LABORATORY MANUAL
OF GENERAL MICROBIOLOGY
V
LABORATOEY MANUAL f
OF
GENERAL MICROBIOLOGY
WITH SPECIAL REFERENCE TO THE
MICROORGANISMS OF THE SOIL
BY
EDWIN BROUN FRED, Ph.D.
Professor of Agricultural Bacteriology, University of Wisconsin
AND
SELMAN A. WAKSMAN, Ph.D.
Professor of Soil Microbiology, Rutgers University
C
FiKST Edition
Second Impression
McGRAW-HILL BOOK COMPANY, Inc.
NEW YORK: 370 SEVENTH AVENUE
LONDON: 6 & 8 BOUVERIE ST., E. C. 4
1928
Copyright, 1928, by the
McGraw-Hill Book Company, Inc.
PRINTED IN THE UNITED STATES OF AMERICA
THE MAPLE PRESS COMPANY, YORK, PA.
PREFACE
This laboratory manual has been designed for students in
General Microbiology, and especially for those working with soils
or with organisms isolated from the soil. Although various
exercises are described primarily for students in soils, the methods
of isolation and cultivation of bacteria, fungi, actinomyces,
algae and protozoa, and the determination of the biochemical
activities of these organisms can be used by the student in
General and Agricultural Microbiology. Special attention has
been paid to the physiology of microorganisms, including the
various so-called fermentation processes, in an attempt to bring
out not only the qualitative but also the quantitative relation-
ships of the various organisms.
It is assumed that the student has had previous training in
general botany, zoology, bacteriology and chemistry; a knowledge
of organic and physical chemistry, of mycology, and of proto-
zoology will prove of great assistance in carrying out the experi-
ments and in understanding the results.
Out of the numerous media suggested for the cultivation of
non-pathogenic organisms, the authors selected the simplest,
preferably the synthetic inorganic media, and those that have
been found to be most useful. It is impossible to give all the
chemical methods of analysis employed in microbiological
investigations. Only a few of the most essential methods,
dealing largely with the transformations of carbon and nitrogen,
and to a less extent of phosphorus and sulphur, have been
described.
In addition to references given in the text, frequent use
has been made of the various nianuals in bacteriology, a list
of which is appended.
The authors are indebted to their associates for many helpful
suggestions, especially to Dr. I. L. Baldwin, Miss Ehzabeth
McCoy, Dr. J. Blom, and Dr. R. L. Starkey.
Madison, Wis. ^dwin Broun Fred.
New Brunswick, N. J. Selman A. Waksman.
November, 1928.
CONTENTS
Page
Preface v
PART I
Culture Media
Principles of Microbial Nutrition and Composition of Culture Media
for Microorganisms 1
General Directions for the Preparation of Culture Media 3
General Media for Isolation and Cultivation of Bacteria 8
Media for Anaerobic Spore-forming Bacteria 11
Media for Fungi 12
Media for Yeasts 16
Media for Actinomyces 18
Media for Protozoa 19
Media for Algae 20
Media for Urea Bacteria 21
Media for Nitrifying Bacteria 22
Media for Nitrate-reducing and Denitrifying Bacteria 25
Media for Reduction of Sulphates and Other Sulphur Compounds . . 26
Media for Sulphur-oxidizing Bacteria 28
Media for Hydrogen and Methane Bacteria 30
Media for Iron and Manganese Oxidizing Bacteria • . . . . 31
Media for Nitrogen- fixing Bacteria 32
Media for Cellulose-decomposing Bacteria 35
Special Media 39
Preserving Stock Cultures 45
Favorable Conditions for the Development of Anaerobic Bacteria. . . 46
PART II
Methods of Staining
Methods of Staining Bacteria 47
PART III
Qualitative and Quantitative Methods of Analysis
Preparation of Reagents 52
Preparation of Standard Solutions 54
Qualitative Methods for Determining Various Forms of Inorganic
Nitrogen 56
Determination of Moisture in Soil 61
vn
40291
viii CONTENTS
Page
Moisture-holding Capacity of Soil 61
Quantitative Methods for Ammonia Determination 62
Quantitative (Colorimetric) Methods for Determining Nitrates ... 63
Quantitative Methods for Determining Total Nitrogen 65
