secured, without any increase in equipment or 
room space. Other advantages of this method of operation are the ma- 
terial decrease in the amount of labor involved in the initial planting and 
in the subsequent care of the room. The necessity for frequent waterings 
with their attendant troubles from overwatering or drying out is elimi- 
nated. In addition, the tuber itself serves as a host of the more mature 
mealybugs. This method, so far as now understood, is limited, however, 
to late-season use when completely mature seed tubers are available; 
otherwise the prolonged period of sprout development makes it un- 
practicable. The development of more effective methods of accelerating 
sprout growth in new tubers would give this method of host plant culture 
preference over any other.* 

♦Editor's Note: The Cobbler, Green Mountain, and Russett Rural varieties of 
potatoes have been sprouted successfully when taken from the field in mid-season and 
stored at 40°F. for a month. The use of ethylene as an aid in speeding up maturity 
has also been successful. — G. F. MacLeod, Cornell University. 



Coccidae 329 

There are several attendant troubles in the growing of the host plant 
material, the more important of which are "tip burn" or "tip dieback"; 
damping-off due to Rhizoctonia; aphis, and potato tuber moth. "Tip 
burn," which affects the sprouts soon after they are out of the ground, 
produces numerous weak laterals, which in turn are themselves often 
affected. Its cause has not been determined. It is, however, believed to 
be entirely physiological and due to a combination of conditions in the 
rooms, particularly relating to ventilation and humidity. 

Damping-off is induced by overwatering and slow evaporation, in spite 
of the fact that each tray is drained through a %-inch crack through the 
entire length of its bottom and that care is taken to avoid saturating the 
soil. Some insectary operators report successful control of damping-off 
by spraying with Semesan, but in general it may be prevented by avoid- 
ing the humid conditions which favor it. Rhizoctonia usually gains access 
because of non-elimination of infected seed when planting. It girdles the 
sprouts at the surface of the soil and, if it does not kill them immediately, 
it so weakens them that they soon succumb to the attack of the mealybug. 

The common mealybug (Pseudococcus citri) is best adapted to labora- 
tory production. This fact is due to its restricted migratory habits, its 
inclination to remain on its host even under over-infested conditions, its 
short life cycle under laboratory temperatures, and a possible high de- 
gree of infestation with least injury to the potato sprouts. 

An ample supply of mealybugs should be made available for evenly 
infesting the host plant material as it matures. 

Infestation of the sprouts is accomplished through the use of temporary 
host material such as the stems and leaves of sunflower and mallow. This 
material is distributed over trays of hatching mealybugs and left there 
until completely infested. It is then collected and placed on new trays 
of sprouts to which the bugs migrate as the temporary host dries. The 
proper degree of infestation is determined by experience. 

When mealybug material is collected in the field and used to infest the 
sprouts it should be placed in cloth bags or perforated paper sacks to per- 
mit the egress of the young mealybugs and retain any parasites that may 
be present. 

METHODS USED IN REARING THE MEALYBUG, 
PSEUDOCOCCUS COMSTOCKI 

W. S. Hough, Virginia Experiment Station 

TWO methods were used in rearing Pseudococcus comstocki on ca- 
talpa: the vial method, and by using potted seedlings 6 to 18 inches 
in height. In the vial method I used straight edged vials i-inch in 
diameter and 4 or 5 inches long. Either a newly hatched mealybug or 



330 Phylum Arthropoda 

an egg was placed on a catalpa leaf which was then inserted in a vial 
and tightly corked. At intervals of two or three days each individual 
was transferred by means of a camel's hair brush to a fresh leaf and the 
old leaf removed from the vial. When the females reached maturity an 
adult male which was still in its cocoon was placed in each vial. The 
eggs were removed daily from each vial during the period of oviposition. 

The potted catalpa seedlings were used to a limited extent in the second 
method. A narrow paper band was placed on the trunk of each seedling 
and each band was kept covered with a fresh coating of tanglefoot to 
prevent migration on or off the tree. Above the tanglefoot band on the 
main trunk or on one of the laterals a band of burlap was loosely tied 
in order to afford a convenient place in which the males could cocoon. 
Sometimes the females also preferred to crawl beneath the burlap band 
just before beginning oviposition. When one individual shifted its posi- 
tion between molts so that it could no longer be distinguished from an- 
other specimen, both specimens were considered lost from the records 
for that stage of their development. No difficulty was encountered in 
rearing the mealybugs on the potted seedlings, but the fact that the in- 
dividuals shift about from place to place makes observations very difficult 
to obtain if a consecutive record from molt to molt is desired for each 
specimen. Usually two to ten mealybugs were kept on each seedling and 
as far as possible not more than one individual was allowed to remain for 
any length of time on a leaf. The occurrence of a molt was indicated by 
the cast skin beside the insect. For a continuous record of the same in- 
dividual from egg to adult it was necessary to employ the vial method of 
rearing rather than rearing a few individuals on each potted seedling. 

It was learned that burlap bands placed on limbs of Catalpa bungei as 
well as on potted seedlings of catalpa offered favorable feeding places for 
mealybugs in all stages of development. A gall-like growth usually de- 
veloped beneath each band after the insects had been feeding for a short 
time. 

Reference 

For a convenient caging method for the Coccidae see p. 320. 



Neuroptera 331 

Order neuroptera 

METHODS OF COLLECTING AND REARING NEUROPTERA 

Roger C. Smith, Kansas State College 

CHRYSOPIDAE (Lace-wings, aphis-lions) 

ADULT chrysopids may be most successfully collected at lights in the 
. early evening and by sweeping vegetation or beating bushes and 
trees during the days (Smith, 1922, 1934). Gravid females have dis- 
tended abdomens and the more common species deposit their stalked 
eggs very readily in captivity. It is desirable to line the bottle or vial 
with leaves or paper so that the eggs will be deposited on them. The 
leaves or paper may then be removed and cut in pieces with an egg to 
a piece for placing in small individual vials. This is desirable because the 
larvae are cannibalistic. 

Adult chrysopids require food. Place leaves with a few aphids on 
them or a pledget of cotton made wet with water or dilute sugar solution 
in the bottle. The vials or bottles must not be so moist as to allow drop- 
lets of water to form because the wings will be caught in the water and 
the adults will die. Adults may be kept for a week or more in a bottle, 
and eggs may be deposited over most of the period. The food should 
be changed daily. The rarer the species of chrysopid the greater the re- 
luctance to oviposit and live satisfactorily in confinement. Hibernating 
prepupae and adults may be brought into the laboratory after they 
have experienced a cold stimulus. They will then develop or lay eggs 
during the winter months. 

Eggs of this family may be collected in the normal habitat of the 
species. Eggs of the tree-inhabiting species may be taken on leaves, 
where they occur generally on the under side, and on the trunks and 
limbs of the tree. Those lace-wings inhabiting low vegetation lay their 
eggs on the under-side of leaves (generally), or on the stems. 

Larvae may be taken by sweeping and beating. Use a regular beating 
or sweeping net and examine the contents carefully. Practice is required 
to see them readily in a mass of net contents. Larvae often occur in or 
near aphid colonies, in aphid-curled leaves, and under or on the bark of 
trees infested with aphids, psyllids, some scale insects, and mealybugs. 

Cocoons may be collected by pulling off pieces of bark on oak and 
maple trees, especially during the winter or spring. Cocoons should be 
placed in large bottles in which there are some twigs or vegetation so the 
pupae may climb up, shed their pupal molts and spread their wings. 
Otherwise many reared adults will have crumpled wings. 

While chrysopid adults and larvae feed on all kinds of aphids, the 
smaller green aphids are more satisfactory for feeding than the larger 



33 2 Phylum Arthropoda 

ones. The cabbage aphid (on cabbage and turnip) is one of the easiest 
of all aphids to maintain as a constant supply for continuous rearing in 
summer and winter. During the season aphids on Spiraea, apple, dock, 
shepherd's purse, and the corn-leaf aphid (also on young sugar cane) are 
wholly satisfactory. 

HEMEROBIIDAE AND BEROTHIDAE 
(Brown lace-wings) 

Collecting and rearing methods described for chrysopids apply 
almost equally well for hemerobiids (Smith, 1923). Members of 
these families are more difficult to collect and all stages are generally 
less plentiful than are chrysopids. 

Adults are best collected by sweeping low vegetation and beating the 
limbs of pines and oaks with a strong insect net. Larvae may be col- 
lected in the same way particularly from pines and oaks. Eggs and 
pupae are very difficult to collect and are discovered usually only when 
making close observations on aphid-infested branches. Some brown 
lace-wings are known to overwinter as adults, and so may be collected 
during the fall and spring. 

CONIOPTERYGIDAE 

Adults and, less commonly, larvae of the "mealy-winged Neurop- 
tera" have been taken by beating the foliage of pine, apple, and oak 
trees with a beating net. These insects are difficult to see in a net be- 
cause of their small size. The net contents must be sorted over carefully. 
It is well to familiarize oneself with the appearance of these insects by 
looking at specimens in collections repeatedly or studying pictures before 
doing much collecting. They occur from midsummer to fall but the 
writer has never found them plentiful. The females may be confined in 
small glass vials plugged with cotton. Place in the vial a portion of a 
leaf with some small aphids, young scale insects, or young mealybugs 
on it for food. A little moisture is taken by adults also. Eggs may be 
laid on the leaf or cotton plug. This same food is accepted by the larvae 
which should be kept in individual vials because of their cannibalistic 
habits. 

MYRMELEONIDAE 

The writer has been unable to obtain eggs by confining mid-western 
ant-lion adults in various kinds of cages. The larvae of the pit-forming 
species may be collected readily by scooping up the sand pits, with 
a coarse tea strainer or other strainer, or with a large spoon, being careful 
to dip below the larvae. Then sift or pour off the sand until the larva 
is found. When thus disturbed the larvae are difficult to see because they 



Nenroptera 333 

feign death or remain immobile for a time and their color blends per- 
fectly with the sand. 

Since many of the ant-lions are known to overwinter as larvae, they 
may be collected in the late summer or fall and allowed to form pits in 
pans of sand. When half grown or larger they will survive several weeks 
to four or five months without food. The pans of sand may be kept 
in a greenhouse and the larvae fed on sow bugs which abound under pots 
or in other places in greenhouses. Any kind of insect is accepted unless 
too large or active to be subdued by the larvae. 

The overwintering ant-lions complete their development in the spring 
and spin their cocoons during May, June, and July (Smith, 1934). 
Large, nearly grown larvae may be collected in the spring and raised 
with little effort. The writer has used a self-feeding device with fair 
results. A small receptacle filled with a sugar solution or diluted 
molasses is set on the sand in the pan. Some of the solution is poured 
around the pan and on the sides of the pan to attract ants. Eventually 
they come and, in attempting to reach the syrup receptacle among the 
pits, many of the ants drop into the pits. Usually it is necessary to 
supplement this method with other feeding, however. 

The non-pit-forming species may be collected under stones, sticks, 
and sometimes under loose soil or dust (Wheeler, 1930). Since there is 
little or no external evidence of their presence, their discovery is largely 
accidental. The writer has been unsuccessful in rearing mid-western 
non-pit-forming ant-lions. 

Cocoons of ant-lions should be placed in some sort of cage provided 
with vegetation upon which the emerging adults may cling to shed the 
pupal molts and spread their wings. Unless this is done a large percentage 
of reared adults will be imperfect. 

MANTISPIDAE 

Eggs have been obtained on several occasions by the hundreds and 
even thousands by confining gravid mantispid females in bottles and 
jars. The eggs were laid on the cloth tops or, less commonly, on 
vegetation and sides of the bottles. The young larvae have refused to 
eat any insects offered or to enter spider egg sacs. The writer has never 
reared these insects to adults. The mantispid adults eat small insects 
readily in confinement and live for a week or two. 

Pupae and cocoons have been taken in the nests of a spider {Philaeus 
militaris) during October (Smith, 1934). No doubt they might be 
taken during the early spring in spider egg sacs and nests on shrubs, 
weeds, and under stones up to early May in the mid-west. The adults 
are ordinarily first taken in May. 



334 Phylum Arthropoda 

ASCALAPHIDAE 

The writer has been unable to rear from egg to adult this uncommon 
group of Neuroptera. Eggs are sometimes deposited by adults in almost 
any kind of container (Smith, 1931). They are unstalked and adhere 
to the substratum. They hatch readily enough, but the young larvae 
have so far refused to feed. 

Larvae of this group have been collected by sweeping grass and other 
vegetation. They live on the leaves largely exposed and sometimes 
almost in colonies. If one is swept up, the search for others should be 
continued. The writer has attempted to rear them through by placing 
them on aphid-infested plants but without success. 

Larvae of other species are found sometimes in digging in loose soil or 
in examining dust layers. Since they do not form pits there is no 
readily perceived evidence of their presence, and their discovery has 
been largely accidental. 

SIALIDAE 

Adults of Corydalis have been reared by bringing in nearly grown 
larvae and placing in an aquarium in which one end is water and the 
other soil. The larvae migrate to the moist soil when fully grown, form 
a pupal cell, and emerge usually in May or June in the mid-west. 

Egg masses of Corydalis are readily recognized as white blotches on 
trees, foliage, rocks, and bridges over water. They hatch normally in 
the laboratory but the writer has never attempted to rear them. They 
have a three year life cycle and are aquatic carnivores which would 
require an elaborate equipment for normal living conditions. 

Bibliography* 

Anthony, Maud H. 1902. Metamorphosis of Sisyra. Amer. Nat. 36:615. 
Bristowe, W. S. 1932. Mantispa, a spider parasite. Ent. Mo. Mag. 18:222. 
Braur, F. 1851. Verwandluggsgeschichte des Osmylus maculatus. Weigm. Archiv. 

i7:255- 
KrxLiNGTON, F. J. 1934. On the life histories of some British Neuroptera. Trans. 

Soc. Brit. Ent. 1:119. 
1932. The life history of Hemerobius atrifrons McLach. The Entomologist 

65:201. 

1932. The life history of Hemerobius simnlans Walk. Ent. Mo. Mag. 



18:176. 

1932. Notes on the life history of Hemerobius pini. Trans. Ent. Soc. South 



Eng. 8:41. 
Hagen, H. A. 1852. Die Entwicklung und der inners Bau von Osmylus. Linn. 

Entomol. 368-418. 
Hungerford, H. B. 1931. Concerning the egg of Polystoechotes punctatus Fabr. 

Bull. Brooklyn Ent. Soc. 26:22. 
Moznette, Geo. E. 1915. Notes on the brown lace-wing. J. Econ. Ent. 8:350. 

♦Writings on Neuroptera in which some reference is made to rearings. 



Mecoptera 335 



Smith, Roger C. 1917. The Chrysopas or Golden Eyes. Nat. Study Rev. 6:261. 

1920. The process of hatching in Corydalis cornuta Linn. Attn. Ent. Soc. 

Amer. 13:70. 

- 1922. Hatching in three species of Neuroptera. Ibid. 15:169. 
1922. The Biology of the Chrysopidae. Cornell Univ. Agric. Exper. Sta. 



Mem. 58:1291. 

1923- The life histories and stages of some hemerobiids and allied species. 

Ibid. 16:129. 

1926. The life history of Eremochrysa punctinervis (Nerr.). Bull. Brooklyn 



Ent. Soc. 21:48. 

1926. The trash-carrying habit of certain lace-wing larvae. Set. Mo. 23:265. 

- 1931. The Neuroptera of Haiti, West Indies. Ann. Ent. Soc. Amer. 45:798. 
1934. Notes on the Neuroptera and Mecoptera of Kansas with keys for 



the identification of species. J. Kans. Ent. Soc. 7:120. 
Townsend, Lee H. 1935. Key to the larvae of certain families and genera of 

Neartic Neuroptera. Proc. Ent. Soc. Wash. 37:25. 
Welch, Paul S. 1914. The early stages of the life history of Polystoechotes punc- 

tatus Fabr. Bull. Brooklyn Ent. Soc. 9:1. 
Wheeler, W. M. Demons of the Dust, A Study in Insect Behavior. N. Y. 

1930. 
Withycombe, C. L. 1922. Notes on the biology of some British Neuroptera. 

Trans. Ent. Soc. London, p. 303. 
1924. Note on the economic value of the Neuroptera with special reference 

to the Coniopterygidae. Ann. Appl. Biol, n: 112. 



Order mecoptera 

BITTACUS 

Laurel R. Setty, Park College, Parkville, Missouri 

Method oj Collecting. An insect net is not a satisfactory instrument to 
use in collecting adult hanging-flies (Bittacus) which are to be used for 
experimental purposes. These long-legged, soft-bodied insects ordinarily- 
fly among weeds and grasses. Many individuals are injured by sweep- 
ing the foliage or on being removed from the net. 

The instrument which I have frequently employed is a very large 
lamp chimney, the top end of which is closed with a piece of cheesecloth 
held in place by a rubber band. By holding the lamp chimney at the 
elongated, closed end, the large basal opening may be quickly thrust 
over the insect as it hangs quietly from some support. Then, after 
making a successful grab in this fashion, one may hold the palm of his 
hand over the opening and carry the fly to a prison box. 

A convenient box to take into the field for holding the flies that are 
collected is one made of wood, about 18" x 16" x 12", and provided 
with several fresh twigs of some woody or semi-woody plant to serve as 
supports for the insects. The box should have a closable opening about 
the size of the base of the lamp chimney. When the opening of the 



336 Phylum Arthropoda 

chimney is placed against that of the box, the hanging-fly will drop into 
the container. 

Not more than twenty or thirty specimens should be placed at one 
time in a prison box of this size, for there is danger that the flies will 
entangle their long legs with those of the other flies. 

Individual Cages. The ordinary Mason glass fruit jar was found to 
be a very satisfactory cage for an individual or a single pair of hanging- 
flies. The jar must be furnished with a twig of buck-brush, or of some 
other woody plant, to serve as a support from which the insect may 
suspend itself. The opening may be closed with a small piece of cheese- 
cloth fastened in place by a rubber band. All jars should be kept in a 
shady part of the laboratory and preferably upon the floor or upon low 
stools, in order to make conditions as nearly natural as possible. 

Such cages I have found advantageous for the following reasons: 
(1) they are convenient to handle; (2) the hanging-flies seem satisfied 
in this sort of a cage; (3) the insects may easily be transferred from 
one jar to another; (4) the eggs may be collected readily by inverting 
the jar; and (5) small flies and other insects placed in the jar for food 
cannot escape. 

Care of the Immature Stages. The most convenient containers for 
keeping the hanging-fly eggs are small or medium sized clay flower pot 
saucers lined with moist, white cellu-cotton over each of which the 
flower pot itself is inverted. By adding a small amount of water each 
day to the cellu-cotton, the eggs may be kept moist. 