Determination of Amino Nitrogen 69
Quantitative Determination of Carbohydrates 69
Complete Analysis of Natural Organic Material 76
Humus Determination 78
Carbon Dioxide Evolution 79
Determination of Total Carbon 80
Seed Sterilization 82
PART IV
The Study of Microorganisms in TjHE Soil
A Suggested List of Arrangement of Class Exercises 87
Apparatus for One Student 89
Laboratory Rules 89
A Black Finish for Table Tops . 91
General Characteristics of the Soil Population 92
Microscopic Examinations of Microorganisms 95
Methods for Counting Numbers of Microorganisms 97
Nitrogen-fixing Bacteria (and Nitrogen Fixation in Soil) 106
Denitrifying Bacteria 121
Nitrification 123
Urea and Protein Decomposition 126
Sulphate-reducing and Sulphur-oxidizing Bacteria 128
Iron Bacteria 132
Cellulose-decomposing Bacteria 133
Evolution of Carbon Dioxide from Soil 136
Literature 137
List of Laboratories where Cultures Can Be Secured 139
Index 141
LABOEATORY MANUAL
OF
MICROBIOLOGY
PART I
CULTURE MEDIA
PRINCIPLES OF MICROBIAL NUTRITION AND COMPOSITION OF
CULTURE MEDIA FOR MICROORGANISMS
Since the work of Pasteur in the 'fifties and 'sixties of last
century on pathogenic and non-pathogenic microbes and since
1881, when the first soUd culture medium was suggested by R.
Koch for the isolation and cultivation of bacteria, numerous
solid, liquid, and semi-solid media have been recommended.
The composition of these media depends entirely upon the food
requirements of the specific organism that is to be isolated or
cultivated. Since microorganisms vary in reference to the
nature of the nutrients which they require for their growth and
reproduction, the composition of the media which are to be used
for the isolation and cultivation of different organisms will
therefore vary. Some media are adapted to the growth of a
maximum number of different organisms, not favoring any
particular kind in preference to others. These are the media
which are used for counting the numbers of microorganisms in the
soil. Other media are highly selective in nature, allowing the
development of only one very limited group of organisms.
These selective or enrichment media are used largely for the
isolation of certain specific organisms, utilizing their specific
physiology.
In general, a medium must contain a source of energy, a source
of carbon, a source of nitrogen, and various mineral elements
1
2 LABORATORY MANUAL OF MICROBIOLOGY
(P, K, S, Ca, Mg, Fe, etc.), which are required by the organisms
for the synthesis of their cell substance. Some organisms are
very specific in the requirements of these nutrients, while others
can derive their energy, their carbon, their nitrogen, and the min-
eral elements from a great variety of substances.
Microorganisms are divided broadly into two large groups, on
the basis of their energy requirements: (1) The autotrophic
organisms, which can obtain the energy required for their
activities from the oxidation of inorganic elements or their
compounds or from simple compounds of carbon, their carbon
from carbon dioxide, and their nitrogen and other minerals
from inorganic compounds. In addition to those organisms
which can obtain their energy from the oxidation of inorganic
substances or simple compounds of carbon (chemosynthetic),
the chlorophyll-bearing plants, which obtain their energy photo-
synthetically are also classified with this group. (2) The
heterotrophic organisms which obtain their energy and carbon
from complex organic substances.
Among the heterotrophic organisms, however, there is also
very considerable specificity. Some organisms can obtain their
energy and carbon only from celluloses, while the great majority
of microorganisms cannot attack celluloses and cannot utilize
them either as sources of energy or of carbon. Some organisms
can obtain their nitrogen from inorganic compounds, such as
ammonium salts or nitrates; others require only organic nitrogen
sources; still others can use even gaseous atmospheric nitrogen.