After hatching, each larva should be transferred to a similar container, 
that differs only in having a little pile of about a tablespoonful of rich 
garden soil upon the cellu-cotton. Moisture and tiny pieces of beef 
steak must be provided daily on the cellu-cotton near the soil. 

When the larvae enter the soil to pupate, less moisture should be pro- 
vided, but by no means should the soil be allowed to dry out. 

Order trichoptera 

ON REARING TRIAENODES* 

CADDISWORMS of this genus are common in beds of the water- 
weed Elodea. They swim freely about, carrying their slender, 
spirally-wound cases with them. They eat plant tissues, especially 
Elodea, and are easily maintained in aquaria, even in tap water, and 
at all seasons. When grown they attach their cases lengthwise to the 
stems or to other solid support for pupation. The pupal stage lasts 7 
to 12 days. When ready for final transformation the pupa leaves its 

* Abstracted from a thesis by Wynne E. Caird in the Cornell University Library. 



Lcpidoptera 337 

case and swims to the surface of the water. It often meets with difficulty 
in breaking through the surface film, and may, after a brief struggle, 
fall exhausted to the bottom. 

The slender, straw-yellow adults may be placed on emergence in an 
aquarium cage having a celluloid window for observation in one side 
of the wirecloth top, and a sleeve for ingress and egress at the other 
(see figure 61 on page 268). They cling immobile to the sides and the 
top of the cage during the day, and run about the walls in the early 
evening, rarely flying. 

J. G. N. 



Order lepidoptera 

A METHOD OF COLLECTING LIVING MOTHS 
AT SUGAR BAIT* 

IN COLLECTING adults of the army worm and other moths for ovi- 
position studies, a simple method of capture at baits was found useful. 
[See also p. 364.] An electric flashlight, having a flat lens, was used 
with a flat-bottomed vial % inch in diameter, straight-sided and about 5 
inches deep. When the bottom of the vial is placed against the lens of 
the flashlight both may be firmly grasped in the right hand. The en- 
circling fingers prevent the spread of light rays to the sides while full 
illumination is given in a narrow beam through the bottom of the vial. 
When this is placed over or close to a moth feeding at bait, the tend- 
ency of the moth is to dash towards the light. Entering the vial, it goes 
at once to the bottom and does not try to escape. The vial may be 
closed with a cork or the moth may be examined before the vial is closed 
and, if not wanted, it may be allowed to escape by moving the vial from 
the light. A number of vials may be carried in a coat pocket and a 
number of individual moths collected in a short time. In the closed vial 
the moths remain quiet for a number of hours and may be removed to 
breeding cages. 

The same method was found useful in collecting the large and shy 
Catocala moths. A wide-mouthed bottle was used in this case, the 
flashlight directed through the bottle, and chloroform or cyanide placed 
in holes in the cork. [See also p. 364.] 

Reference 

Family Eucleidae 

For rearing see p. 365. 

♦Reprinted, with slight changes, from Canad. Ent. 60:103. 1928, by R. P. Gorham, 
Dominion Ent. Lab., Fredericton, X. B. 



338 Phylum Arthropoda 

Family tineidae 

A METHOD FOR BREEDING CLOTHES MOTHS 

ON FISH MEAL 

Grace H. Griswold, Cornell University 

WHERE large numbers of clothes moths are used for testing 
various methods of control, it is often difficult to keep on hand a 
supply sufficiently large to meet all needs. With fish meal as the sole 
food supply, an efficient method for rearing them has been devised at 
the Cornell Insectary. 

The meal which has been found so satisfactory is "white fish meal" 
and is manufactured by the Dehydrating Process Company, Boston, 
Massachusetts. 

Cylindrical cardboard cartons, such as are used for packing butter 
and cheese, make good rearing cages. The gallon-sized carton is about 
7 inches high and 6% inches in diameter. The cartons are inexpensive, 
and, since they may be stacked upon shelves, they occupy very little 
space. 

In the center of a large square of cloth (about 22 by 22 inches) is 
pasted a circular piece of heavy cardboard, slightly smaller than the 
bottom of the carton. The cloth is placed in the container so that the 
cardboard rests on the bottom. A layer of fish meal, about half 
an inch thick, is spread evenly over the circle of cardboard. The cloth 
is then folded back against the sides of the carton and the cover is put 
on. Adults are admitted to the container through a small opening cut 
in the cover. This opening is closed on the outside with a piece of 
cheesecloth secured by a little paste. To infest a new container it 
is only necessary to catch a few adults in small vials and drop them 
into the opening in the cover of the carton. The females have easy 
access to the fish meal and will lay quantities of eggs. At temperatures 
similar to those of an ordinary living room, adults will begin to emerge 
within about two months after a container has been infested. 

When one wishes a supply of larvae, the carton may be opened and 
the square of cloth carefully lifted out. In this way the entire contents 
of the carton may be removed without disturbing the layer of fish meal, 
which rests on the circle of cardboard in the center of the cloth. Larvae 
will be found crawling all over the cloth and may be removed with 
the aid of a camel's hair brush. If the cloth is black, the white bodies 
of the larvae stand out clearly. 

Since adults shun the light, they will be found hiding in the folds of 
the cloth where they are easy to catch. Some, of course, will escape 
when the carton is opened. To obviate this, one may have a second 



Tineidae 339 

piece of dark cloth ready and throw it over the carton as soon as the 
cover is removed. Adults will hide in the folds of this second cloth 
as well as in those of the cloth already in the carton. If the folds of the 
two cloths are turned back slowly and carefully, a number of adults 
may be caught in a few minutes. 

It has been found that larvae are more easily removed from small 
containers than from large ones. The half-pint size, commonly used at 
stores for cottage cheese and ice cream, is just about right for this pur- 
pose. Several pieces of flannel, on which eggs have been laid, are placed 
in one of these small containers, each piece of flannel being sprinkled 
lightly with fish meal. The larvae get enough to eat but cannot bury 
themselves in a thick layer of meal such as is put in the large containers. 
By using a number of these small containers, one can keep on hand 
a supply of larvae of any age desired. 

Adults are difficult to catch in the small containers, since they have 
no place in which to hide and can so easily escape. For rearing adults 
it is advisable to use the gallon size and black cloth. Hence at Ithaca 
both the large and small containers are in constant use. 

In the summer of 1933 some of our containers became infested with 
predacious mites. Although the point was never proved, mites were 
probably carried from old containers to new ones on the bodies of adult 
moths. It therefore appeared advisable to start new colonies with eggs 
instead of with adults. 

To get a supply of eggs, moths are placed in a glass jar or vial with 
pieces of flannel. Each day fresh flannel is put in and the pieces on 
which eggs have been laid are removed. If the flannel is cut into squares, 
about 3x3 inches, the pieces are easy to examine under a binocular 
microscope, and any mites present may be removed. The gallon card- 
board containers are prepared as explained above. After examination 
under a binocular, the pieces of flannel, on which the eggs have been laid, 
are placed on the layer of fish meal in the bottom of the container. Each 
piece of flannel is sprinkled lightly with some of the fish meal before 
another piece is added. Since this method of starting new colonies was 
adopted, no further trouble with mites has been experienced. 

By this method literally hundreds of clothes moths may be reared in 
a very small space. At the Cornell Insectary, one or two large cartons 
and several small ones are infested each month, in order to insure an 
adequate supply of insects for all needs that may arise. Just how long 
an infestation will keep itself going, has not yet been determined. If 
a little fresh fish meal is occasionally added to each container, it seems 
probable that the various colonies will maintain themselves for a con- 
siderable period of time. 

Although the method was developed for breeding the webbing clothes 



340 Phylum Arthropoda 

moth (Tineola bisselliella) , the case-making species (Tinea pellionella) 
may be reared satisfactorily in a similar manner. 



Family gelechiidae 



THE GOLDENROD GALL-MAKER, GNORIMOSCHEMA 
GALLAESOLIDAGINIS* 

THE writer has never encountered any difficulty in securing eggs 
from adults of G. gallaesolidaginis in the fall. Moths have been 
caged time after time with living goldenrod stems in glass cylinders with 
cheesecloth top, and in large outdoor cages 5x5x4 feet in size built over 
clumps of goldenrod. In this way thousands of eggs have been secured. 

Female moths begin to deposit eggs within 4 or 5 days after emergence 
when they have been previously confined with males. The eggs are 
placed on both surfaces of a leaf, but usually on the under side and 
preferably on one that is dried, and on the stems among the hairs. 

Moths have been observed to partake of nourishment in the form of 
sweetened water when placed in indoor cages. It may be this factor 
which causes females to deposit a greater average number of eggs in 
indoor cages than under the semi-natural conditions of outdoor cages. 

The first determination of the time of hatching of the eggs followed 
the failure in three successive winters of attempts to keep them through 
the winter under semi-natural conditions. During the fourth winter of 
experiment, eggs deposited regularly on dried leaves and stems of golden- 
rod in the fall were carried through by placing these leaves and parts of 
stems in small shell vials stoppered loosely at both ends with cotton; 
the vials were in turn placed in larger vials similarly stoppered. This 
method solved the problem of continued excessive moisture previously 
encountered. The eggs hatched normally, the larvae producing galls 
which were found on the new goldenrod shoots at the same time that 
the galls of a similar size were present in the field. 

M. E. D. 

MASS PRODUCTION OF SITOTROGA CEREALELLA 

Stanley E. Flanders, University of California 

AMONG the Lepidoptera, those that feed on stored products are 
l most readily reared. The grain moth, Sitotroga cerealella, is the 
most adaptable for continuous reproduction in large numbers. This is 
largely due to certain habits and tropisms that permit efficient mechan- 
ical manipulation of the species. Also Sitotroga appears to have a much 

♦From an article in /. N. Y. Ent. Soc. 30:81, 1935, by R. W. Leiby, North Carolina 
State Department of Agriculture. 



Gelechiidae 341 

greater immunity to disease epidemics than any other species of grain 
moth. 

Sitotroga feeds on a number of different grains, among which are 
wheat, corn, and oats. The dry kernels are used in bulk for rearing pur- 
poses. The hygroscopic moisture within the kernels should not be 
below 8' , . During the feeding and pupal stages, the insect occupies 
the interior of the individual kernels so that the interspaces between the 
kernels are not clogged with excreta or webbing. The newly emerged 
adult moths, therefore, always have free egress from the mass of grain. 
Upon emergence from the kernels the moths worm their way upward 
and outward until they reach a space of sufficient size to permit their 
wings to expand and mating to take place. 

By day the moths tend to climb upward and come to rest in positions 
of positive thigmotropism and negative phototropism. By night they 
become active and oviposit in crevices about 0.23 mm. in width. The 
newly hatched larvae are negatively phototropic and positively geotropic, 
so that they readily permeate a mass of grain. If the moths are col- 
lected daily, few, if any, deposit their full quota of eggs or die within 
the production unit. 

The type of unit used depends to some extent on the rearing medium 
and the size of the grain used. Small kernels yield smaller moths than 
do large kernels, and the smaller moths deposit fewer eggs. Moths 
reared in corn deposit over three times as many eggs per moth as those 
reared in wheat. On the other hand, given equal weights of grain, the 
rapidity with which the grain is utilized varies inversely with the size of 
the kernels; so that a much greater number of moths is obtained in a 
shorter period of time with wheat than with corn. This is due to the 
fact that a kernel of grain, irrespective of its size, is usually inhabited 
by only one larva at a time. 

Laboratories producing for seasonal field requirements find that soft 
varieties of wheat form the most satisfactory rearing media. In experi- 
mental work where a uniformly non-seasonal supply of moths is needed, 
without the necessity of frequently replenishing the food, soft varieties 
of corn are used. 

The "tilted bin" type of production unit adapted particularly for 
holding corn, has been developed by the University of California. A 
"vertical bin" type of unit for holding wheat was developed by the 
U. S. Bureau of Entomology. 

Of these two types of production units, the "tilted bin" type (Fig. 66) 
is the simplest in construction but is not as readily refilled as the "ver- 
tical bin" type; therefore it is only well adapted for use with a slow 
productive medium such as corn. Essentially it consists of a series of 
similar shallow trays (36" x 26'" x 3") placed one on the other and 



342 



Phylum Arthropoda 



topped with a cover. It rests on a base frame so constructed that the 
trays are tilted to an angle of 22.5 to the horizontal. The slope thus 
formed is slightly less than the "angle of flow" of corn when piled in a 
loose heap. It is sufficient, however, to direct the egress of the moths 
from the trays, and to provide good ventilation. 

The floor and two sides of each tray should be made of material of 
sufficient strength so that when the tray is filled the bottom will not 

sag, and when stacked each tray will 
support the entire weight of the trays 
above. 

The upper end of each tray should 
be at least %6 mcn below the top 
edge of the other three sides, in order 
to provide exit for the moths. The 
lower end should be formed of 30- 
mesh copper screening for ventilation. 
Across the top of the tray and flush 
with the rear end is firmly fastened 
a wooden "stop," 2 inches wide, in 
order to hold each tray and the top 
cover in position. The upper ends of 
the trays should be formed so that 
when the trays are stacked in place 
they form a flat, vertical wall. 

In setting up a unit, each tray is 
filled with corn before adding the next 
tray. Care should be taken to make 
the top surface of the corn even with 
the top edge of the upper end and 
parallel with the bottom of the tray 
above, thus leaving a space of % 6 
inch in height over the entire surface of the corn. 

When the unit is filled and the cover in place, a strongly woven cloth 
is loosely draped over the vertical front of the stack and fastened on 
top and at the sides with felt-lined batten placed flush with the front 
surface. The lower edge is fastened to a flange of the collecting tube 
extending along the entire front below the bottom tray. 

The tube is made of sheet iron, 1% inches in diameter, with flanges 
2% inches in width, which diverge outward from a slot % 6 inch wide 
extending along the top side of the tube for the width of the trays. 
The tube is held in position by the inner flange, which is inserted 
between the base and the bottom tray. It is placed so that the inner 
side of the slot is flush with the upper end of the bottom tray. Attached 




Fig. 66.- — Diagrammatic cross section 
through the center of the "tilted bin" 
type of production unit. 



Gelechiidae 343 

to the outer flange by small bolts and winged nuts is a rigid wooden bar. 
The cloth is fastened against this by the pressure of the bar against the 
flange. The cloth then forms an enclosure with the front of the stack 
within which the moths collect. 

The collecting tube extends several inches beyond each side of the 
stack. One end is plugged with a cork and the other connects with a 
tube extending through the cover of the moth trap. This trap consists 
of a Royal electric hairdryer fastened permanently to the cover of a 
gallon mayonnaise jar. The air intake opening of the dryer is connected 
by a short right-angle tube to an aperture in the cover containing a 
30-mesh copper screen to retain the moths in the jar. When the jar is 
screwed onto the cap it completes an air suction circuit between the 
hairdryer fan and the moth enclosure. 

The moths are collected in a few seconds by starting the fan and 
shaking the cloth to dislodge the moths accumulated on it and on the 
front of the stack. This is possible because the moths when suddenly 
disturbed habitually drop down and slide through the slot at the bottom. 
The air suction carries them into the jar without injury. Moths that 
remain in the tube are pushed through with a bottle brush inserted 
through the opposite end. 

In setting up a production unit it is necessary to use a pure culture of 
Sitotroga. Contamination by other grain insects or by enemies of 
Sitotroga, such as the mite Pediculoides ventricosus and the chalcid 
Habrocytus cerealellae, must be avoided in order to insure continuous 
and economical production. This may be accomplished by using clean 
rooms and by sterilizing equipment and grain with carbon bisulfide in a 
vacuum fumigator. The grain should not be inoculated with Sitotroga 
until the fumigant has dissipated and the proper hygroscopic moisture 
has been reached through holding the grain at 70% relative humidity 
for several days. Then it should be well infested with a pure culture of 
eggs. This may be done while the grain is in the sacks. If the units 
are to be operated continuously they should be refilled with infested grain 
that has produced one or more generations of moths. At a temperature 
of 8o° to 85 ° F., the period from egg to adult is from 25 to 30 days. 

The "vertical bin" unit (illus. in Peterson, 1934) consists of a series 
of narrow trays of 12 -mesh wire (24" x 12" x %") open at the top 
and suspended in a row in a metal box about 30 inches square having 
sides of 60-mesh copper screen. Six-inch holes in the top provide open- 
ings for the moths to enter the traps placed above each hole. These 
traps consist of coffee tins with inverted funnels soldered to the rims of 
the screw tops. The percentage of moths collected through such 
"geotropic" traps used with this type of unit is probably less than in the 
suction trap used with the ''tilted bin" type. 



344 Phylum Arthropoda 

The geotropic method of collecting the moths may also be used with 
the "tilted bin" unit (Fig. 66). Instead of hanging loosely, the cloth 
across the front of the stacked trays should be stretched taut and 
fastened securely about % inch away from the trays. The enclosure 
thus formed should open at the top and at the bottom into sheet iron 
funnels. The base of each funnel should be % inch wide and as long 
as the width of the stack, and the small end of each funnel should fit 
into the opening of the moth trays. 

The temperature and humidity in the rooms housing the production 
units should be regulated so that the air surrounding the kernels in the 
mass of grain will range in temperature between 75 ° and 85 ° F., and in 
humidity between 60% and 70%. Such humidities provide optimum 
hygroscopic moisture for the development of the insect larvae, and the 
temperatures mentioned result in the most rapid succession of genera- 
tions. 

The activity of the larvae themselves is a source of heat that raises the 
temperature of the grain mass higher than the surrounding air (heat of 
infestation). If the grain is not permitted to heat as a result of ex- 
cessive moisture (heat of respiration) the status of the infestation may 
be roughly gauged by taking the temperature of the mass. The tempera- 
ture of the room housing the production units should be comparatively 
low. 

Reference 

For the culture of the Angoumois grain moth see also p. 497. 

Bibliography* 

Bailey, C. H. 1921. Respiration of shelled corn. Univ. Minn. Agric. Exper. Sta. 
Tech. Bull. 3. 

Flanders, S. E. 1927. Biological control of the codling moth. J. Econ. Ent. 
20:644. 

1929. The mass production of Trichogramma minutum Riley and observa- 
tions on the natural and artificial parasitism of the codling moth eggs. Trans. 
Fourth Inter. Congress Ent. 2:110. 

1934. Sitotroga production. J. Econ. Ent. 27:1197. 



Peterson, Alvah. 1934. A Manual of Entomological Equipment and Methods. 

Part I, plate 137, fig. 1-4. 
Schread, J. C, and Garman, P. 1933. Studies on parasites of the Oriental fruit 

moth. Conn. Agric. Exper. Sta. Bull. 353:691. 
Simmons, P., and Ellington, J. W. 1932. A biography of the Angoumois grain 

moth. Ann. Ent. Soc. Amer. 25:265. 

*Post script: Since the completion of this Compendium there has appeared Circular No. 
376, U. S. Department of Agriculture: New Equipment for Obtaining Host Material for 
the Mass Production of Trichogramma minutum, an Egg Parasite of Various Insect Pests, 
by Herbert Spencer, Luther Brown, and Arthur M. Phillips, all of the U. S. Bureau of 
Entomology and Plant Quarantine. 



Tortricidae 345 

SITOTROGA EGG PRODUCTION 

Stanley E. Flanders, University of California 

IT is estimated that the moths produced from one pound of corn will 
deposit about 40,000 eggs, while those produced from an equal 
weight of wheat will yield about 80,000 eggs under mass production 
conditions. 

The moths will oviposit in almost any narrow crevice, such as found 
between strips of paper fastened together by clips or between the bodies 
of the moths themselves in crowded containers. Almost any type of 
container may be used, provided it is small enough to cause the moths 
to crowd together. The cover should consist of a 20-mesh screen if 
moths are from corn, or a 30-mesh screen if from wheat. They deposit 
most of their eggs within 72 hours after emerging from the grain if 
held at a temperature of 8o° F. and relative humidity of 70%. At the 
end of this period the moths should be vigorously stirred and shaken to 
dislodge any eggs adhering to them. The eggs are then sifted out through 
the screened cover and winnowed to free them from moth scales and 
debris. The finest of the scale dust may be eliminated by keeping the 
moth container inverted over a trough in a constant current of air. 

By this method loose eggs are obtained. They are then evenly 

spread in a single layer over cards of uniform size thinly coated with 

shellac. This affords a fairly accurate means of measuring production. 

The chorion of the egg is relatively tough so that they may be handled 

in mass as readily as grains of rice. At 8o° F. the moth larvae hatch in 

about 5 days. 

Bibliography 

Flanders, S. E. 1928. Developments in Trichogramma production. 7. Econ. 

Ent. 21:512. 
1930. Mass production of egg parasites of the genus Trichogramma. Hil- 



gardia 4:465. 



Family tortricidae 



NOTES ON BREEDING THE ORIENTAL FRUIT MOTH, 
GRAPHOLITHA MOLESTA 

W. T. Brigham, Connecticut Agricultural Experiment Station 

THE present need of large numbers of larvae of the Oriental fruit 
moth for mass production of parasites has led to the adoption of 
the following procedure in breeding at the Connecticut Agricultural Ex- 
periment Station. 

The original moths were obtained from infested twigs collected from 
orchards during the first and second broods. Infested terminal twigs 



346 Phylum Arthropoda 

are easily observed due to their wilting. Clip these twigs to a length 
of 3 to 4 inches, stripping off the leaves to prevent overheating when 
put in bags for convenient carrying. Upon completion of collections 
and arrival at the laboratory, the twigs are spread on pans of i%-inch 
green apples so that larvae emerging from the drying twigs may have 
sufficient food for normal development. Pans used are galvanized re- 
frigerator pans, 15 inches in diameter, placed in an incubator regulated 
for 8o° F. and 50% relative humidity. Twigs are removed after three 
days and corrugated paper strips, % mcn m width, are fastened around 
the pan tops by means of metal strips doubled to form a V, one end 
being longer and bent over the pan lip to hold it in place. Unbleached 
muslin covers are tied over the pans to prevent mature larvae from 
crawling away, as not all find openings in the strips. As soon as the 
larvae begin spinning, the corrugated strips are removed every other day 
or, at times, daily, and new strips substituted. Strips containing the 
spun larvae are treated in two ways. Those for immediate emergence 
are placed in refrigerator pans, covered with muslin and kept in the 
emergence cage. The others are clipped into jelly jars and held at a 
temperature below 45' F. 

An emergence cage may be constructed by making a framework of 
two by fours with a door hung at one side. Shelves are built along the 
side to hold the refrigerator pans and a bench for working. The frame 
is enclosed with black cambric except for one end (covered with white 
cheesecloth), which faces the room windows. A temperature of 75 ° 
to 8o° F. and a humidity of about 50% is maintained by means of air 
conditioning apparatus. After about 6 days from date of spinning moth 
emergence starts and the pans are left open, allowing the moths to fly 
directly to the white screen. Covering the pans is an emergency measure, 
as parasites attacking the moths in the prepupal stage may become 
numerous. 

The moths, being phototropic, are easily collected from the white 
screen by means of a suction apparatus. An ordinary hand hair dryer 
with the heating unit removed is used for creating a suction sufficient 
to draw the moths from the screen into the prepared containers. The 
containers used are ordinary pint Fonda cartons with a i-inch hole cut 
in the bottom and stopped up by a cork covered with cheesecloth. The 
top is partly cut away, leaving just the outer ring and a narrow section to 
help stiffen the cover on its being replaced. A section of cheesecloth 
placed over the carton and the cover forced back in place makes a tight 
container with plenty of ventilation. The cheesecloth may be dampened 
to aid in preserving the confined moths. A ring of cardboard slightly 
larger than the pint containers mentioned is fastened to the side of the 
hair dryer where the air is sucked in. Elastic bands with clips attached 



Tortricidae 347 

hold the Fonda carton in place on the dryer, and a cork bored with a. ]/ 2 - 
inch hole in which is inserted a 2 -inch length of glass tubing is sub- 
stituted for the cloth-covered cork. By using a rheostat to regulate the 
speed of suction, moth collection is quickly and easily carried out. As 
many as 400 moths may be caught in one carton if transfer to the ovi- 
position cages is made immediately. 

The oviposition cage found most effective is a 15" by 16" by 28" 
frame covered with cheesecloth. The cloth should be on the inside of the 
frame, as the moths will deposit eggs on the smooth frame surface. 
Temperatures should be maintained at 8o°-85° F. This cage is placed 
on damp sand directly below a window of a celotex incubator in the 
greenhouse. A strip of turkish toweling with one end immersed in a 
pan of water, the other end lying on the top of the cage, furnishes mois- 
ture for the moths. Egg laying begins in 2 days, reaching a peak at 6 
days. Peach seedlings grown from pits, or seedlings obtained from 
orchardists and grown in the greenhouse in 5-inch pots are used for 
egg deposition. The pots are introduced and removed daily. The 
leaves are stripped from the plants and estimates of egg depositions 
kept. In exposing the pots the leaves should rest against the top or 
sides of the cage, as a much larger egg production results when the moths 
crawl about from the cloth of the cage to the leaves for oviposition. A 
temperature of 8o° to 85 ° F. and humidity of 50 to 60% should be 
maintained at dusk when maximum egg deposition takes place. 

The leaves on which the eggs have been deposited are placed in jelly 
jars or may be refrigerated at 50 F. for 2 weeks without reducing 
below 80% the number of eggs hatching. Usually the jars are placed 
directly in an incubator, temperature 8o° F., relative humidity 50%. 
After 3 days the eggs appear to be black-spotted because of development 
of the embryos. At this time they are removed from the jelly jars and 
spread out upon green apples in refrigerator pans. Approximately 4,000 
eggs are used for each pan of about 70 apples. To facilitate entrance to 
the apples, punctures are made in the apples or they may be sliced, as 
described later. It has been found that a larger number of larvae tunnel 
into the fruit after the tough outer skin has been broken and a 
depression, into which larvae may crawl, made available. A 6-inch knife 
with a thin flexible blade is used in cutting the apples into slices of 
%-inch thickness. The apples are not cut all the way through, so that 
the slices stay in place. The green apples used are 1% to 2 inches in 
diameter, gathered when orchardists are thinning, and placed in cold 
storage for use at any season of the year. 

As the larvae develop it is necessary to divide the apples into two 
pans and add a fresh supply of green apples to make sure sufficient food 
is available for normal growth. Corrugated strips are fastened around 



348 Phylum Arthropoda 

the edge of the pans and held by clips as described earlier. These strips 
are removed and handled like the original stock. 

Another method of augmenting the stock of moths in the fall is by 
the use of infested quinces placed in butter tubs prepared for their 
reception. Three or four %-inch holes are bored in the bottom of the 
tubs and covered with a fine mesh wire screen. This aids in ventilating 
and delays fruit rot. Two inches below the tub rim two rows of cor- 
rugated paper strips are tacked completely encircling the sides. The 
tubs are half-filled with infested quinces, and muslin covers tied over 
the top. The fruit moth larvae, upon completing development, leave the 
fruit and spin in the corrugated paper. The strips must be removed 
when filled with larvae and new strips tacked in place. These tubs are 
left in the insectary and the spun larvae allowed to remain in covered 
refrigerator pans where hibernation takes place as a normal procedure, 
due to low temperatures. Hibernation may be broken by the end of 
December, the spun larvae being placed in a refrigerator at 55 F. for 
about 3 weeks and then brought out into the emergence cage. A gradual 
change in temperature is found better than a sudden change. 

There are numerous pitfalls along the path of fruit moth breeding. 
Ants and spiders in the greenhouse oviposition cages and spiders in the 
dark emergence cage may cause trouble. Parasites may develop unless 
pans are kept covered, Dibrachys boucheanus being especially trouble- 
some at Connecticut for two years. There is need in the fall to control 
temperatures since at that time larval hibernating tendencies increase 
and temperatures fluctuating below 6o° F. may cause hibernation of 
developing larvae. Once hibernation has started, we have been unsuc- 
cessful in breaking it until after approximately two months have elapsed. 
Oviposition-cage temperatures at dusk are very important if a fair ratio 
of increase in numbers is to be maintained. 

Fruit rot is another difficulty causing serious losses of larvae. Apples 
from different sources rot differently; that is, some will produce a wet 
slimy mass which apparently drowns the larvae before they can escape 
to other fruit or reach maturity. Other apples rot with less moisture, 
becoming pithy or corky, allowing the larvae to escape when the fruit 
becomes unsuitable for food. Green Baldwins are plentiful in Connecti- 
cut, keep well in storage, and are used by us almost exclusively. Other 
varieties, however, may be employed. 

In refrigeration of spun larvae or eggs, it is necessary to guard against 
the loss of moisture which may prevent transformation of the larvae 
and hatching of the eggs. 

Maintenance of seedlings for use in the oviposition cages requires 
a certain amount of thought. By use of fertilizers and cleaning off any 
eggs laid upon the stems of the seedlings, these plants may be used at 



Pyralidae 349 

least twice. In summer small branches are clipped from peach trees 
and placed in vials of water. These remain fresh long enough for ex- 
posure to the moths. Forsythia and privet are also used, the moths lay- 
ing on the leaves of such shrubs nearly as well as those of peaches. Lilac, 
pear leaves, and probably others, may be used successfully. 

Careful adherence to the above methods of breeding has led to suc- 
cessful year around production of the Oriental fruit moth in Connecticut. 
The numbers which may be reared are apparently only restricted by the 
amount of space, time, material, and labor available. 

Our average ratio of increase is 17 to 1, although in some cases 30 
to 1 may be obtained. The sex ratio is about 50' ,' c each, males and 
females. Maximum production was reached during the month of August 
1934, when 376,000 fruit moth eggs were obtained. 

Reference 
For the culture of Harmologa fumiferana see p. 488. 



Family pyralidae 



BREEDING METHODS FOR GALLERIA MELLONELLA 

T. L. Smith, College of the Ozarks, Clarksville, Arkansas 

The Wax Moth, Galleria mellonella, is found wherever bees are cul- 
tured. The adults never eat; the larvae feed only on honeyless bee 
combs, preferably the old brood combs. To the apiarist, it is a very 
potent pest and quickly destroys his weaker colonies. Throughout the 
larval stage during which there are eight instars with their terminating 
ecdyses, food is consumed in great quantities. The larvae also spin silken 
threads from the time they hatch to the time of pupation. During the 
last few days of their larval life each larva spins about itself a compact 
silken cocoon. The duration of the larval stage is about 35 days, the 
pupal stage 12 to 14 days, and the adult stage about 10 days at ordinary 
room temperature. Mating ordinarily occurs the first day after eclosion. 
The females lay from 200 to 1,000 eggs or more. 

The adults show a distinct sexual dimorphism. The male is generally 
lighter in color than the female and the posterior edges of the fore wings 
are notched at their ends while the ends of the wings of the female are 
almost straight. The antennae of the male are 10% to 20% shorter 
than those of the female. In the male the labial palps are hooked 
inward while those of the female protrude forward and slightly upward 
giving a pointed or beak-like appearance to the front end of the head. 
The female has a distinctly long ovipositor which is almost prehensile in 
its use for seeking out places for oviposition. The eggs are whitish in 



35° Phylum Arthropoda 

color and generally spherical or slightly ovoid and are deposited in any 
available niche or corner in the comb. About 9 to 10 days are required 
for the eggs to hatch. The newly hatched larvae are very small — about 
1 mm. in length and perhaps % mm. in diameter. The young larvae 
begin to eat and to spin their webs immediately after hatching. They 
have a tendency to congregate into a rather compact colony, and in this 
colony they often have the power of raising their temperature some io° 
to 15 C. above that of the environment. 

For culturing purposes, the regular milk bottle used for Drosophila 
has been found very adaptable. Instead of using the cotton plugs 
(which will not retain the larvae), the regular waxed cardboard milk 
bottle tops with attached lifters have been found preferable. These fit 
the top of the bottle very tightly and serve to prevent the young larvae, 
which crawl up the sides of the bottles, from crawling out and leaving 
the culture. Since mating may occur within a few hours after eclosion, 
it is necessary to isolate the female pupae before eclosion if virgin females 
are desired for mating purposes. This may be done very readily by 
careful observation of the pupa cases. The wing and antennal distinc- 
tions of the adults may be detected, by close observation, in the portions 
of the pupa case which cover these respective parts of the adult. By 
this means it is possible to segregate males and females before eclosion. 
It is not safe at any time to regard a female as virgin if it is found that 
a male is also present in the culture bottle. However, a newly hatched 
moth always has a tuft of loose, fluffy hair-like scales on its head and 
if these are present, it is a fairly safe indication of the moth's virginity. 

The young larvae upon hatching start eating the portion of the comb 
known as the "midrib," which is the sheet of wax at the base of the cells. 
They bury themselves in the wax and throw out piles of masticated wax 
at the surface. This is a sure indication of their presence in the combs. 
Since offspring of a given female are fairly numerous, it becomes neces- 
sary, as the larvae grow older, to segregate portions of the colony into 
subculture bottles; for instance, if a colony has 300 larvae present, after 
they are about 2 weeks old, it is better to divide them into cultures of 
about 40 or 50 larvae each. Also at this stage, it becomes necessary 
to arrange an aerating device for the culture bottles. For this purpose 
a special ventilator cap has been devised. It consists of two regular 
milk bottle caps with a 1% or 1% inch hole cut through the center of 
each. Then a piece of 40-mesh copper wire screening cut to about the 
same size as the cap itself is inserted between the two perforated caps 
and these layers are stitched together with some wire stapling device.* 
This special aerating cap permits free circulation of air through the 
bottles. The metal screen between the caps is necessary since the larvae 

♦The Hotchkiss or Bostitch letter stapler is very satisfactory for this purpose. 



Pyralidae 351 

otherwise would cut their way through the caps and escape. These 
aerating caps may be used repeatedly. 

Special care must be taken to prevent the food from becoming in- 
fected with wild or stray larvae. The comb, when it is received from 
the apiarist, is sterilized by placing in an oven at 6o° to 70 C. and 
leaving for about 2 hours. The sterilizing process is primarily to kill any 
wild larvae which might be present; otherwise, sterilizing would not be 
necessary since fungus and other disturbing organisms do not attack 
dry bee comb. After sterilization the combs are packed into metal cans 
with close-fitting metal covers. The ordinary pretzel or lard can is very 
satisfactory and easily handled. For the current supply of the comb 
a smaller can is preferable. It may be refilled as needed from the 
supply in the large can. The large storage cans should be kept, in a 
separate room or at least some distance away from the culture bottles 
containing the larvae and the moths. This is necessary since the young 
larvae, if they escape from the culture bottles, may go a considerable 
distance and may chance to get into the stored supply. If several stray 
larvae get into the stored supply and are not apprehended, the next 
generation will likely be sufficient to destroy the whole lot of combs. 

The larvae flourish in a temperature around 30 to 35 C. The regular 
Drosophila incubator, or any other incubator which has a temperature- 
regulating device, is satisfactory for incubating purposes. If an in- 
cubator is not available a very serviceable one may be made of ordinary 
fibrous building board. A convenient size is 4 feet wide, 3% feet high, 
and 14 inches deep. Material 2x2 inches is adequate for the frame- 
work and, if it is lined both inside and outside, leaves a 2-inch dead air 
space in the wall. The shelves should be about 2 inches narrower than 
the space inside and perhaps 2 inches shorter, permitting a free circula- 
tion of air all through the incubator. It is helpful so to arrange the 
shelves that the free space on one shelf is on the opposite side of the 
incubator from the free space on the next shelf above. Also, if a small 
fan is placed in the incubator it keeps the air disturbed and aids in the 
aeration of the cultures. The DeKhotinsky thermo-regulator has proved 
a very efficient and dependable mechanism for regulating the tempera- 
ture. This regulator, attached to a 40-watt electric light bulb for each 
shelf in the incubator, is a very adequate heating device and is easily 
installed. 

The handling and inspecting of Galleria is comparatively simple and 
easy, but different from that of Drosophila. Galleria may not be shaken 
from the culture bottles into an etherizing chamber as in the case of 
Drosophila. The food in the culture bottles is loose and dry, and also 
the moths adhere firmly to the walls or to pieces of the comb. Further- 
more, it is not necessary to anesthetize them in handling and examining 



352 Phylum Arthropoda 

them. Normally these moths move about at night but sit motionless 
throughout the day. Thus, their natural characteristics lend them ex- 
cellently to handling unanesthetized. The apparatus for handling con- 
sists of the "moth dipper," a quantity of % x %-inch shell vials and of 
the cardboard milk bottle caps. The dipper may easily be made by 
gluing one of the % x %-inch vials to a wooden tongue depressor, or 
any strip of wood whittled down to about % x % x 6 inches. It is used 
to "dip" the moth from a piece of comb, or from the walls of the culture 
bottle. This is done by gently placing the dipper over the moth and 
moving it slightly. This generally will cause the moth to shift in one 
movement to the side or bottom of the dipper and remain there, and it 
may thus be moved to any part of the room or to the binocular for 
examination. If the dipping causes the moth to become irritated, one 
may slide the dipper up the side of the culture bottle to its top, then 
slip the thumb of the same hand over the mouth of the dipper, thus 
confining the moth. It has proved to be a saving of time to transfer the 
irritated moths to the % x % vials and invert each of these on a milk 
bottle cap. In a few minutes the moths ordinarily will come to rest on 
the milk bottle cap. Once the moth becomes quiet the vial may be 
removed and the moth taken to the binocular or handled in any way 
desired. 