Some can grow at a wide range of reactions; for the activities of
others only a very limited range of hydrogen-ion concentration
exists. Some organisms are able to grow on a great variety of
media, liquid or solid; others will develop only on very specific
media, when the particular nutrients are available.
In addition to the nature of the nutrients, it is also important
to keep in mind that a specific osmotic pressure is required; in
other words, the concentration of the nutrients must not vary
within too wide limits. Solid media are often required; for this
purpose either inorganic gels, largely silicic acid (also mag-
nesium-gypsum blocks), or organic gels of a carbohydrate
nature (agar-agar, etc.) or of a protein nature (gelatin, coagulated
egg albumen, coagulated blood serum) are employed.
CULTURE MEDIA 3
These considerations can help one to reaUze why such a great
variety of culture media have been proposed at various times and
are used in microbiological studies. These media are frequently
modified as regards the concentration of certain of the nutrients,
reaction, buffer content, elimination of one nutrient and sub-
stitution of another.
GENERAL DIRECTIONS FOR THE PREPARATION OF CULTURE
MEDIA
Of the numerous formulae of various culture media, which have
frequently been only briefly described, only the most essential
and those which have been tried repeatedly and found useful
are given in this manual. It is assumed that the reader has at
hand a manual of general bacteriology, such as the Manual of
Methods Prepared by the Society of American Bacteriologists;
hence, all directions for the preparation of standards, for measur-
ing the hydrogen-ion concentration, buffer content, etc. are
omitted. The formulae are arranged according to the general
physiological characters of the microorganisms.
For general purposes, the reaction of culture media for bacteria
should be about the neutral point, or pH 7.0. Some will grow
at a considerably lower pH value, others will grow only at the
neutral point or even at a more alkaline reaction. In some
cases, bacteria can be separated from one another, by merely
adjusting the reaction to such a point as to eliminate one group
of organisms without injuring the other. Fungi are able, as a
rule, to grow at a much higher acidity than bacteria. This fact
is utilized frequently for the separation of these two groups of
organisms: by adjusting the reaction of the medium to pH 4.0,
the bacteria are practically eliminated, while the great majority
of fungi are not affected.
When it is necessary to adjust the reaction of a medium, the
scheme outlined here will be found convenient. Place 2 cubic
centimeters of the medium and 8 cubic centimeters of water in
a test tube and add 4 or 5 drops of phenol red or any other of the
desired indicators. Now add 0.1 iV or 0.05 N sodium hydroxide
from a burette until the color of the solution matches that of a
known standard. Calculate and add to the medium the amount
4 LABORATORY MANUAL OF MICROBIOLOGY
of normal sodium hydroxide required to make the reaction
pH 7.0. Check the results by repeating titrations. All attempts
to adjust the culture to an exact pH are unnecessary since the
changes in reaction due to heat, glassware, etc., will be found
greater than a pH 0.1.
Filtration. Funnel Filter. — A cotton filter is prepared as
follows: In the base of a large funnel place a small amount of
clean excelsior. In place of the excelsior a small spiral of copper
wire or iron-wire netting may be used. On top of this put two
or three layers of absorbent cotton. Split a piece of absorbent
cotton, somewhat larger than the top of the funnel, horizontally
into two layers of equal thickness. Place one layer of cotton
above the other, so that the fibers are at right angles and wash the
cotton filter with boiling water. Pour the medium, slowly at
first, on to the filter. (In order to avoid breaking the filter use
a glass rod to direct the fluid to the center of the filter). When
the filtrate begins to come through the cotton, fill the funnel.
If the first filtrate is not clear, the turbid liquid should be refil-
tered through the same cotton.
If several liters of media are prepared the suction method given
below will be found convenient.