LABORATORY BREEDING OF THE EUROPEAN CORN 
BORER, PYRAUSTA NUBILALIS* 

CONSIDERABLE time has been devoted to the development of a 
technique for rearing larvae of the European corn borer, Pyransta 
nubilalis, in quantity. This work was necessary for the breeding of 
parasites working on the first instars of the borer, such as Microgaster 
tibialis. 

All material is incubated at a temperature of 85 ° F. with a relative 
humidity of 60% or higher. This has been found to provide optimum 
conditions of propagation. 

Full grown larvae collected in the field are used to rear moths for egg 
production, since all the young larvae reared in the laboratory are used 
in the parasite breeding. The cages that are now used for this purpose 
have proven to be very satisfactory and are, at the same time, very simple. 
They consist of lantern globes closed at both ends with fine mesh wire 
screening to facilitate air circulation, and strips of corrugated cardboard 
one inch wide which are placed side by side in a single row across the 
diameter of the globes so that they form a partition. 

The full grown borer larvae are put into the globe and crawl into the 

♦Reprinted, with slight changes, from Canad. Ent. 61:51, 1929. by L. J. Briand, 
Parasite Laboratory, Belleville, Ontario. 



Pyralidae 



353 



corrugations, which open to the sides, where they remain to pupate. 
Not more than 200 larvae should be put in each cage. Before being 
placed in the incubator the globes and their contents are immersed in 
water for from five to ten seconds. All excess water is drained off 
and the water absorbed by the cardboard supplies sufficient moisture for 
pupation. 

During the time of incubation it is necessary to look the material 
over two or three times a week and remove the dead larvae which would 




LI- 



B 




D 

Fig. 67. — Oviposition cage for Pyrausta nubilalis. A, end view: b, frame; c. wood 
disc closing end of cage. B, front view with lower roller removed: d, upper roller; 
e, second roller; f, cylinder forming the cage; g, wire rods (4); w, wire screening; i, 
wire screening cylinder for feeding cotton. C, inside end view of cage. D, lower 
roller (hinged): r, wire rod. 



otherwise contaminate the healthy ones. The mortality among larvae 
varies between 5% and 10%. When overwintered larvae are used the 
moths begin to emerge in three weeks' time, but if using fresh larvae 
collected during the late summer and fall a much longer period of in- 
cubation is required. Emergence extends over a period of about 2 weeks. 
A cage for oviposition (Fig. 67) consists of a cylinder 12 inches long* 
and 5 inches in diameter suspended in a small frame (A, b) in a hori- 
zontal position by means of a central axle on which it revolves. The 
frame also supports a series of three rollers which hold the waxed paper 
in place. The cage is so constructed that its sides, which represent the 
greatest surface, are of waxed paper. The ends of the cage (A, c) are 

♦This dimension must correspond to the length of the roll of waxed paper used: not 
all rolls are 12 inches long. 



354 Phylum Arthropoda 

made of two discs of laminated wood % inch thick and 5 inches in 
diameter held apart by four stiff wires (B, g), and when the waxed paper 
is placed around the wires it forms an oblong cage, 12 inches long by 3% 
inches square. The ends, which expose the smallest surface, are covered 
with wire screening (C, and B, w) in order to keep the moths from lay- 
ing eggs on that part of the cage. They are also provided with a wire 
screen cylinder (B, i) 1 inch long and 1 inch in diameter to hold a 
feeding cotton which is kept moistened with water. When putting paper 
on the cage a roll of waxed paper is placed on the top roller (B, d), the 
paper is then passed over the second roller (B, e), run around the wires 
and brought out over the lower roller (D), the loose end being held in 
place by a clip to the wire rod (D, r). It is so arranged that the paper 
containing the egg masses is pulled out and replaced in the same opera- 
tion. The lower roller (D), which is hinged and removable, serves as a 
door to the cage. When opened it gives an opening 2% inches wide, and 
when closed against the second roller the distance between them is just 
enough to let the egg masses on the waxed paper go through without 
being crushed. 

The newly emerged moths are picked from the globes every day and 
put in the oviposition cages. The operation requires a light which is 
placed on the side opposite to the opening of the cylinder so that the 
moths, which are positively phototropic to artificial light, are held in the 
cylinder by the light through the waxed paper and very few fly out. 
About 40 females and 30 males are put in each cage. If kept in the in- 
cubator the females begin to lay 4 or 5 days after emergence and they 
live from 10 to 12 days. The dead moths are taken out of the cage every 
day and replaced by newly emerged ones. Since the moths are more 
active in the dark it is necessary to cover the cage with a heavy, dark 
paper. 

The eggs are taken out of the oviposition cages every morning and 
left in the incubator 2 or 3 days until they are ready to hatch; then they 
are cut from the paper and placed in 4-inch flower pot saucers with 
food. To make a container one saucer is inverted on another and, in 
order that they may fit closely, their edges are ground until an even 
surface is obtained. Before being used the saucers are immersed in 
water for 24 hours; this method has proven to be satisfactory since 
a large quantity of water is absorbed by the pottery which supplies suffi- 
cient moisture for the hatching of the eggs. Twenty egg masses are 
put in each container with food and then the containers are placed in the 
incubator. They are kept in the incubator until the larvae have reached 
the proper stage for parasitism, which takes about 4 or 5 days. During 
that period it is necessary to examine them daily and replace food ma- 
terial when it begins to decay. 



PyraUdae 355 

After the 3rd instar it is necessary to isolate the larvae on account of 
their cannibalistic habits. They are, therefore, fed in 2-inch glass vials, 
wire screen stoppers being used in order to provide for ventilation. 

A large number of foods have been experimented with including the 
following: curled dock (Rumex crispus), string beans (yellow and green 
pods), celery, corn, beet, turnip, rhubarb, apple, cabbage, chard, potato, 
corn flakes, lima bean, and sprouted grains. Of these the first named 
have proven most satisfactory with others in the order given. 

Reference 
For the breeding of Pyransta nubilalis see also p. 512. 

REARING EPHESTIA KUEHNIELLA LARVAE IN QUANTITY 

P. W. Whiting, University of Pennsylvania 

Equipment Needed. A warm room, 25 ± C, for starting the cater- 
pillars; a cool room, 20 — C, for slowing development of the cater- 
pillars; a refrigerator for cold storage of full-grown caterpillars; wash 
boilers or other convenient, covered vessels for raising humidity; paste- 
board boxes (wire-stitched, not glued) of convenient size, 8" x 5" x 4" 
high, giving a fair amount of floor space; glass tumblers or tin cans; a 
few shell vials, 70 x 20 mm., for collecting the moths. 

Directions. Hold the box or cover with moths resting on it in the 
left hand. Collect the moths in a vial held in the right hand. The vial 
must be held approximately upright while the vessel containing moths 
is adjusted so that the moth will drop down into the vial. There is 
thus little danger that the moth will escape. Twenty-five to fifty moths 
may be collected in one vial. If the moths fly actively they may be 
quieted by cooling the box in the refrigerator or on a window sill in cool 
weather. A pasteboard box with the bottom spread thinly with rolled 
wheat (Pettijohn's Breakfast Food), 1 cup per 40 square inches, is 
opened and the cover held in the left hand. The vial of moths, held in 
the right hand, is emptied into the box by a quick motion so that the cover 
may be replaced before the moths escape. 

Boxes containing moths are placed in covered boilers in which one or 
two glasses of water are set. The boilers are then left for about 2 weeks 
in a warm room. Boxes should then be inspected and the webbiness of 
cereal and the presence of young caterpillars noted. Add more cereal 
according to the needs of the larvae. The advantage of having a small, 
measured amount at first (1 cup to 40 square inches) is that thus the 
number of caterpillars may be better judged. Enough food should be 
present so that all larvae have plenty, but the whole mass should become 
matted and the cereal for the most part be consumed. 

After the boxes have been in the boilers about five weeks they should 



356 Phylum Arthropoda 

be removed so that the water of metabolism may escape and thus mold 
be avoided. Some boxes may be retained in the warm room for more 
rapid growth of caterpillars and to obtain a further supply of moths. 
Larvae pupate in cocoons in the cereal or on the cover of the boxes. 
Moths will emerge in four or five weeks after the two weeks' inspection. 

If the boxes are placed in a cool room after the two weeks' inspec- 
tion, pupation is delayed but growth continues so that the caterpillars 
attain much larger size. Boxes may be placed in a refrigerator, 5 or 
6° C, and used when desired. 

Dangers. Rats or mice, if present, will gnaw into the boxes, eating 
cereal and caterpillars. 

Beetles or mites may infest the cereal and eat the moth eggs. Habro- 
bracon or other parasitic wasps may attack the caterpillars. Protozoan 
and other diseases may infect the caterpillars. All of these dangers may 
be avoided by using clean cereal, by using new boxes or by returning 
the boxes to the warm room only after sterilizing in a hot oven, and by 
removing from the warm room all cultures showing any signs of infec- 
tion. 

"The White Plague." Chalky, opaque, white caterpillars are full of 
Coccidian spores, which may be seen as fusiform bodies if the cater- 
pillar is macerated on a slide and placed under a microscope. 

"The Black Death." Black spots may appear on the caterpillar 
or the whole body may become black and disintegrate with a foul odor. 
The infection may appear in one corner of the box while caterpillars 
in other parts may be unaffected. Wash and sterilize hands before 
handling cereal in uninfected boxes after contact with infected ones. 

Mold. If cultures become very moldy, caterpillars will not develop 
well. Avoid by lowering humidity and by disposing of infected cul- 
tures. When caterpillars were reared in tin boxes the danger from mold 
was much greater and the water of metabolism sometimes condensed in 
crowded cultures to such an extent that caterpillars would drown. Even 
when there was no mold, caterpillars did not grow as well in tin boxes. 

Failure of cultures to set may be due to extreme aridity. It is im- 
portant to have humidity high when cultures are being started. This is 
conveniently arranged by means of covered boilers and glasses of water 
as above directed. In summer and in humid regions it is unnecessary to 
use boilers. 



Saturniidae 357 

Family saturniidae 

REARING POLYPHEMUS MOTHS 

R. W. Dawson, University of Minnesota 

THE polyphemus moth is for the most part easily handled under 
laboratory conditions. It is short-lived, does not feed, and readily 
and promptly deposits its full quota of eggs in any cage without the 
stimulus of appropriate food plants. The eggs are relatively hard-shelled 
and may easily be removed by hand and placed in small boxes for in- 
cubation. 

Care oj Moths. The chief difficulty in caring for the moths lies in 
getting them to mate in captivity. The method of clipping the wings of 
a recently emerged female and placing her in the open on a small tree 
or a bush during the season of flight of the species is attended by the 
highest per cent of success in securing matings. Caging both sexes to- 
gether in a large screen enclosure out-of-doors is the next best procedure. 
If this cannot be arranged smaller cages by an open window should be 
tried. Only a relatively small number of matings will occur under un- 
modified laboratory conditions.* 

Care of Eggs. Eggs are best collected from the cages daily and kept 
in small tin boxes. Beginning about the sixth or seventh day fresh pieces 
of leaves should be placed in the boxes each day to bring the relative 
humidity up to a favorable level. Frequent changing and cleaning of 
the boxes is necessary to check molds and bacteria which might other- 
wise develop in the moist, enclosed chamber. From eight to fourteen 
days will be required for incubation at ordinary temperatures. 

Care of Larvae. The most successful method of starting newly hatched 
larvae is to keep them in tight containers, glass jars or tin boxes, for 
the first 24 hours with fresh leaves of the intended food plant. The 
wandering instinct seems to weaken during this time and the larvae 
settle down to feed. Glass cylinders, open at the ends, are excellent 
for confining the larvae during their early stadia, and sometimes longer 
if they are not crowded or if they are being reared at low temperatures. 
Ventilation must be provided by covering the top of the containers with 
open-meshed cloth or screening. Larger larvae are best confined in screen 
cages. A remarkably satisfactory cage may be made by taking window 
screening and cutting it into appropriate lengths, bringing the cut edges 
together and tacking them to a strip of wood. The lower selvage edge of 
the cylinder so formed will fit close to the floor or table so that no 
bottom for the cage will be needed. The upper selvage edge will make 

♦Editor's Note: W. T. M. Forbes states that mating takes place normally about 
2-3 A.M. and that if a flown male and a fresh female are held together at that time they 
will usually mate. It has been known to succeed even earlier in the evening. M. E. D. 



358 Phylum Arthropoda 

an even support for a loose lid, best made with a loop of heavy wire 
covered by some open-meshed cloth. If need be the lid may be weighted 
to keep a tight contact with the cylinder. Such a cage is easily and 
quickly prepared at a low cost, gives perfect ventilation, and affords 
the maximum facility for cleaning and reprovisioning. If several cages 
are to be used the diameters should be in a graded series so that 
the cages will telescope one inside the other for storage until again 
needed. 

A few items of the utmost importance in rearing the larvae should be 
noted. Food should be furnished either by growing plants, or by twigs 
kept fresh in jars or bottles of water. In the latter case the water sur- 
face must be blocked off or the larvae will drown themselves. Most 
food plants will not support their full quota of leaves on twigs kept in 
water. If possible choose a species of plant that keeps well when cut. 
For T. polyphemus basswood, hazel, dogwood, and birch are especially 
good. In any case, always reduce the leaf surface to one-half or one- 
third the normal. The leaves also keep better if washed or dipped in 
water. This procedure removes dirt and supplies the larvae with a cer- 
tain amount of drinking water comparable to that available in nature in 
the form of dew and rain. That water is important for their best de- 
velopment is evident from the greed with which they drink when the 
above precaution has been neglected. On one occasion the writer ob- 
served the drinking of 25 drops of water in succession by a large T. poly- 
phemus larva that had been reared without drinking water. 

All twigs, whether their leaves are largely consumed or not, should 
be discarded on the second day, or at the latest on the third day, and 
the larvae transferred to fresh foliage. This procedure is a fundamental 
factor in avoiding diseases among the larvae. The only other factors of 
parallel importance are proper ventilation of the cages, and the imme- 
diate destruction of all abnormal larvae. The transfer of larvae to fresh 
food may be quickly accomplished by closely trimming the leaves or 
stems upon which they are clinging with the scissors, and laying them 
on the new leaves. This procedure is necessary because the larvae will 
not move from the old twigs to fresh ones promptly of their own accord, 
and may not be transferred forcibly without injury. 

As the larvae come to maturity their droppings become large and moist. 
Finally a large amount of partially digested food and fluid is evacuated. 
Immediately after this the larvae seek a place to spin their cocoons. The 
great majority will begin to spin upon the food plant. As soon as well 
settled they should be removed from the cage and the leaves and twigs 
supporting the cocoon pinned to the margin of the lid covering the cage. 
It is important to keep the emergence end of the cocoon upward until 
spinning is complete — about 2 or 3 days. Otherwise the structure of the 



Saturniidae 359 

emergence end may be modified so that the moth cannot later escape 
from the cocoon. 

Care of the Cocoons. The cocoons are best kept out-of-doors in woven 
wire or screen cages if they are to overwinter. Otherwise the pupae are 
injured or killed by a shortage of moisture, and without exposure to cold 
do not usually come out of hibernation satisfactorily. Refrigeration is 
possible in keeping the pupae over winter. On one occasion the writer 
kept thirteen hundred cocoons in a refrigerator room where the humidity 
was high and the temperature held constantly at the freezing point. 
Ninety-eight per cent of emergence occurred the following spring, mostly 
during a brief period of about two weeks. Refrigeration at about 42 ° F., 
however, sometimes proves highly fatal to the pupae, and should be 
avoided. Apparently this temperature is too high for complete 
dormancy and too low to sustain development. 

Reference 
For the rearing of Hemileuca, Automeris, Tropaea lima, and Callosamia see p. 365. 

BREEDING LYMANTRIID AND SATURNIID MOTHS* 

William Trager, Rockefeller Institute for Medical Research 

EGGS and pupae of lymantriid and saturniid moths may be readily 
obtained from professional collectors. 
Eggs are placed for hatching in small covered glass dishes (diam. 7 
cm., volume 150 cc). The newly hatched larvae are kept on twigs of 
food plant placed in the same dishes, the dishes being inverted over pieces 
of blotting paper to absorb excess moisture and facilitate removal of the 
excreta. The larvae are transferred to fresh leaves as often as the old 
leaves dry out. In transferring, they are gently shaken off the old leaves 
on to the fresh ones and are handled as little as possible, and then only 
with a soft brush. Larvae which cling tightly to their substrate, and those 
about to molt, should be put on fresh food together with the leaf to 
which they are attached. After the first molt the caterpillars are put 
in larger containers and it is well to put them in wire gauze cages as soon 
as they are large enough to be unable to escape through the mesh. The 
larvae may also be kept outdoors on growing plants surrounded by suit- 
able cages. When kept indoors, the trouble of providing a constant 
supply of fresh food may be somewhat reduced by placing twigs of the 
food plant in a bottle of water with the opening plugged with cotton to 
prevent the caterpillars from crawling in and drowning. Leaves supplied 
in this way must be changed at least every other day. As the larvae get 

♦Abstracted from "Die Zucht der Lymantriidae und Saturniidae'' by K. Pariser in 
"Methodik der wissenschaftlichen Biologie," T. Peterfi, Verl. Julius Springer, Berlin, 
1928, Bd. II, S. 290-300. 



360 Phylum Arthropoda 

larger, mass-cultures should be divided up into smaller ones to reduce 
the chances of disease epidemics. When the caterpillars are ready to 
pupate it is especially important that they should not be too crowded. 

Lymantriidae* The nun moth (Lymantria monacha) and the gypsy 
moth (L. dispar) overwinter in the egg stage. Eggs should be kept 
through the winter in cotton plugged tubes held in a protected place out- 
doors, as the eggs require a prolonged frost period. In the early spring, 
the eggs are placed in an icebox and kept there until leaves of the food 
plant become available. The eggs hatch very soon after being brought 
to room temperature. Crataegus serves as food for both species of 
Lymantria, although L. dispar will feed on oak, fruit trees, pine, and 
larch, and L. monacha on beech and oak. Polyhedral virus diseases are 
the chief difficulty to be overcome in the rearing of Lymantria. The 
spread of these diseases is favored by overcrowding, excess heat and 
moisture, and poor or insufficient nutriment. Breeding cages, before 
being used for new cultures, should be washed with hot water or with 
10% formalin in 50% alcohol, or some similar disinfectant. The pupae 
of Lymantria, when several days old and sufficiently hard, are placed in 
suitable combinations (depending on the purpose of the experiment) in 
glass jars having blotting paper on the bottom and sides, and strips of 
cardboard. The pupal stage lasts 2 to 3 weeks. Copulation and egg- 
laying follow soon after emergence. The eggs are laid on the cardboard 
strips, enabling subsequent convenient handling. 

Saturniidae. The moths of this family overwinter in a cocoon in the 
pupal stage (with the exception of Anther aea yamamai which overwinters 
in the egg stage). Some species (Saturnia pavonia, S. spini, and S. pyri) 
require frost and should be kept outdoors during the winter, while other 
species (of the Samia group) do not require frost and may be kept in 
any cool place. [See also P. 00.] For purposes of mating, the cocoons 
are placed in large breeding cages in the spring. Where special crosses 
are to be made, and it is necessary to know the sex of the insects before 
they emerge, the pupa may be removed from its cocoon, or better, the 
cocoon may be opened just enough to reveal the posterior tip of the pupa. 
Here in the genital region, the two sexes may be readily distinguished. 
The moths mate soon after emergence and the female then begins laying 
eggs. For most of the species the eggs hatch within 10 to 21 days of the 
date of laying. European species are easily reared on Crataegus; An- 
theraea on oak, apple, or hawthorn ; and Samia on Prunus or apple. Like 
the Lymantriidae, Saturniidae larvae are susceptible to polyhedral dis- 
eases, and the same prophylactic measures should be taken. 

♦Editor's Note: W. T. M. Forbes cautions that Lymantria is a very serious pest and 
L dispar should be handled only in areas already infested. L. monacha should not be 
handled at all in this country. M. E. D. 



Noctuidae 361 

References 



Family Lasiocampidae 

For the rearing of Malacosoma see p. 365. 
Family Lymantriidae 

For the rearing of Notolophus, and Hemerocampa see p. 365. 



Family noctuidae 



THE COLUMBINE AND IRIS BORERS 

Grace H. Griswold, Cornell University 

DURING a study of the columbine borer (Papaipema pur pur i fascia) 
and the iris borer (Macronoctua onusta) it was found to be almost 
impossible to tell the sexes apart in living adults, but an examination of 
cast pupal skins revealed that the genital opening in the female pupa is 
further cephalad than is that in the male pupa. This fact made it easy to 
differentiate the sexes in the pupal stage. 

The rearing work was carried on in an outdoor cage. Full grown 
larvae of both species were collected and placed in individual salve boxes 
with damp sphagnum. When pupation occurred each pupa was examined 
to determine the sex and then removed to a jelly glass with the sex marked 
on the cover. Each jelly glass contained, in addition to some damp 
sphagnum, a piece of wire netting about 3 inches long and % of an inch 
wide. This netting rested on the bottom of the jelly glass and extended 
up the side nearly to the top. Almost without exception, every adult 
that emerged walked up the netting and rested there until its wings were 
spread. Since the sex of each moth had already been determined it was a 
simple matter to place pairs of moths in individual cages. 

The most satisfactory cage used for the columbine borers was a 
cylindrical glass cage 12 inches high and 5% inches in diameter with a 
piece of cheesecloth tied over the top. Each cage rested on a circle of 
paper toweling placed in a large shallow saucer. A wide-mouthed bottle 
held the columbine foliage. About each bottle was wrapped a piece of 
wire netting to provide a rough surface on which the moths could easily 
walk. A long strip of absorbent cotton extended to the bottom of the 
bottle, the upper end being wrapped several times around the petiole of 
the columbine leaf. The cotton acted as a wick and the part at the top 
of the bottle was always wet. Thus the moths were constantly supplied 
with drinking water. 

Pairs of iris borer moths were placed in the same type of cylindrical 
glass breeding cage. To insure an adequate food supply each cage was 
also provided with a small watch glass of the Plant Industry type, con- 
taining a s r < solution of dextrose in water. Fitted into the top of each 



362 Phylum Arthropoda 

watch glass was a circle of wire netting to prevent the moths from fall- 
ing into the liquid. Black cloth was substituted for wire netting as a 
cover for the bottles containing the foliage and the wick of absorbent 
cotton. Since the moths of both species are very sluggish during the 
day no difficulty was experienced in moving them from one cage to 
another. 

Although leaves and other debris were placed in the cages of the 
columbine borer, eggs were rarely laid anywhere but on the bottom. No 
debris of any kind need be provided, thus making it easy to find and 
count the eggs. Eggs of the iris borer are placed in clusters of from 2 
to as many as 150, carefully glued down. Although females deposited 
their eggs on dried leaves, they also pasted them on practically every- 
thing in the cage that had a rough or crinkled surface. The females 
evidently prefer to lay their eggs between two surfaces as clusters were 
often placed in folds of cloth or between two pieces of paper toweling. 

Eggs were collected from the cages in the fall and placed in salve 
boxes. These small boxes were buried in dead leaves in large cartons 
and these in turn were kept out of doors all winter. When hatching was 
desired in the spring the eggs were brought inside and placed on the soil 
beside iris or columbine plants growing in pots. Larvae were hatched as 
early as March, although outdoors they did not hatch so soon. These 
larvae found their way to the plants and developed satisfactorily under 
these nearly natural conditions. 

References 

For the trapping of Catocala see p. 337. 

For the trapping of Noctuidae in general see p. 364. 
Family Arctiidae 

For the rearing of Utetheisa, Euchaetias, Hyphantria, Halysidota, and Ammalo 
see p. 365. 



Family bombycidae 



METHODS FOR THE LABORATORY CULTURE OF THE 
SILKWORM, BOMBYX MORI 

William Trager, Rockefeller Institute for Medical Research 

METHODS for the large scale rearing of Bombyx are readily avail- 
able in books and government publications. I will attempt here 
merely to describe the technique used by Dr. R. W. Glaser and those in 
his laboratory in connection with studies on silkworm diseases. 

The silkworm eggs (which may be obtained from a dealer) are kept at 
room temperature in groups of about 50 in small glass dishes. The larvae 
hatch within 10 to 14 clays, and are provided with a mulberry leaf on 



Bombycidae 363 

which they crawl and begin to feed. Only young, tender mulberry 
leaves should be used for the small caterpillars. In all cases clean, healthy 
leaves should be selected, and it is usually well to wash the leaves. The 
mulberry leaf with the young larvae on it is transferred to a shallow tray, 
about 6 inches square with sides i%-inches high, made out of previously 
autoclaved cardboard. No cover is needed. Fresh mulberry leaves are 
supplied and the trays cleaned at least once a day. The young larvae 
are not handled but are removed together with the leaves to which they 
cling. The excreta and dry leaves are shaken out of the tray. The leaves 
with the larvae are replaced and covered with a few fresh leaves, to which 
the larvae crawl. Caterpillars which must be handled directly are best 
taken up with a camel's hair brush. The larval stage of the silkworm 
consumes from 4 to 6 or 7 weeks, depending on the temperature, and con- 
sists of 5 instars. When a group of about 50 larvae enters the 3rd instar 
it should be transferred to a cardboard tray about 1 2 inches square with 
sides 3 inches high. Larvae in the 4th and 5th instars are large and may 
be handled directly. Larvae entering the 5th instar should be thinned 
out to not more than 20 to 30 to a large tray. 

When one is working with a silkworm disease, the healthy stock larvae 
should be kept in a separate room in which no diseased material is pres- 
ent. They should be fed in the morning before diseased material has been 
handled. If it is necessary to go to them again later, one's hands should 
first be thoroughly washed. Observation of these simple precautions pre- 
vents infection of the stock from the diseased experimental worms. The 
washing of the leaves and autoclaving of the cardboard for the trays re- 
duce the chances of infection from external sources. If an occasional 
caterpillar does get sick or dies, it should immediately be removed. If 
a number of larvae in one tray die, this tray with all the larvae in it 
should be discarded. 

For careful studies of disease, the experimental worms should be kept 
singly in % pint bottles, previously capped with paper and sterilized 
by dry heat. When it is desired to feed known quantities of infective 
material with a pipette, the worms (preferably in the 5th instar) should 
be starved for one day. 

The stock may be perpetuated by allowing the full grown worms not 
needed for experiments to spin cocoons. The cocoons are conveniently 
kept in large cardboard trays. The moths, which emerge about 2 weeks 
after spinning of the cocoon, mate readily, and each pair should be placed 
on a large sheet of cardboard under a % pint bottle or a small lamp 
chimney. Eggs laid by the female moths adhere to the cardboard, which 
may later be cut into squares, each square holding the eggs laid by one 
moth. The newly laid eggs are yellow but, unless they are sterile, they 
gradually darken and become gray within 7 to 10 days. When the eggs 



364 Phylum Arthropoda 

are gray and fully embryonated they should be stored in an icebox. Eggs 
of the ordinary univoltine race will not hatch unless they have been kept 
about 2 months in the cold. They may be kept a year in the cold and 
will still hatch when brought to room temperature. Eggs of the Japanese 
multivoltine race hatch within 10 to 14 days after the date of laying and 
require no cold period. They may, however, be stored in the cold with- 
out injury. 

FURTHER NOTES ON BREEDING LEPIDOPTERA 

W. T. M. Forbes, Cornell University 

COLLECTING moths at bait [see also p. 337] is very nearly a special 
method for Noctuidae, although it is successful also for a few mem- 
bers of other families. A diffuse light is better than a torch since there is 
less likelihood of a sudden bright illumination frightening them. When 
frightened many moths dash down or sideways instead of to the light. 

A useful collecting outfit consists of a wide-mouthed (6 oz.) bottle 
with a vial of cotton inverted in the cork. Wet the cotton well with 
ether. If the moths are wanted alive ether should be used and not 
chloroform or cyanide. Catch moths from the bait in the bottle and 
remove when the legs are still twitching. They may then be examined 
safely for sex, etc., and either transferred to pill boxes, freed, or cy- 
anided. The bottle should be kept open between periods of use to 
avoid poisonous decomposition products of the ether. 

In sexing the moths, note that the female has an extensile fleshy ovi- 
positor and the male two chitinous valves which may be seen by squeez- 
ing the abdomen gently. The male has a frenulum running through a 
hook near the base of the fore wing while the female has interlocking 
scales and bristles only. 

Most moths which do not feed in the imago may easily be reared 
for successive generations by the general method described for Lyman- 
triidae and Saturniidae [see p. 359]. Others usually give trouble in 
mating. Moths seek their mates by smell and a slight drift of 
wind may help them find each other. Confining them in a very small 
box may also be successful. Some males need to fly a time before mating 
and should have a large emergence cage. Avoid a strong light from one 
side or moths will congregate there. 

To avoid handling in transferring caterpillars to fresh food, some silk 
breeders use a light frame carrying a coarse net. The fresh food is put 
on this and laid over the tray of caterpillars. When they have climbed 
up to the fresh food the tray is lifted off and the lower tray discarded 
with any larvae too sick to move. At molting times extra time must be 
allowed as they will not then move up to the fresh food. This method 
is used on a commercial scale in Asia Minor and China. 



Bombycidae 365 

Eucleidae. Slug caterpillars spend the winter as larvae in the cocoon 
and need careful protection at that stage. According to Dyar most 
members of this family mate only on the day of emergence and fairly 
large lots are necessary if matings are to be obtained. 

Lymantriidae. Tussock moths, Notolophus and Hemerocampa, may 
be reared by the same technique as the Lymantrias [see p. 360] . They 
also overwinter in the egg stage. Young larvae are extremely active and 
especially fine screening may be needed for cages during the first few 
days. 

Saturniidae. Hemileuca hibernates in the egg stage. Most pupae 
need no protection during the winter, even in a dry room. A few with 
light cocoons (Tropaea luna, Automeris) should be protected during the 
winter [see p. 360]. To determine the sex pupae of this family I 
prefer to make an opening in the side of the cocoon in order to note the 
width of the antenna. The opening may be resealed with a small cover 
glass if it is desired to keep track of the stage of development. 

The following are somewhat specialized in their food preferences: 
Callosamia feeds on members of the family Magnoliaceae and Sassafras; 
C. promethea is less particular and will also eat lilac, etc.; Samia Colum- 
bia feeds by exception on Larix and Tsuga; Hemileuca feeds on oak in 
the east and willow in the west, while Hemileuca lucina prefers Spiraea. 

Lasiocampidae. Tent caterpillars (Malacosoma) are particularly easy 
to breed indoors, even under adverse conditions of temperature and 
humidity and thus form excellent material for the school classroom. 
They give no trouble even with mating. For M. americana the food 
should be supplied in water or a growing plant should be provided. 
When the first food grows stale or is exhausted the twig and tent should 
be removed from the water but not discarded. New food may be put 
beside it and discarded when stale unless it is too much involved in the 
tent. Eggs are laid in early July and need no care during the winter, 
though they should be exposed to the winter weather. 

Arctiidae. Most members of this family are general feeders. Eu- 
chaetias and Ammalo are limited to the Asclepiadaceae and Apocyna- 
ceae; Halysidota needs tree foods as a rule; Hyphantria makes a tent 
and should be treated like Malacosoma, with apple forming a satisfactory 
food; Utetheisa is partial to seed pods of Crotalaria. 

BUTTERFLIES 

John H. Gerould, Dartmouth College 

THE stage of development in butterflies at which experiments in 
genetics are most conveniently begun is the adult. Any captured fe- 
male, in species which I have bred, may be safely assumed to be already 
fertilized, though very likely by more than one male. 



366 Phylum Art hr op oda 

Paper bags, carried into the field and inflated, make useful receptacles 
for gathering live specimens. The wings of large, active species such as 
the "Monarch" should be clamped together over the back with paper 
clips. 

Live butterflies may be sent even across the continent by parcel post 
or express in a large mailing tube or tin can lined with wet blotting 
paper, sewed securely, through punched holes, to the inner side. The 
strap-hanging butterfly should have a good foothold, and there should 
be no loose projecting walls upon which to beat its wings. Never more 
than two or three live specimens may be shipped together, for they stimu- 
late one another and are likely to be smashed to pieces. 

Plenty of pure (unsweetened) water should be used in wetting the 
lining of the mailing tube or other receptacle, which must be kept moist 
throughout the journey. Five days en route is about the maximum. 
For a long trip, a tin container should be packed in excelsior to avoid 
jarring. 

On arrival, the female should be set over a growing food plant, pref- 
erably potted and in a greenhouse, or insectary, to avoid predatory ants, 
wasps, mice, birds, etc. The atmosphere in the greenhouse should be 
moist, or at least never very dry. 

The cage, a wooden frame lined with soft, black netting, should fit the 
potted plant closely enough so that the butterfly will hover close to the 
foliage. She is fed conveniently by a bunch of flowers frequently dipped 
in clean water sweetened slightly with brown sugar to a consistency re- 
sembling that of maple sap. The bouquet should be kept in a tall, wide- 
mouthed bottle close to the plant on which she is to lay.* 

For cultures of Colias (= Eurymus), circular patches of white clover 
turf are cut with a rounded spade to fit a bulb pan 10 or n inches in 
diameter and 5 or 6 inches high. This is then covered by a cage 15 inches 
square and 10 inches high. The frame may be made of pine strips 
i%" x %", supported by corner posts %" square, with the exposed 
vertical edge of each smoothed off. 

To avoid caterpillar diseases, it is of utmost importance to have the 
eggs thinly distributed, with not more than one egg to a leaf. If the 
butterfly is actively flying in the sunlight, there is not likely to be much 
crowding, but the pot should be rotated 180 when the side toward the 
sun is sufficiently furnished with eggs. Six or eight pots of clover may be 
required for the eggs of a single female during the week or fortnight of 
her laying. 

♦Editor's Note: W. T. M. Forbes says that he has had surest results in obtaining 
oviposition (with some risk of killing the butterflies by overheating) by covering the 
butterfly with a glass bell jar with some twigs of the proper food-plant and setting in the 
sun. Also that in many long-lived butterflies there are no mature eggs when the butter- 
fly emerges, and there must be a long period of feeding and flying; this is true of angle- 
wings for instance. M. E. D. 



Bombycidae 367 

If the eggs are at all crowded, dispersion of the larvae as widely as 
possible over the food-plant must be undertaken after the first molt; 
later, when the foliage becomes sparse or covered with aphids, the larvae 
must be transferred to a fresh pot of clover. 

Even though the caterpillars are sedentary, as in Colias, the pots 
carrying a culture must be covered with cages, or, better, with a single 
large cage extending over all the pots of the culture. Such a cage may be 
made of a wooden frame covered internally with fine black netting (white 
is too opaque) and accurately fitting the surface of the table. This 
should be as flat as possible and preferably of cement, so that it may be 
washed frequently with a hose. 

Most caterpillars are subject to diseases corresponding to flacherie 
and pebrine of the silkworm. If a green, naked caterpillar is off-color or 
slightly pale, it should immediately be isolated, for "polyhedral disease," 
resembling flacherie, is highly contagious, and the leaves near the sick 
caterpillar soon become contaminated. If the disease does not kill the 
larva, it slows up development, the suspended caterpillar later droops and 
dies, or the pupa turns black and purulent. 

The healthy full grown larvae usually leave the plants and pupate on 
the walls of the cage, though some will be suspended on the plants. The 
cage, with all the pupae, may now be removed to any laboratory table, 
preferably in a cool, dark room where the emerging butterflies will not 
mate. 

On emergence, it is desirable to mark each one with a letter designating 
the brood and a number indicating the individual. This may be done 
with a stub pen and India ink diluted with 50% alcohol to make it flow 
freely. I regularly mark the ventral side of the right hind wing. The 
date of emergence having been recorded, the butterfly's age is always 
accurately known. The males and females of any brood are now kept in 
separate cages in a cool, dark room with adequate humidity. Once or 
twice a day they should be brought into the sunlight and allowed to feed 
on moistened flowers. 

Mating is easily brought about by placing a few individuals of each sex 
together in a cage of the dimensions mentioned or others 15" x 15" x 15". 
The cage is set upon a square wooden tray (16" x 16" inside), with sides 
about 2" high to prevent the escape of flying butterflies when the edge of 
the cage is lifted. The cage upon its tray is then put into the sunlight; 
pairs are taken out and isolated as soon as they form. Mating lasts from 
a half hour to several hours, usually one or two. The same male may be 
mated again the next day with a different female. This is important 
in testing a male's genetic make-up. Females known to be impregnated 
are thus ready to set over potted food plants, or are kept in reserve for 
use if others prove sterile. A female requires only a single mating. 