Suction Filter. — Prepare an absorbent cotton filter on the top
of a heavy walled glass bottle; milk bottles are satisfactory.
Place a layer of cheesecloth on the top of the bottle, then one or
two layers of absorbent cotton and cover with another layer of
cheesecloth. Be sure that the filter is tied tightly around the
neck of the bottle. Now invert bottle in a saucepan containing
the medium. It is sometimes necessary to use a weight to hold
down the empty bottle. Place in the autoclave and heat to 10
or 15 pounds pressure for 20 minutes. Allow to cool slowly.
The heat will exhaust the bottles and as it cools the medium is
drawn up through the filter pads. Since the vacuum in the
bottles is never complete, the bottles are usually found to be
about two-thirds full after filtration.
Solid Media. — For the preparation of solid media, agar, gelatin,
or silica gel are commonly employed. Some of the differences
between these substances are shown in the following table:
CULTURE MEDIA
General Properties
Agar
Gelatin
Silica gel
Source
Chemical nature
Reaction
Melting point — usual concentra-
tion
Solidifies, usual concentration . . . .
Tryptic digestion
Water of condensation
Usual concentration
Plant
Carbohydrate
Faintly acid
96°C.
40°C.
Not affected
Present
1 to IH per
cent
Animal
Protein
Acid
25°C.
25°C.
Liquefied
None
10 to 12
per cent
Inorganic
Silicic acid
Acid
None
5 to 6 per
cent
Agar-agar is prepared by extracting certain seaweeds (largely
Gelidium corneum) with hot water. It consists almost entirely of
polysaccharides (largely galactans with some pentosan^) and a
small admixture of protein and mineral matter.
Approximate Chemical Composition of Agar and Gelatin
Agar, 2 Gelatin,
Per Cent Per Cent
Moisture 16.0 14 to 15
Ash, dry basis 4.4 0.6
Calcium oxide 1.15 0.0
Magnesium oxide 0 . 77 • 0.0
Nitrogen 0.40 18.3
Pure agar has practically no buffer effect within the range com-
monly employed.
Washed Agar. — This is prepared as follows: Take 1,000 grams
of agar shreds, place in an enamel pail or glass vessel with 10 liters
of distilled water and allow to stand for 24 hours at room tem-
perature. Place a piece of cheesecloth over the top of the vessel,
pour off the water and once more add fresh distilled water and
allow to soak another 24 hours. Now pour off the water, and
allow the agar to air dry in thin layers.
Agar treated in this way is much lower in calcium and mag-
nesium, as well as in soluble organic matter. In the preparation
1 KoNiG and Bettels, Ztschr. Unters. Nahr. u. Genussm., 10: 487, 1905.
2 Fellers, C. R., /. Ind. Eng. Chem., 8: 1128, 1916.
6 LABORATORY MANUAL OF MICROBIOLOGY
of certain kinds of culture media, it has been found that the
removal of these inorganic salts prevents the formation of a
precipitate during sterilization and hence gives a clearer medium.
Preparation of Silica Gel — Prepare a 25 per cent solution of
equal parts of sodium and potassium silicate (C.P.). Adjust
the specific gravity of the solution to 1.06. Five cubic centi-
meters of this solution is then poured into a Petri dish, and a
drop of phenolphthalein added. An approximately 5 per cent
solution of HCl is added to this silicate, from a graduated pipette,
until the color is just discharged. The acid is then adjusted so
that 5 cubic centimeters of HCl solution will just neutralize 5
cubic centimeters of the sihcate. If care is taken not to overrun
the acid, the gel will set in a few minutes.
To make the Petri plates, add a definite volume of the silicate,
solution to an equivalent amount of acid, placed in a large flask.
Shake the mixture vigorously and immediately pour 20- to 50-
cubic centimeter portions into Petri dishes. Allow the plates
to harden upon a flat surface. They are then placed in deep flat
vessels and dialyzed in running tap water until free from chlorides.
About 24 hours is required for this purpose. The dishes are then
removed and transferred to a sterile vessel containing boiled
distilled water. This is replaced several times. After they have
been properly washed, the dishes are drained and treated with
the nutrient medium.
Sterilization. — Unless otherwise stated, it is recommended
that all media be sterilized in the autoclave. In the case of
liquid and agar media, about 120°C. for 20 minutes will be found
sufficient for complete destruction of all microorganisms. In
sterilization, it must be remembered that the time required to
kill bacteria depends upon the degree of heat at the center of the
vessel and the nature of the medium. This degree of heat is
determined by the size of the container, the original temperature
and viscosity of the contents, and also by the free access of the
steam to the surface of the container. All sterilizers should be
equipped with temperature controls and with air outlets at the
bottom. To secure the best results, place the medium .n small
containers and space in the autoclave in such a way as to give
free access of the steam to the surfaces. The time of steriliza-
tion must be determined for the various types of media. Agar
CULTURE MEDIA
media are best sterilized at 15 pounds pressure for 15 to 30
minutes. Gelatin media are sterilized in flowing steam on three
consecutive days or in the autoclave at 10 pounds pressure for
30 minutes. Soil is extremely difficult to sterilize. Small
containers with about 5 to 10 grams of soil are heated in the
autoclave at 15 to 20 pounds pressure for 2 hours one day, or 1
hour on each of 2 consecutive days, or in flowing steam on at
least 7 consecutive days.
The relation between pressure and temperature is shown in
the table below:
Pressure
Temperature
Pounds per
square inch
Atmosphere
100°C
0
7.5
15.0
22.5
30.0
0
112°C
0.5
121°C
1.0
128°C
1.5
135°C
2.0
The reaction of a medium has a decided influence upon the
decompositions brought about by sterilization. The lower the
pH of the medium, the greater will be the hydrolyzing effect of
the sterilization, not only of gelatin, but also of agar, so that at
pH 4.0, as much as 3 per cent agar has to be used to obtain a
soUd medium. When the medium is alkaline, the iron of the
solution will be precipitated out. This may necessitate the
addition of a small amount of sterile iron salt, or the sterilization
of the medium by filtration through a Berkefeld filter.
It is recommended that culture media be inoculated as
soon as possible after sterilization. Because of the adsorbed
air, old culture medium is unfit for the growth of strict anaerobic
bacteria.
Silica gel media need not be sterilized. The washing with
sterik \ vter, followed by the use of highly selective media is
sufficient to assure sterility, as far as air contaminations are
concerned. Their use is adapted only to the very selective
organisms.
8 LABORATORY MANUAL OF MICROBIOLOGY
GENERAL MEDIA FOR ISOLATION AND CULTIVATION OF
BACTERIA
Medium 1
Nutrient Broth or Bouillon
Beef extract 3.0 gm.
Peptone 5.0 gm.
Distilled water 1,000.0 cc.
Heat until extract and peptone are dissolved.
Adjust reaction to pH 6.6 to 7.0 using Brom thymol blue.
Medium 2
I
Nutrient or Beef-extract Agar
The same composition as nutrient broth (Medium 1) plus 15
grams of agar.
Medium 3
Nutrient or Beef-extract Gelatin
Gelatin 100.0 to 150 . 0 gm.
Beef extract 3.0 gm.
Peptone 5.0 gm.
Distilled water 1,000.0 cc.
In a convenient vessel measure 1,000 cubic centimeters of
nutrient broth.
Add 10 to 15 per cent gelatin. Let the gelatin soak 5 to 10
minutes.
Heat over water bath until dissolved.
Adjust the reaction as directed in the preparation of nutrient
broth. Gelatin is decidedly acid and has a high buffer value;
it will, therefore, require more NaOH for neutralization than
bouillon or agar.
If properly prepared, gelatin may be filtered through filter
paper. Otherwise it will be necessary to use an absorbent-
cotton filter.