368 Phylum. Arthropoda 

Hibernation presents no problem in Pieris, Euchloe, and other butter- 
flies which winter as chrysalids. It is more difficult in Colias spp., which 
usually hibernate as half grown caterpillars. Loss of moisture and 
haemolymph by the chrysalids of these species during the winter usually 
prevents eclosion in the spring; the wings cannot expand. 

The natural method of hibernation, burying the caterpillars deeply in 
moist, sandy soil, sometimes succeeds, if they are placed in boxes or tins 
inverted so that moisture will not collect, and packed in a bushel or more 
of excelsior; but even in a cool, well-shaded excavation they may be 
killed by molds in early spring before clover in the fields begins to grow. 
An electric refrigerator, with atmosphere well humidified, is far more 
likely to give dependable results. 

Order Diptera 
Superfamily tipuloidea 

CRANEFLIES 

J. Speed Rogers, University of Florida 

APPROXIMATELY 700 to 800 species of craneflies, distributed among 
l 4 families* and about 120 genera and subgenera are known from 
North America. Many of the species are rare, restricted in distribution, 
or little known, but a large number are common and widely distributed. 
They often form a considerable element of the insect fauna of stream- 
courses, swamps, marshes, woods, and grasslands where their larvae occur 
in a wide variety of aquatic, semi-aquatic, wet, or moist situations. In- 
deed, along the shaded banks of upland rills and seepage areas and on 
wet, mossy cliffs craneflies are frequently the dominant forms of insect 
life. 

A partial knowledge of the life histories and immature stages of repre- 
sentative species of each of the families and subfamilies and of more 
than % of all the genera has been obtained; but the life histories of 
more than % of the species are wholly unknown, and few of the others 
have been carried through a complete life cycle within the breeding cage. 
The following discussion of rearing and culturing methods is thus limited 
and conditioned by a very incomplete knowledge of the life histories of a 
large majority of the species. 

For the purpose of rearing adults or maintaining cultures, craneflies 
may be divided into five broad and overlapping groups: 

♦Tanyderidae, i genus, 3 species; Ptychopteridae, 3 genera, 7 species; Trichoceridae, 
3 genera, about 12 species; Tipulidae, 3 subfamilies, some no to 115 genera and sub- 
genera and more than 700 species. 



Tipuloidea 369 

I. Immature stages inhabiting rotten wood and fungi: The larvae 
mycetophagous, xylophagous, or nekrophytophagous. 

II. Immature stages inhabiting saturated silt, mud, or sand: 

a. The larvae herbivorous, phytophagous, or geophagous (detritus) . 

b. The larvae carnivorous and predacious (cannibalistic under cul- 
ture conditions). 

III. Immature stages inhabiting wet or damp soil: This group in- 
cludes both grassland and woodland species; the larvae herbivorous 
(chiefly rootlets and leaves in contact with the soil) or phytophagous. 

IV. Immature stages inhabiting wet or damp growths of algae, liver- 
worts, and mosses: A large assemblage of species that overlaps some- 
what with Groups I and II, and markedly with Group V; principally 
from hygropetric or neuston situations; the larvae feeding upon the living 
plants and the accumulations of detritus and micro-flora. 

V. Larvae aquatic, pupae aquatic (Antocha) or semi-aquatic: A con- 
siderable number of genera and species with habitats that range from 
strictly aquatic to those of groups I, II, and IV. Typically lotic and 
lenitic habitats and both herbivorous and predacious food habits are rep- 
resented. 

SOME GENERAL CONSIDERATIONS 

Cultures may be started from either the immature stages or from 
fertile females taken in the field. In most instances (all unless the larval 
habitat is known) it is more practicable to begin with the larvae or 
pupae.* Within each of the groups listed, the various species have more 
or less specific requirements for culturing that are best learned by observ- 
ing the conditions of the larval habitat. 

Except for some of the species in Group V, wide-mouthed glass jars or 
small aquaria with loosely fitting glass lids make satisfactory cages. 
Stacked finger bowls are useful for the smaller species, and for the younger 
larvae of larger species if their contents can receive enough diffused day- 
light to permit photosynthesis (Groups II, III, IV), and if they may 
be stacked within a large aquarium or other glass enclosure where the 
evaporation rate may be held uniformly low. Maintenance of the 
requisite moisture is of first importance. The medium in which the larvae 
live will need to be moist to saturation, and the air in the space above, 

*The following papers give considerable data on the habitats and methods of collecting 

the immature stages: 

Alexander, C. P. 1920. The craneflies of New York: Part II; Biology and phylogeny. 
Cornell Univ. Agric. Exper. Sta. Mem. 38:691. 

1931- Deutsche Limnologische Sunda-Expedition; The craneflies. Arch.' fur 

Hydrobiologie, Suppl. Bd. IX, Tropische Binnengewasser 2:135. 

Rogers, J. S. 1933. The summer cranefly fauna of the Cumberland Plateau in Ten- 
nessee. Occas. Papers Mus. Zool. Univ. Mich. 215:1. 

■ ■ 1933- The ecological distribution of the craneflies of northern Florida. Ecol. 

Monog. 3:1. 



370 Phylum Art hr op oda 

into which the adults will emerge, should have a relative humidity of 
from 85% to 100%. Marked or rapid changes in temperature, aside from 
any deleterious effect upon the craneflies and their food material, cause 
much difficulty and frequent losses from their effect upon moisture con- 
ditions. Condensed moisture on the sides of the jar traps and kills the 
adults, and, falling on the larval medium, damages its surface and may 
drown larvae or pupae. This is particularly true when the medium is 
silt or agar. Exposure of the rearing jar to direct sunlight is especially 
to be avoided. 

Except for larvae from markedly acid or basic habitats (sphagnum 
bogs and swamps or seepage from limestone cliffs) considerable varia- 
tions in pH are tolerated. Since it is necessary to keep the jars closed 
to maintain a high humidity, it is advisable to have a comparatively large 
air space in proportion to the space filled with the larval medium, and to 
include a few small green plants (usually mosses, liverworts, or algae) for 
photosynthesis. 

All predacious larvae are strongly cannibalistic in rearing jars, even if 
well fed, and should be isolated. Stacked finger bowls or jelly glasses, 
the centers of the tin lids of which have been replaced by fine meshed 
wire gauze, make good culture dishes. The younger instars of many 
species of Tipula are also somewhat cannibalistic when crowded and, since 
the females lay from 200 to 300 eggs, are very likely to be crowded if the 
full complement is oviposited in one jar. However the survivors appear 
to thrive upon such a diet and, if the culture is well supplied with food, 
the number of larvae is only reduced to the proper population for the 
space provided. 

For a number of species in groups I, II, III, and IV, it has been pos- 
sible to maintain a continuous culture of successive and overlapping 
generations in one large rearing jar with no more attention than to 
maintain proper conditions of moisture and food. For a maximum pro- 
duction of individuals, however, it is better to start one or more new 
cultures from each mated female. 

If sufficient space above the larval medium is provided in the rearing 
jar (500 cc. or more for large species, 4,000 cc. for the larger Tipula), 
the adults of the majority of species will mate and oviposit. It is usually 
preferable, however, to remove the newly emerged adults to a large glass 
jar, kept humid by a carpet of wet filter paper, where mating and often 
oviposition will take place. If one can obtain recently emerged males 
and females at the same time there is no necessity to provide food, since 
mating and oviposition are usually completed within 48 to 72 hours and 
water is obtained from the wet filter paper. If adults are emerging 
slowly so that individuals of both sexes are not always available, the 
adults may be kept alive for one or two weeks by providing food and 



Tipidoidea 371 

keeping the jar at a moderately low temperature, 12 ° to 20 C. A 10% to 
15% solution of cane sugar, or honey, or the juices pressed from over- 
ripe fruit may be fed in small saturated pellets of filter paper or from a 
drinking fountain made by closing the upper end of a short length of 
small glass tubing. 

Although the females of a number of species will oviposit in or on wet 
filter paper, others require a layer or clump of the same or similar ma- 
terial that is utilized in nature. The species that will oviposit on filter 
paper will oviposit more freely in a more natural medium. If mud or silt 
is required, it is convenient to provide a layer several millimeters deep of 
fine silt that has been washed through a sieve fine enough to retain 
the eggs (a mesh with openings about 0.25 mm.). The eggs may then 
be secured readily by washing the silt again through the sieve. Species 
that oviposit in algae and mosses, on wet rocks or elsewhere, will usually 
oviposit freely in tufts or "ropes" of filamentous algae, about the size of 
a lead pencil, coiled about on the wet filter paper. A considerable num- 
ber of species that normally oviposit in wet rotten wood, fungi, silt, or 
algae, oviposit freely in a stratum of rather soft agar, where the eggs 
are easily seen and from which they are readily removed. 

When adults are scarce one male may be mated with two or three 
females in succession. If males are plentiful this practice is not advis- 
able for, in a number of instances and in several species, a considerable 
proportion of the eggs of the 2nd and 3rd females were infertile. 

GROUP I. IMMATURE STAGES INHABITING ROTTEN WOOD AND FUNGI 

Representative genera, subgenera, or species: Rotting hardwoods (feed- 
ing mainly on mycelia) — Atarba, Elephantomyia, Epiphragma, Gno- 
phomyia, Limonia (Rhipidia) fidelis, Orimarga (Diotrepha) mirabilis, 
Teucholabis complexa, Tipula trivittata. 

Rotting wood and fungi — Limonia (Limonia) cinctipes, L. (L.) rara. 

Fungi — Limonia (L.) globithorax, L. (L.) macateei, Via. 

Rotten wood- and fungus-inhabiting forms are probably the easiest 
of all craneflies to rear and may, in many instances, be carried through 
repeated generations within the breeding cage. Since any sample of 
rotting wood or fungus in which larvae and pupae are found may contain 
the immature stages of still other species (as well as possible predators), 
if one wishes to culture but a single species it is necessary to sterilize the 
habitat material for arthropods without destroying the microfungus or 
algal flora. This is best done by air drying. Part of the material is 
dried for several days or weeks, while the possibly mixed culture is kept 
alive in another portion. The dried material may then be placed on 
pads of wet filter paper or on a layer of wet sand in the rearing jars 
where it will become re-saturated with moisture within two or three days ; 



372 Phylum Arthropoda 

then the selected larvae or pupae may be added. It is necessary to keep 
the filter paper or sand wet, but not submerged, during the life of the 
culture. Since it is probable that the actual food material of nearly all 
wood- and fungus-inhabiting larvae consists of mycelia and other soft 
parts of the mixed flora of fungi (and sometimes algae) that grows on the 
wood or larger fungous body, it would seem logical to re-infect the 
previously dried material with scrapings from the original, undried wood 
or fungus, but this procedure usually seems unnecessary. 

With some exceptions, the precise species of wood or fungus does not 
appear to be a specific requirement. Limonia cinctipes, L. macateei, and 
L. rara have been carried through 4 to 12 successive generations in a 
supply of dried and re-wetted Polyporus tsugae. A large supply of this 
fungus was collected and dried in North Carolina and then brought to 
Florida where it does not occur. In it a stock of L. cinctipes, obtained 
in Florida from a Polyporus sp. on a sweet-gum (Liquidambar) ; also 
a stock of L. macateei, obtained in Florida from Poria sp.; another stock 
of L. macateei, obtained in North Carolina from Polyporus tsugae; and a 
stock of L. rara, obtained in Florida from mycelium-riddled wood of a 
Magnolia log, were all successfully maintained as long as the supply of 
the re-wetted fungus was provided. For L. cinctipes, a new supply was 
required about every two generations ; for the other species, about every 
four generations. 

A "Polyporus agar," made by steeping shreds of fresh Polyporus for 
a week or so in tap water and then boiling, straining, and adding suffi- 
cient dry, plain agar to make the infusion set when re-boiled and cooled, 
forms a medium in which L. macateei and L. rara will oviposit freely; 
the eggs will hatch and the larvae feed and grow. The agar is poured 
into the usual petri dishes and, when set, the cover is temporarily re- 
placed by a lantern globe with screened top, set directly on the agar. 
Mated females, introduced from the rearing jars containing fungus, not 
only oviposit but infect the agar with spores so that mycelia soon pene- 
trate the agar in all directions. It is the mycelia that furnish food for 
the young larvae, and the movements and feeding of the latter may be 
watched under a binocular microscope until the agar becomes opaque 
from fungous and bacterial growth. Transfers of the larvae to fresh 
plates inoculated with smears from decaying fungus may easily be made, 
but if visibility of the larvae is not required they will thrive in the original 
plates as long as the agar neither liquifies nor dries out. Accumulations 
of moisture on the surface of the agar are likely to drown the larvae. 



Tipuloidea 3 73 

GROUP II. IMMATURE STAGES INHABITING SATURATED SILT, MUD, OR SAND 
Representative genera, subgenera, or species: Non-predacious larvae— 
Erioptera (all subgenera), Gonomyia, Helius, Helobia, Molophilus, Ormo- 
sia, Pseudolimnophila, Tipula annulata, T. jacobus, T. sayi, T. subeluta, 
T. synchroa, T. tricolor, Trimicra. 

Predacious larvae — Adelphomyia, Hexatoma (Eriocera) albitarsis, Lasi- 
omastix, Phylidorea, Pilaria, Polymera, Ulomorpha. 

Here, perhaps more than in any other group, silt, mud, or sand from 
the larval habitat is likely to contain the immature stages of several 
species of craneflies as well as various predators upon them. If it is 
only desired to carry late larvae through to the adult stage, the wet silt, 
mud, or sand may be placed in a layer an inch deep on top of a layer 
of coarse wet sand in the rearing jars and pasteurized. When cool, any 
loss of water should be made up and the larvae introduced into the jars. 
If younger larvae or eggs are to be reared, or a continuous culture is to be 
maintained, the silt, mud, or sand should be washed through a wire sieve 
of 12 to 16 meshes per inch into a jar of water. When the material has 
settled* the water may be decanted and the semi-fluid silt or muddy sand 
may be placed in a layer an inch deep on about an equal depth of coarse, 
wet sand in the rearing jars. (A cylindrical glass jar, 150 x 150 mm., 
Y 4 filled with sand and silt, is of ample size for a score or more of Eriop- 
tera larvae.) Some small green plants that will thrive in the wet silt or 
sand should be provided, but not allowed to choke the surface. Small 
pond-margin grasses (Websteria), Hydrocotyle, and silt-inhabiting liver- 
worts and algae have been used successfully in rearing Erioptera, Molo- 
philus and Pseudolimnophila. It is important to keep very nearly satu- 
rated cultures in which fine silt is the medium. Once the surface becomes 
dry it is likely to acquire a texture that prevents the larvae from reaching 
the air with their respiratory disks. 

Predacious larvae require much the same culture conditions but should 
be isolated. Several living Erioptera or Pseudolimnophila larvae, some- 
what smaller than the predacious larva, or some small aquatic or silt- 
inhabiting annelid worms should be dropped into the jar once or twice 
a week. 

GROUP III. IMMATURE STAGES INHABITING WET TO DAMP SOILS 
Representative genera and species: Cladura, Dactylolabis cubitalis, 
Dicranoptycha, Nephrotoma eucera, N. jerruginea, N. macrocera, Tipula 
bicornis, T. borealis, T. dietziana, T. disjuncta, T. dorsomaculata, T. du- 
plex, T. fuliginosa, T. georgiae, T. grata, T. oxytona, T. perlongipes, T. 
triplex, T. triton. It is probable that many Nephrotoma and a large 
majority of the species in the subgenera Oreomyza and Lunatipula of 
Tipula belong here. 

*Small larvae or other arthropods that pass through the sieve usually soon rise to the 
surface of the water where they may easily be seen. 



374 Phylum Arthr op oda 

The only species of this group that I have carried through one or more 
complete life cycles are Nephrotoma suturalis and Tipula oxytona, but 
the others have been reared from early or mid-stage larvae to adults and 
would apparently be no more difficult to maintain in cultures than these 
two. Glass jars or aquaria, about as tall as their diameter and a gallon 
or more in capacity, form good rearing jars for about a dozen larvae. 
Soil from the habitat, crumbled in the hands and sifted through a sieve 
of about 10 meshes to the inch, should be placed in a layer i% to 2 inches 
deep over an inch layer of coarse, damp sand and lightly tamped. It is 
advisable to include all leaf fragments and other small bits of decaying 
vegetation that are found in the sieve. The purpose of the sifting 
is to remove unaccounted-for larvae and the various predatory ar- 
thropods that occur in the soil. A few plants (grasses or small herbs) 
from the habitat should be planted but kept from growing so tall as 
to fill the space above the soil. For Nephrotoma suturalis centipede 
grass, small cabbage plants, or young lettuce appeared to serve equally 
well. The soil should be kept about as damp as or slightly damper than 
the habitat from which the larvae were taken, but gravitational water 
should never be present. A small well, formed by a piece of large glass 
tubing, open at both ends and extending to the bottom of the jar, forms 
a convenient well from which any surplus (standing) water may be 
"pumped" with a pipette, and through which required water may be 
added. 

GROUP IV. IMMATURE STAGES INHABITING ALGAE, MOSSES, AND LIVERWORTS 

Representative species: In algae (including diatomaceous sludges), and 
mosses on wet cliffs, rocks and piling (Fauna Hygropetrica) — Dactylolabis 
montana, Elliptera illinoiensis, Limonia humidicola, L. pudicoides, L. stulta, 
L. canadensis, L. distincta, L. rostrata, L. simulans, Dolichopeza carolus, 
Tipula caloptera, T. floridensis, T. furca, T. kennicotti, T. iroquois, T. 
oropezoides. 

In filamentous algae floating in quiet waters (Infraneuston) — Limonia 
distans, Megistocera longipennis, Tipula caloptera. 

In submerged algae (and mosses) of rocky or gravelly stream bottoms — 
Limonia gladiator, L. iowensis, Limonia sp., Tipula caloptera. 

In mosses and liverworts on damp earth or logs — Limonia divisa, L. di- 
versa, Dolichopeza dorsalis, D. obscura, D. sayi, D. subalbipes. 

A number of the species listed are aquatic or semi-aquatic, from the 
standpoint of an ecological classification, but all may be successfully 
cultured with no more water than is required to keep their food plants 
from rapid deterioration. 