CULTURE MEDIA 9
Medium 4
Sodium Caseinate or Nutrose Agar
Agar 12.5 gm.
Sodium caseinate (nutrose) 2.0 gm.
Glucose 1.0 gm.
Dipotassium phosphate (K2HPO4) 0.2 gm.
Magnesium sulphate (MgS04-7H20) 0.2 gm.
Ferrous sulphate (FeS04-7H20) trace
Tap water 1,000.0 cc.
Reaction approximately pH 6.8.
It is not necessary to adjust the reaction of this medium. To
secure a clear agar filter through cotton.
Medium 5
Sodium Albuminate AgarI
Agar 12.5 gm.
Glucose 1.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 0.2 gm.
Ferric sulphate (Fe2(S04)3'9H20) trace
Egg albumen (powdered) 0 . 25 gm.
Water, distilled 1,000.0 cc.
1 Waksman, S. a., Soil Sci., 14: 283-298, 1922.
Make a suspension of the egg albumen in a little water (5
cubic centimeters), add a drop of phenolphthalein and enough
0.1 iV NaOH to dissolve and bring solution to a permanent
pink color ; the sodium albuminate is then added to the remainder
of the medium. Reaction is about pH 7.2.
Medium 6
Nahrstoff-Heyden Agar2
Agar 12.5 gm.
Nahrstoff-Heyden 7.5 gm.
Distilled water 1,000.0 cc.
2 Hesse, W., and Niedner, Z. Hijg., 29: 454-462, 1898.
To 500 cubic centimeters of cold distilled water in a flask add
7.5 grams of Nahrstoff-Heyden. Shake until a good suspension
is obtained and allow the mixture to stand for 30 minutes or more.
10 LABORATORY MANUAL OF MICROBIOLOGY
Heat in steamer or double boiler for 1 hour, or until the upper
portion of the solution is clear.
While hot filter through paper.
Dissolve 12.5 grams of agar in 500 cubic centimeters of water.
Filter and mix the Nahrstoff-Heyden and agar solutions.
It is not necessary to adjust the reaction of this medium.
Medium 7
Soil-extract Agar
Agar 12.5 gm.
Glucose 1.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Soil extract (stock) ^ 100 . 0 cc.
Tap water 900.0 cc.
1 Stock Solution of Soil Extract. — This is prepared by heating 1,000 grams
of garden soil with 1,000 cubic centimeters of tap water in the autoclave for
30 minutes. A small amount of calcium carbonate is added and the whole
is filtered through a double paper filter. The turbid filtrate should be poured
back on to the filter until it comes through clear.
Dissolve the agar in 900 cubic centimeters of water by heating
in the steamer for 1 hour or longer. Add 100 cubic centimeters
of the stock soil-extract solution.
Add the glucose just prior to tubing.
The reaction should be pH 6.8.
Medium 8
Soil- EXTRACT Gelatin 2
Gelatin 150.0 gm.
Glucose 1.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Soil extract (stock) 100.0 cc.
Tap water 900.0 cc.
2 Conn, H. J., N. Y. Agr. Expt. Sta., Tech. Bull. 38, 1914.
Dissolve the gelatin in the diluted soil-extract solution by
heating slowly in the steamer.
Clarify the medium with egg albumen.
Add 1 gram of glucose and adjust the reaction to pH 6.8.
To prevent the spread of liquefying colonies add 5 grams of
sodium chloride (NaCl) per liter.
CULTURE MEDIA 11
Medium 9
ASPARAGIN MaNNITOL AgAR^
Agar 15.0 gm.
Dipotassium phosphate (K2HPO4) 1.0 gm.
Potassium nitrate (KNO3) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 0.2 gm.
Calcium chloride (CaGl2) 0.1 gm.
Sodium chloride (NaCl) 0.1 gm.
Ferric chloride (FeCl3-6H20) trace
Asparagin (C4H8N2O3) 0.5 gm.
Mannitol 1.0 gra.
Water 1,000.0 cc.
1 Thornton, H. G., Ann. Appl Biol, 9: 241-274, 1922.
After the agar and salts have been dissolved add the mannitol,
adjust the reaction to pH 7.4 (Brom thymol blue) and filter.
Mannitol is recommended in place of glucose or related sugars
because it decomposes less than the common sugars during
sterilization.
Medium 10
Tap- WATER Gelatin 2
Gelatin 120 to 200.0 gm.
Tap water 1,000.0 cc.
2 Conn, H. J., N. Y. Agr. Expt. Sta., Tech. Bull 57, 1917.
Reaction pH 6.5.
Incubate the plates at 18°C. for 7 days. This medium is
especially useful in qualitative studies of soil flora.
MEDIA FOR ANAEROBIC SPORE-FORMING BACTERIA
Medium 11
Glucose-phosphate Nitrogen Free Medium for Anaerobic Bacteria^
Dipotassium phosphate (K2HPO4) 1.0 gm.
Magnesium sulphate (MgS04-7H20) 0.2 gm.
Sodium chloride (NaCl) 0.01 gm.
Ferrous sulphate (FeS04-7H.20) 0.01 gm.
Manganese sulphate (MnS04-4H20) 0.01 gm.
Glucose 20.0 gm.
Calcium carbonate (CaCOa) 30 . 0 gm.
Distilled water 1,000.0 cc.
3 Winogradsky, S., Centr. BakL, II Abt., 9: 49, 1902.
12 LABORATORY MANUAL OF MICROBIOLOGY
In order to prevent decomposition of the glucose, it is well to
sterilize the calcium carbonate separately. Tube in deep layers.
For a solid medium add 15 grams of agar to each liter.
Medium 12
Winogradsky's Glucose-peptone Agar
Agar 15.0 gm.
Peptone 5.0 gm.
Beef extract 3.0 gm.
Glucose 5.0 gm.
Tap water 1,000.0 cc.
Reaction, pH 7.0. Deep tubes.
Medium 13
Peptone-mannitol Solution for Anaerobic Bacteria^
Peptone 2.5 gm.
Beef extract 2.0 gm.
Mannitol or glucose 10.0 gm.
Manganese sulphate (MnS04-4H20) 0.004 gm.
Tap water 1,000.0 cc.
1 Truffaut, G. et N. Bezssonoff, Compt. rend. Acad. Sci., 173: 868, 1921.
Adjust the reaction to pH 7.0.
MEDIA FOR FUNGI
Medium 14
Raulin's Solution
Ammonium nitrate (NH4NO3) 4.0 gm.
Ammonium phosphate ((NH4)2HP04) 0.6 gm.
Ammonium sulphate ((NH4)2S04) 0 . 25 gm.
Potassium carbonate (K2CO3) 0.6 gm.
Magnesium carbonate (MgCOs) 0.4 gm.
Zinc sulphate (ZnS04) 0.07 gm.
Ferrous sulphate (FeS04-7H20) 0 . 07 gm.
Potassium sihcate (K2Si03) 0 . 07 gm.
Tartaric acid 4.0 gm.
Sucrose 70 . 0 gm.
Distilled water 1,500.0 cc.
This medium is given here due to its historical significance.
The cultivation of fungi can be carried out successfully on a much
simpler medium.
CULTURE MEDIA 13
Medium 15
Ammonium Nitrate-sucrose Solution
Ammonium nitrate (NH4NO3) 3.0 gm.
Monopotassium phosphate (KH2PO4) 1.0 gm.
Magnesium sulphate (MgS04-7H20) 1.0 gm.
Ferrous sulphate (FeS04-7H20) trace
Sucrose 50 . 0 gm.
Distilled water 1,000 . 0 cc.