The rearing jars should have a thin layer (5-10 mm.) of sand covered 
with one or two sheets of filter paper, the sand and filter paper being 
saturated, or barely submerged in water. If algae are to be the food 
plants, a small quantity of filamentous strands should be spread one (or 



Tipidoidea 375 

at most a few) filament deep over the filter paper. Mosses and liverworts 
should be strewn in as thin a layer as possible. If the culture dishes are 
placed in a strong north light the plants will remain green and suitable 
for food for a maximum time — several days to a week or more for the 
algae, several weeks to indefinitely for the mosses and liverworts. If more 
than a very few or very young larvae are present, the plants will be 
eaten more rapidly than they will spoil. Frequent inspections are ad- 
visable and fresh material must be added as needed. Accumulations of 
detritus are not harmful unless considerable quantities of plant material 
are undergoing rapid decomposition. Young larvae, in fact, appear to 
thrive best on a thin, brownish-green detritus that accumulates from the 
decomposition of a small excess of plant material. The newly hatched 
larvae of L. distorts, L. rostrata, D. subalbipes, T. caloptera, and others 
do best on detritus from algae or mosses, or on algae, but may be fed on 
mosses after they have reached the 2nd instar. Stock supplies of 
algae, mosses, and liverworts may be maintained in large jars or aquaria 
that are kept in strong, diffused light. Almost any species of filamentous 
or colonial alga that is available appears to be satisfactory for food. 

GROUP V. THE LARVAE AQUATIC, PUPAE AQUATIC OR SEMI-AQUATIC 

Since it is generally tedious and expensive to simulate aquatic, espe- 
cially lotic, habitat conditions in breeding cages, Group V includes only 
the residue of aquatic species that may not be reared or cultured by the- 
methods used for groups II and IV. 

Some or all of the known larvae of the various species of Dicranota, 
Hexatoma, Pedicia, Protoplasa, and Longurio live in the gravel, sand, or 
sandy silt of rill, creek, or shallow river bottoms and margins but mi- 
grate well above the water-line to pupate. The larvae of Dicranota, 
Hexatoma, and Pedicia are predacious and very active. Half grown or 
older larvae of Hexatoma juliginosa, H. aurata, H. tristis, Pedicia in- 
constans, P. johnsoni, and P. paludicola have been carried through to the 
adult stage in rearing jars in which wet sand or sandy gravel was piled an 
inch or more above the water-line on one side of the jar. Water from a 
reservoir was allowed to drip slowly onto this emergent bank, while 
a low water level was maintained in the jar by means of a constant- 
level siphon. These larvae were all provided with tubificid or lumbriculid 
worms for food. 

Tipula abdominalis larvae, 6-8 mm. long and probably early 2nd 
instar, have been carried through to the adult stage in a small artificial 
sand-bottom stream. A stream course with emergent banks was molded 
in coarse wet sand in a cypress trough, 3 feet by 2 feet by 10 inches deep. 
The trough was nearly divided longitudinally by a wooden partition 
so that about 6 feet of stream course, in the form of a "U" was provided. 



376 Phylum A rthropoda 

Tap water was run through the stream in a very slow current and leaf 
drift from natural streams was allowed to form drifts against pilings 
made from skewers. This arrangement was most successful in a shaded 
position out of doors, where it received enough light to permit the growth 
of diatoms and blue-green algae on the submerged leaves. Pedicia al- 
bivitta larvae have lived in a somewhat similar arrangement for over 3 
months and then pupated and emerged, but this species is predacious and 
should be provided with annelid worms or small larvae for food. 

Bittacomorpha clavipes and Ptychoptera rujocincta are easily reared 
in aquaria provided with water from the habitat and containing enough 
sphagnum and coarse plant detritus to form feeding and resting places 
within breathing-tube-reach (5-15 mm.) of the surface. For Ptychoptera 
some of the sphagnum should project above the surface. Bittacomorpha 
has been carried through 3 generations in one such aquarium, the females 
ovipositing while in flight above the surface of the water. Some support, 
such as loosely hung horizontal lengths of thread across the top of the 
aquarium, should be provided for the adults. 

Mature larvae of many, probably most, aquatic forms may be carried 
through to the adult stage in loosely packed, wet, but well aerated, mosses. 
Wet, aerated mosses also form the best medium in which to store tempo- 
rarily and to transport aquatic immature stages. Larvae and pupae will 
remain alive and vigorous for several days in such moss but usually 
die within a few hours if placed in water. 



Family 



CULICIDAE 



METHODS OF REARING, MANIPULATING, AND CONSERV- 
ING ANOPHELINE IMAGINES IN CAPTIVITY* 

Mark F. Boyd, T. L. Cain, Jr., and J. A. Mulrennan, Station for Malaria 
Research, Tallahassee, Florida 

CERTAIN earlier publications (Boyd, 1926, 1930) presented im- 
proved methods for large scale rearing of anopheline larvae which 
depended upon feeding the larvae abundant quantities of Fleishmann's 
yeast, placed in accessible positions in the rearing vessel. However heavy 
larval mortality continued and the resulting imagines were smaller than 
those encountered in nature and unreliable in their biting proclivities. 
This problem has been solved by rearing the larvae in vegetable infusions 
kept at a constant temperature. 

The technique was originally developed with Anopheles quadrimacu- 
latus and has been found equally applicable without modification to the 

♦Arranged from articles in Amer. J. Hyg. 16:832, 1932; Amer. J. Hyg. 16:839, 1932; 
and Amer. J. Trop. Med. 15:385, 1935- 



Culicidae 377 

successful maintenance of a colony of A. punctipcnnis. A limited success 
has been achieved in the culture of A. crucians sufficient to learn that some 
of the procedures necessary to the rearing of A. quadrimaculatus and 
A. punctipcnnis must be more or less modified for this species. 

Colonies have been started by the collection of ova from gravid wild 
females confined in small cages above dishes of water. These were given 
a blood meal every three days as long as their ova were required. 

In the insectary a large tank raised 2 feet from the floor is maintained 
as a balanced aquarium filled with water to a depth of 6 inches and 
stocked with aquatic vegetation and snails. Normally the water is never 
changed, though more is added as replacement is necessary to maintain 
the proper level. A small handful of hay is placed in it once a week, as a 
source of food for the snails. A small amount of lime water is added once 
a month to provide calcium for them, and about 5 grams of ammonium 
nitrate in solution is added twice a year as an extra source of nitrogen for 
the aquatic plants. The aquarium attracts the female mosquitoes as a 
place for oviposition. Oviposition is most abundant along the edges and 
where the horizontal vegetation breaks the surface film. The space be- 
neath the tank serves as a dark, humid diurnal shelter for the imagines.* 

In routine insectary operations, ova are collected from the aquarium by 
skimming the water surface (which is an egg trap for ovipositing females) 
with a cereal bowl. Before skimming, the bowl should be wet to prevent 
the ova from sticking to the sides. The ova are dipped from the bowl 
with a bent spoon, and poured into a folded paper in a funnel. After the 
collection has been made, any larvae present are removed with a pipette, 
and the ova are washed to the bottom of the filter cone by a gentle stream 
of water. 

The surplus ova are stored in a Frigidaire at the laboratory for 2 weeks, 
as an insurance against any accident. The moist paper filters bearing the 
ova are kept in % P mt fruit jars. 

As required but before the end of two weeks the ova are taken from the 
Frigidaire and returned to the insectary to be incubated. The ova are 
washed from the filter paper by means of a pipette into the space within 
a cork ring floating in a small bowl of water. The cork ring prevents 
them from stranding on the sides of the bowl as evaporation lowers the 
water level. They are incubated by floating the bowl in the aquarium in 
the summer, and in the water bath with a temperature of 70 F. in the 
winter. They should be kept in the bowl until they enter the second 
stage, and during this period are fed only on yeast. 

In the outdoor insectary optimum temperature is provided by rearing 

* For further details of the construction and operation of the insectary see: Mark F. 
Boyd, T. L. Cain, Jr., and J. A. Mulrennan: "The insectary rearing of Anopheles 
quadrimaculatus." Amer. J. Trop. Med. 15:385, 1935. 



37§ Phylum Arthropoda 

the larvae in a constant temperature water bath which has capacity for 
6 pans. In the winter months this is heated to 70 F. by contained elec- 
trical heating units. In the summer it is cooled to 70 F. by the coils of 
a Frigidaire refrigerating plant. 

The infusions are made, and the larvae subsequently kept when the in- 
fusion is ripe, in white enameled cream pans, about 1 2 inches in diameter 
and 2% inches deep. 

Early successful culture of the larvae on a large scale is attributable to 
the employment of pans of hay infusion for their rearing, the rich auto- 
genous plankton of which, especially when supplemented by a supply of 
yeast, affords an abundance of food and results in the production of large 
vigorous pupae and imagines. While successful, hay infusions have not 
been satisfactory; their qualities vary widely depending on the hay em- 
ployed; they are from a biological standpoint very complex; and they 
may only be employed for larval nutrition after a lengthy process of 
fermentation. It was observed that pans were satisfactory for larval 
nutrition after the reaction had become alkaline and if a dense growth of 
Paramecium and flagellates occurred. Considerable experimentation 
was carried on in an effort to reproduce these conditions in a simpler man- 
ner. After trying extracts from various vegetables and seeds, as well as 
synthetic media, wheat infusions were found to serve as a satisfactory 
base for cultures and now hay infusions are abandoned. 

Wheat infusions are prepared as follows: One or two ounces of sound 
wheat grains are placed in a beaker which is partly filled with tap water. 
The beaker is then placed over a flame and the water boiled for a few 
minutes. When boiled, about 250 grains are placed in an enamel pan 
with 2 liters of tap water. The pan is placed where it will receive very 
diffused light in the laboratory. After 2 or 3 days when a visible bacterial 
growth is present it is heavily inoculated from a previously prepared 
plankton culture in wheat infusion. After 4 or 5 days' further incubation 
at room temperature nebulous masses of flagellates in descending con- 
vection currents are clearly visible and the pan is ready for the introduc- 
tion of larvae. At the time when a series of pans is prepared a separate 
culture is made to serve for the inoculation of the next series. The 
original mixed culture of Paramecium and flagellates was secured from 
a satisfactory ripened pan of hay infusion. Alkalinity is maintained by 
adding 1 gram of powdered calcium carbonate to each pan and neutrali- 
zation of acid is probably facilitated by the gentle stirring of the settled 
CaC0 3 from the bottom once a day. After being placed in service a 
culture will give from 2 to 3 weeks' service before being discarded. 
Water loss by evaporation should be replaced. If a slimy envelope de- 
velopes about 4th stage larvae, the affected larvae may be placed in a 
1% solution of NaCl for 30-45 minutes and then returned to their pan. 



Culicidae 379 

For both A . quadrimaculatus and A . punctipennis an alkaline reaction 
is required. For A. crucians, the hay infusion pans have been used while 
in the acid phase, and later employed for the other species. The acid 
phase may be prolonged by the addition of small amounts of sugar from 
time to time. 

To start a new rearing pan, approximately 250-300 larvae, with the 
larger stages predominating, are transferred to the infusion by means of 
a teaspoon or a pipette. It is imperative that the larger larvae pre- 
ponderate, in order that they may consume any surface pellicle as it 
forms, thereby preventing undue mortality among the smaller larvae. 
Our experience indicates that it becomes necessary at times, to vary the 
number of larvae from the normal limits for a few days in individual pans 
to maintain a balance between food production and consumption. There 
is no criterion that may be given, whereby an inexperienced person would 
be justified in increasing or decreasing the number. This is a matter 
of experience. 

Pans are operated to contribute a quota of pupae daily. If 250-300 
larvae are maintained, 20-25 pupae will be furnished daily by each pan. 
The places vacated by the pupae are filled by adding about 50% more 
small larvae than the number of pupae removed, which is done to offset 
the mortality. 

The food supplied by the infusion does not appear to be wholly ade- 
quate for satisfactory development. Hence in order to obtain imagines of 
maximum size, it is still found advantageous to supplement the diet with 
Fleishmann's yeast. The yeast is placed on a glass slide which is floated 
about %-inch below the surface of the water by means of a cork float. 
The amount of yeast to be placed upon the slide must be determined by 
the rapidity with which it is devoured. Experience indicates that best 
results are obtained when moderate amounts are used and renewed several 
times each day if necessary. The glass slide and float should be washed 
daily to remove the old yeast cells adhering. 

It is found desirable to employ some type of floatage to secure uniform 
distribution of the larvae in the pans. This reduces the opportunity for 
cannibalism and serves to equalize feeding opportunities. Chaff is 
scattered over the surface or several paraffined sticks or cork strips may 
be floated on the surface. 

As transformation into pupae takes place, they are removed daily with 
a pipette to an eclosion pan of tap water. Some type of floatage must be 
used on the water surface of this pan to keep the pupae spaced apart. If 
this is not done they tend to collect around the edge, where they nervously 
bump into one another and jeopardize the safety of those emerging. 
Broken hay, chaff, or ground cork may be used for floatage. The floatage 
also aids the emerging adults by providing support for the tarsi and 



380 Phylum Arthropoda 

thereby reducing the risk of the adults becoming caught in the surface 
film. 

It is advisable to change the water in the eclosion pan daily to prevent 
pellicle formation, which will kill the pupae. This pan is placed beneath 
a specially built eclosion cage in the form of a screened cone that fits 
over the edge of the pan. It is kept in a cool dark place in the summer, 
and in the water bath in the winter. 

Small pupae indicate that the larvae are undernourished. This may 
arise from either the use of pans with insufficient plankton or from keep- 
ing too many larvae in a pan. Undersized pupae should be killed, as the 
imagines they produce will be useless. 

Early attempts at insectary rearing of anophelines indicated that con- 
siderable space was required for nuptial maneuvering. Mating of indi- 
viduals recently derived from wild stock did not occur in small cages, 
and it was not until the imagines were confined in a cage with dimensions 
of approximately 8 by 10 by 13 feet, that the fertilization of sufficient 
females was secured. In addition to adequate space, mating also requires 
that the breeding stock of the colony be composed of large, vigorous 
imagines. 

A. quadrimaculatus can become well adapted to life in such an artificial 
environment. Up to the present (June, 1936) the Florida colony has 
passed through the 48th generation, allowing an average of one genera- 
tion per month in captivity. This adaptation probably accounts for the 
successful establishment of a sub-colony in a small cage with screened 
sides, 30 inches square and 36 inches high. The imagines originally in- 
troduced were taken from an eclosion cage before mating, about 56 hours 
after emergence and had just previously been fed, the males on glucose 
and the females on blood. In the new environment, the colony has al- 
ready been maintained for several generations. 

The breeding colony requires very little attention other than the estab- 
lishment of a reliable source for blood meals. This is most satisfactory 
when furnished from the person of the technician in charge. One of the 
most important factors in establishing a new colony is to have an attend- 
ant who will conscientiously endeavor to persuade all the females to feed 
as often as possible. The males are fed on raisins continuously kept on 
several small trays, or they may be fed on dextrose solution, which is 
a satisfactory food. Dead imagines are picked up daily to keep out 
fungous and bacterial contamination. 

In Florida, density of the colony is maintained at about 5,000 adults 
in the winter, and decreased to about 3,000 in the summer. This is done 
because we find that mortality is greater in the winter months, while feed- 
ing and egg laying are diminished. A ratio of approximately 2 males to 
1 female is maintained. 



Culicidae 381 

The sand on the floor of the space under the aquarium (diurnal shelter 
for imagines) is kept saturated with water the year around to increase 
the humidity. Periods of excessively dry weather necessitate the flooding 
of the floor of the insectary with water in the summer. By so doing it 
has been possible to reduce the imaginal mortality which follows high 
temperature and low humidity. When the mosquitoes are kept in lantern 
chimneys or glass cylinders, considerable trouble arises from the con- 
densation of moisture on the surface of the interior of the containers upon 
their removal to warm air. In order to keep the mosquitoes from contact 
with any solid surface on which moisture might condense, a type of cage 
has been developed consisting of a bobbinet cylinder stretched on a brass 
frame and tied in position. Only a very narrow ring of brass at each end 
is exposed in the interior of the cage, and this is covered with a narrow 
strip of filter paper which absorbs any condensing moisture. When cages 
become soiled, the cylinders are removed, washed, starched, and stretched 
over glass bottles to dry. Closure of the ends of the cages is effected 
with squares of bobbinet secured by rubber bands. 

For the catching of individual mosquitoes in cages there has been 
devised a special catching apparatus consisting of a test tube secured over 
one tine of a spring forceps and a sliding lid placed over the other tine. 

When the mosquitoes are being removed from a cage with a catching 
tube a special cover is placed over the open end of the cage. This is made 
from a heavy rubber bathing cap consisting of two flat pieces of thin 
rubber cemented along the crown. In the center of each flat piece a slit 
is cut. These slits intersect at right angles and form a self-closing orifice. 
The rubber band securing the bobbinet square over one end of the cage 
containing mosquitoes is removed, and the bathing cap is laid over the 
bobbinet with the slits over the center of the cage. The bobbinet square 
is then drawn out from underneath and the cap is secured in place by the 
rubber band. If the mosquitoes are to be transferred to an empty cage 
with the catching tube, the transfer cage is similarly prepared. The cage 
containing the mosquitoes is taken up in the left hand, with the cap- 
covered-end toward the operator. The catching tube is held in the right 
hand ; it is closed by pressure on the forceps tines, inserted through the 
slit orifice in the cap, and opened. The open tube is then gently placed 
over a mosquito resting on the opposite bobbinet end of the cage. The 
mosquito will fly into the tube, and the open end may be closed by pres- 
sure on the tines and the tube removed. If the mosquito is to be trans- 
ferred, the closed tube is inserted through the orifice into the second cage 
and is opened by decreasing the pressure on the tines. The mosquito is in- 
duced to fly out by gentle tapping. If it is desired to kill the mosquito, a 
pledget of cotton saturated with chloroform is placed over the mouth of 
the tube. 



382 Phylum Arthropoda 

When females are required for experimental purposes they are sep- 
arated from the males by means of the catching tube after transfer from 
the eclosion cage to a small bobbinet cage to be transported to the 
laboratory. The males and surplus females are released in the insectary 
to join the colony. A pledget of cotton moistened with a 10% dextrose 
solution is placed over the top or side of the cage. We have found that 
longevity of the females is greatly increased if they have fed on dextrose 
some days prior to their application to a malaria patient. 

The females are carried to the laboratory and placed in a 20 C. in- 
cubator, where they remain 2 or 3 days, or until they have all had a 
feeding of dextrose. They are then put into a large bobbinet storage 
cage holding approximately 300 mosquitoes and placed in a Frigidaire, 
where they remain until required for use. 

When females are to be infected, they are starved about 2 days before 
they are to be given the infecting blood meal. They will take a feed 
better if they are given their opportunity from 7 to 10 days after eclosion. 

It is desirable for various reasons to infect mosquitoes on their first 
feed. On this occasion they can ingest a larger volume of blood than 
at any subsequent time. Immediately after feeding, the insects which 
fed should be separated from those which did not, and the latter are 
discarded. This separation must be done with great care in order to 
avoid injury to the distended mosquito. 

After mosquitoes have received their initial infecting feeding on an 
infectious human subject, their subsequent nutritional or conservation 
feedings are taken from a rabbit. This rabbit is secured to an operating 
board and the hair is clipped from its side over an area sufficient to 
permit the application of the cage end directly to the skin. We find it 
very important to first apply the mosquitoes to the rabbit the day fol- 
lowing their infective feeding, and to then employ great patience in 
order to persuade the maximum number to feed on the animal. There- 
after during the extrinsic incubation period, they are given an oppor- 
tunity to feed every third day by exposing the caged insects to the 
rabbit for about fifteen minutes. After sporozoites are present in the 
salivary glands of the mosquitoes, the cages are transferred to the 
Frigidaire. They are then only allowed to feed on the rabbit once a 
week. Pledgets of moist cotton are kept on each cage to maintain 
humidity at saturation and to give the insects an opportunity to drink. 
All cages are examined daily for the detection of dead mosquitoes, which 
are removed for dissection. After they become infective, they are stored 
in a Frigidaire at a temperature varying from 2 to 17 C. except when 
they are permitted to feed weekly for purposes of conservation. Mos- 
quitoes are transported to and from the hospital in a picnic refrigerator. 

Insects that are incubating an infection are kept in a cool incubator, 



Culicidae 3^3 

adjusted to maintain as closely as possible a temperature from 20 to 
22 C. Close attention must be paid to the operation of incubators, as 
mortality progressively increases as the temperature ascends above 22 C. 
For their control they should be provided with maximum and minimum 
thermometers which are read and set daily. 

Wooden rings are used as cages for the application of individual 
infectious-mosquitoes' inoculation of patients. The rings have the same 
diameter as the large cage frames and a small rim around the outer 
edges so that the bobbinet squares may be retained by a rubber band. 
One square is dyed black to facilitate observation of the mosquito. The 
rings may be turned out of any dense wood. Their interior should be 
sandpapered, painted white and sandpapered again to present a smooth 
surface. A number of these rings, each with its mosquito, may be 
placed in a copper petri dish can during transportation in the icebox. 

Overcrowding of the mosquitoes in a cage is an important cause of 
injury. The risk increases with a rise in the temperature of storage. The 
best results are secured when about 5 cubic inches of space is allowed for 
each insect. The chilling of mosquitoes during transportation is very 
important to immobilize them and reduce the danger from contusion 
arising from collisions especially when they are full of blood. 

Success requires close and conscientious daily attention to the mos- 
quitoes and in all handling and manipulation they should be treated with 
all possible gentleness. 

Bibliography 
Boyd, Mark F. 1926. A note on the rearing of anopheline larvae. Bull. Ent. Res. 

16:308. 
IQ 30. The cage rearing of Anopheles quadrimaculatus. Amer. J. Trop. 



T 



Med. 9:165. 

A MOSQUITO REARING CAGE 

F. C. Baker, U. S. Bureau of Entomology and Plant Quarantine 

HIS indoor mosquito cage (Figs. 68 and 69) was constructed in the 
spring of 1933 as a modification of the Boyd anopheline breeding 
cage. A brief description of this cage is recorded, not because it is an 
ideal type to be copied in detail, but for three other merits: (1) It was 
fairly successful, both in the maintenance of a cage colony of Anopheles 
quadrimaculatus and in culturing Aedes triseriatus and Culex pipiens. 
(2) The cage and equipment cost only about $25. (3) It may help 
someone else in designing his own artificial mosquito habitat to fit pre- 
vailing circumstances when a greenhouse is available. 

The mosquito rearing cage is located in one of the compartments of 
an insectary greenhouse. It is 4' * IO ' and "' hi S n - Its len S tn 
extends in an east- west direction. On the south and west, 3' of its height 



3 §4 



Phylum Arthropoda 



is concrete wall, 4' is glass sash, and 4' is beaver board. On the north 
side the wall comprises the bottom 1.5' of its height; the center 5.5' is 
covered with 16-mesh, iron wire gauze; while the top 4' is beaver board. 
The east end is covered by the same kind of mosquito netting for a 
distance of 7' from the floor; then beaver board for the top 4' of the 

MOSQUITO REARING CAGE 




Fig. 68. — North and south side elevation of mosquito rearing cage. 



wall. The ceiling is beaver board. There are opaque curtains that are 
hung just above the glass or screen on all sides. These may be rolled 
down from the outside, so that light intensity may be fairly accurately 
controlled. The screen door swings outward. Inside, it is guarded by 
three, weighted cheesecloth curtains. 

Across the west end and about 3 feet from the floor level is a perma- 
nent shelf that is 2.5' wide. Its under surface and nether walls are 
blackened. The opening to the space under the shelf is partially closed 
by a concrete foundation wall which extends upward for about half of 



Culicidae 



385 



the distance from the floor to the free edge of the shelf. The bottom 
of this pit is covered by a layer of moist white sand. It was designed as 
a dark, humid, quiet resting place for imagos. From the 1.5' fore-wall 
that is below the edge of the shelf, to the east end of the cage is a 

MOSQUITO REARING CAGE 



w 



J u - s 


P?rm»nen\ 


SUeH /- 


3 /* s — <s 


\ Mov»1 


J Sa. nc\ 


" vv&U S 


a 






6oa,rA 


a 


floor "■ 


FoU\- 








3 


ll 






: 


TanK 


[ 


Uj 


aj 



■ I N 




Plan 



L S t W West 

End Elevation 
(from the East) 



Fig. 69. — Plan and end elevation (from the east) of mosquito rearing cage. 

wooden floor. Across the east end is a 44 gallon wooden tank, the 
bottom of which is 3' above the floor. It has a free water surface of 
6.4 square feet. Four feet directly above the vat is a water vaporizing 
nozzle that is similar to those used on vegetable stands. It is kept 
continuously in action. That fraction of the water jet which is not 
beaten into a mist is conducted out of the cage and allowed to flow over 
the walk of the greenhouse compartment. The element of the mist that 
does not evaporate falls into the tank. The overflow from the vat passes 



386 Phylum Arthropoda 

through a small pipe to the west end where it moistens the sand of the 
grotto. A few potted plants usually are kept on a shelf which is within 
range of the humidifier. It is felt that vegetation adds something to 
the naturalness of the artificial habitat in the cage. 

LABORATORY BREEDING OF THE MOSQUITOES, 
CULEX PIPIENS AND C. FATIGANS 

Clay G. Huff, University of Chicago 

MOSQUITOES of the genus Culex have been bred chiefly for 
studies on the transmission of avian malaria. They have sub- 
stituted for Anopheles and human malaria in many studies where the 
breeding of Anopheles or the experimental inoculation of human malaria 
has proved impossible or very difficult. When more research has been 
devoted to the laboratory breeding of mosquitoes it will very probably be 
found that they are adaptable to many other kinds of research. 

The chief difficulty encountered in trying to adapt either Culex pipiens 
or C. jatigans (=C. quinquejasciatus) to laboratory conditions is the 
failure of the adults of the first generation reared in the laboratory to 
copulate. This is true to a much lesser extent of C. jatigans than of 
C. pipiens. When adults are reared from larvae which have been either 
collected in nature or hatched from wild-caught females, the second 
generation females will readily engorge on blood and will oviposit, but 
only in a very small number of cases will their eggs hatch. This indicates 
that copulation has probably not occurred. It should be stated, however, 
that this is the case when the adults are kept in small containers (no 
larger than lantern globes) . It seems likely that copulation would occur 
frequently in large insectaries such as those employed by Boyd for 
Anopheles. The only method of overcoming this difficulty has been that 
of breeding entirely from the very few mosquitoes which did copulate 
in captivity. At times I have been fortunate in securing a raft of fertile 
eggs from a comparatively small number of females, but at other times 
many thousands of egg rafts have been secured before any fertile ones 
would be encountered. In all cases, however, in which a strain of 
mosquitoes had been secured from females which had mated in captivity, 
no further difficulty was met in securing copulation of the adults of the 
succeeding generations. 

FOOD REQUIREMENTS OF ADULTS 

Adults of these two species will survive for fairly long periods on many 
different fruits, but the one which has proved most satisfactory in the 
laboratory is cooked raisins. The large varieties are most satisfactory. 
A beaker is half filled with raisins and covered with water. They are 



Culicidae 387 

boiled until almost dry and the water replenished and the boiling con- 
tinued until the raisins are suspended in their own syrup. These may be 
kept in a refrigerator in good condition usually until they are all used 
up. One or two of these large raisins placed on top of the gauze netting 
of the breeding cage, if moistened each day, will usually not need to be 
replaced for 4 to 10 days. 

Although it has been shown (Huff, 1929) that females of C. pipiens 
will ovulate and produce viable eggs upon diets other than blood (such as 
egg yolk, ox gall, and potato, carrot, and apple juices) as well as the 
various fraction's of blood, it is advisable to feed them upon living 
animals if one wishes to use them or their progeny in blood feeding 
experiments. 

Gravid females require the free surface of water, or completely sat- 
urated substances such as cellucotton, for oviposition. For maintaining 
stock I use lantern globe cages placed over crystallizing dishes into 
which the globes fit snugly (a dish 90 mm. in diameter and 50 mm. deep 
is required for most makes of globes). For isolation of progenies the 
females may be placed, one each, in standard-sized bacteriological test 
tubes Y 3 filled with water and plugged with cotton. Oviposition occurs 
much earlier and in a larger number of cases when test tubes rather than 
larger cages are used. The period elapsing between the blood meal and 
oviposition may also be appreciably shortened by selection of those 
progenies derived from the eggs earliest laid. 

The feeding and care of the larvae is the most difficult part of the 
whole technique of caring for these two species. The larvae may be 
grown satisfactorily in white enameled pans or the square refrigerator 
dishes which nest one in the other when many different lots must be 
stored in a small space. Although these mosquitoes normally live in 
foul water, great care must be exercised to prevent the cultures from 
becoming too viscous and turbid when grown in the laboratory. This 
precaution is much more important, too, while the larvae are young. 
By judging the amount of food carefully it is often possible to rear a 
brood of larvae to the pupal stage without changing the water. As they 
grow, the amount of food should be greatly increased. If the pabulum 
becomes viscous or a surface pellicle forms upon it, the larvae must be 
transferred to fresh water. This is most conveniently done by pouring 
the entire contents of the pan upon a piece of fine-meshed bolting silk. 
The larvae are thus filtered out and may be transferred to the clean 
water by turning the bolting silk over onto the surface of the water. 
A variety of substances may be employed as food, such as banana, old 
protozoan cultures, and dehydrated blood serum, but the one found most 
satisfactory is dehydrated, skimmed milk. This is available at all times 
in the same condition and may be administered in carefully graded 



388 Phylum Arthropoda 

amounts. It should be added daily from the time of hatching until 
pupation, in very small amounts at first and then in increasing amounts 
as the larvae become able to keep the water clear. The amounts needed 
must be learned by experience. 

When the pupae appear they are removed daily by means of a pipette 
with a large opening. They are placed in containers exactly like the 
ones previously described for oviposition. Successful emergence from 
the pupal stage requires that the surface film of the water be quiet, clean, 
and free from oils. An example of the extreme susceptibility of pupae to 
oils is afforded by my discovery at one time that eclosion could not occur 
in a room containing pigeons. The dandruff or scales from the feathers 
of these birds which settled from the air upon the surface of the water 
disturbed the surface tension sufficiently to effect this result. 

The newly-emerged adults may be kept in lantern globe cages placed 
over petri dishes into which pads of cellucotton have been fitted and 
then moistened. While adults of C. pipiens and C. jatigans will live and 
grow in a wide range of temperatures, they thrive best at about 80 ° F. 
Although they will probably live longer at the lower temperatures used 
by Boyd for Anopheles, they will live for two months or more at 8o° F. 
This, of course, is an advantage, since a cool incubator is not required for 
storing them. The females may be fed upon birds or other experimental 
animals by allowing them to bite directly through the gauze netting of 
their cages. Separation of the unengorged from the gorged females may 
be accomplished by catching them singly by means of a small vial from a 
catching bag made of netting and provided with a sleeve. I have for 
some time been anesthetizing them with ether in the globe cages and 
then separating them quickly by picking them up carefully by the wings 
or legs with small forceps having very flexible points. If the minimum 
exposure to ether sufficient for immobilizing them is used they very 
quickly recover from the anesthesia and show no harm as a result of it. 

When attention is directed to the essential requirements of these 
species it will be found that they may be grown with ease. Indeed, the 
simplicity of the task is the chief argument for employing them in many 
types of experiment. 

Bibliography 

Huff, C. G. 1927. Studies on the infectivity of Plasmodia of birds for mosquitoes, 
with special reference to the problem of immunity in the mosquito. Amer. J. 
Hyg. 7:706. 

— — — 1929. Ovulation requirements of Culex pipiens Linn. Biol. Bull. 56:347. 

1929. The effects of selection upon susceptibility to bird malaria in Culex 

pipiens Linn. Ann. Trop. Med. and Paras. 23:427. 

■ 1 93 1. The inheritance of natural immunity to Plasmodium cathemerium in 



two species of Culex. J. Prev. Med. 5:249. 



Culicidae 389 

THE CULTURE OF MOSQUITO LARVAE FREE FROM 
LIVING MICRO-ORGANISMS 

William Tracer. Rockefeller Institute for Medical Research 

A METHOD has recently been described (Trager, 1935) whereby 
bacteria- free larvae and adults of the yellow fever mosquito, Aedes 
aegypti, may be readily obtained. The essentials of this method will be 
briefly summarized here. 

STERILIZATION OF THE EGGS 

This is accomplished by a slight modification of the method of Mac- 
Gregor (MacGregor, 1929). "Boats," made out of coverslips by heating 
their edges in a flame to make them curl down and sterilized by dry heat, 
are placed in small sterile petri dishes containing a 5% solution of Cas- 
tile soap. From 5 to 20 eggs (as desired) are put in each boat and left in 
the soap solution 5 to 7 minutes. With sterile forceps each boat is then 
lifted out (the eggs coming with it), drained of excess liquid and placed 
in a sterile petri dish holding 80% alcohol. The eggs are left 15-17 
minutes in the alcohol and are then transferred, as before, to a sterile 
petri dish holding sterile water. Finally, the boat with its contained eggs 
is lifted with sterile forceps and dropped into a tube of culture medium. 
This method is nearly 100% successful as long as the eggs used have 
been laid recently (within 1 to 3 days) on filter paper partly immersed in 
distilled water. 

THE CULTURE MEDIUM 

Liver extract. To every 100 cc. of distilled water, 0.5 gm. of Eli Lilly 
and Company liver extract #343 is added. The somewhat turbid mix- 
ture, when filtered through paper, gives a clear amber-colored filtrate. 
The pH of this solution is adjusted to 7.0 by the addition of N/i NaOH 
(about 0.25 to 0.3 cc. per 100 cc. of solution). The medium is then 
sterilized either by passage through a Berkefeld "N" filter or by auto- 
claving. 

Yeast. A strain of Fleischmann's bakers' yeast may be used. Two- to 
4-day growths from the surface of Blake bottles of dextrose agar are 
suspended in sterile tap water, centrifuged down, re-suspended in sterile 
distilled water, again centrifuged, and finally suspended in enough sterile 
distilled water so that 10 to 11 cc. of suspension contain the yeast from 
one Blake bottle. The yeast suspension is then killed by heating at 
8o'°-8s° C. for 30 minutes. Ordinarily, 1 cc. of such a yeast extract is 
added to each large test tube (22 x 180 mm.) containing 12 to 14 cc. 
of the liver extract. 

An even simpler but equally effective medium may be made by using 



390 Phylum Arthropoda 

a i% suspension of Harris brewers' yeast (dry powder) in the liver 
extract, the mixture being tubed and autoclaved. If not more than 5 to 
10 larvae are present in a tube containing 14 cc. of the liver extract-killed 
yeast medium, the larval stage of Aedes acgypti, at a temperature of 
2 7°-2 8° C, consumes about 8 days. 

Bibliography 

MacGregor, M. E. 1929. The significance of the pH in the development of 

mosquito larvae. Paras. 21:132. 
Trager, W. 1935. The culture of mosquito larvae from living micro-organisms. 

Amer. J. Hyg. 22:18. 

References 
For the rearing of mosquito larvae see also pp. 376, 383, and 386. 



Family psychodidae* 

PSYCHODA ALTERNATA AND P. MINUTA** 

A STUDY of the breeding habits and life history of Psychoda alter- 
nata was made with the view of determining whether it might not 
be used for studies in genetics. The effort was attended with unusual 
success both in breeding the flies and in the discovery of at least two 
mutations. 

The adults are about 2 mm. in length. They ordinarily breed in decay- 
ing vegetation, but dung from either horses or cattle has proved to be 
an excellent medium. Breeding takes place readily under laboratory 
conditions, the life cycle being completed in from 12 to 16 days. Adult 
females are favorably stimulated by the culture medium, so that ovi- 
position takes place quickly. The eggs hatch in a little less than 2 days 
into active, eyed larvae resembling those of midges. The larvae feed 
for about 10 days, after which they become quiescent and pupate. 
Adults emerge 2 days later. 

Pedigreed strains were maintained in test tubes and small flasks, while 
battery jars were employed for large mass-cultures. 

M. E. D. 

* Editor's Note: The reader may find an excellent summary of what is known of the 
biology of other members of this family in an article entitled Aquatic Diptera Part I. 
Nemocera, exclusive of Chironomidae and Ceratopogonidae, Cornell University Agricul- 
tural Experiment Station Memoir 164, pp. 23-24, 1934, by O. A. Johannsen. 

** Abstracted from an article in Science 60:338, 1924, by C. L. Turner, Beloit College. 



Ceratopogonidae 391 

Family ceratopogonidae 

METHODS OF COLLECTING AND REARING 
CERATOPOGONIDAE 

Lillian Thomsex, Bethany College, Lindsborg, Kansas 

1ARVAE were collected from various localities and from such materials 
^ as blanket algae in ponds; algae in springs, streams, ponds, and 
lakes; mud taken from the shores of springs, ponds, and lakes; water 
in tree-holes; and the ooze of ulcers on maple and elm trees. 

The material was examined in different ways after being brought in. 
The larvae in algae could be removed by placing the algae on a screen 
over a silk bolting cloth bag and playing a strong stream of water on 
them. This method removed about 75% of the larvae of the swimming 
type, but such larvae as the Atrichopogon, Stilobezzia, and Dasyhelea 
had to be removed singly from the algae, the material being examined 
filament by filament under the binocular microscope. The larvae found 
in mud were obtained in like manner by sifting the mud. The pebbles 
were retained by the screen while the mud was washed into a shallow 
enameled pan and the larvae were then readily seen swimming at the 
edge or over the bottom of the pan. The larvae