Medium 16
Sodium Nitrate-sucrose Solution (Czapek's)^
Sodium nitrate (NaNOs) 2.0 gm.
Monopotassium phosphate (KH2PO4) 1 .00 gm.
Potassium chloride (KCl) 0 . 50 gm.
Magnesium sulphate (MgS04-7H20) 0 . 50 gm.
Ferrous sulphate (FeS04 -71120 0.01 gm.
Sucrose 30 . 00 gm.
Distilled water 1,000.00 cc.
1 CzAPEK, F., Beitrage z. chem. Phijsiol. u. Path., I: 538-560, 1901.
To prepare an agar medium, add 15 grams of agar to 1,000 cubic
centimeters of solution. Dissolve and filter. The reaction is
left unadjusted.
Medium 17
Asparagin-glucose Agari
Agar 15.0 gm.
Glucose 10 . 0 gm.
Asparagin 0.5 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Distilled water 1,000 . 0 cc.
1 Krainsky, a., Centr. Bakt. II Abt., 41: 649-688, 1914.
This medium is also used for the cultivation of actinomyces.
Medium 18
Peptone-glucose Acid Agar^
Agar 25.0 gm.
Monopotassium phosphate (KH2PO4) 1.0 gm.
Magnesium sulphate (MgS04-7H20) 0.5 gm.
Peptone 5.0 gm.
Glucose 10.0 gm.
Water 1,000 . 0 cc.
1 Waksman, S. a., /. BacL, 7: 339-341, 1922.
Reaction pH 3.8 to 4.0.
14 LABORATORY MANUAL OF MICROBIOLOGY
Dissolve the salts, peptone and agar by steaming for H to 1
hour, adjust the reaction to pH 3.8 to 4.0 with N/1 H2SO4
(about 5 to 7 cubic centimeters of N/1 per liter) and then add
the glucose. Filter through cotton, using a suction flask.
Because of the acid reaction steriUze in the autoclave at 110°C.
(7 to 8 pounds pressure) not more than 10 minutes. Avoid
excessive heat.
The medium may be prepared without addition of acid,
filtered, then placed in 100-cubic centimeter portions, in Erlen-
meyer flasks and sterihzed. Whenever needed for use, the agar
is melted by placing flasks in boiling water. One cubic centimeter
of N/2 H2SO4 is then added to each flask giving the desired
reaction.
Medium 19
Potato-glucose Agar
Agar 30.0 gm.
Potato 200.0 gm.
Glucose 20.0 gm.
Tap water 1,000.0 cc.
Peel and shce 200 grams of potatoes. Cook in 1,000 cubic
centimeters water for 1 hour in the steamer. Strain or decant
the clear Hquid and restore it to the original volume. Add 20
grams glucose and 30 grams agar. Heat in steamer until the
agar is dissolved. Filter through cotton filter.
Medium 20
ASPARAGIN-STARCH AgAR^
(Brown, modified for color formation)
Agar 15.00 gm.
Potato starch 10.00 gm.
Asparagin 0 . 20 gm.
Potassium phosphate, ortho (K3PO4) 1-25 gm.
Magnesium sulphate (MgS04-7H20) 0 . 75 gm.
Water 1,000.00 cc.
1 Brown, W., Ann. BoL, 39: 405, 1925.
CULTURE MEDIA 15
Medium 21
Clover-sucrose Agar
Agar 25 . 0 gm.
Clover (green tops) 500 . 0 gm.
Sucrose 2.0 gm.
Potassium nitrate (KNO3) 0.5 gm.
Tap water 1,000.0 cc.
Extract the clover tissue by heating for 1 hour in the steamer;
filter and add the other ingredients. Add 1 to 2 drops of concen-
trated lactic acid to each tube of the medium just before pouring
plates.
Reaction approximately pH 3.5.
Medium 22
Peptone-malt Agar^
Agar 20 . 0 gm.
Peptone 1.0 gm.
Glucose 20.0 gm.
Malt extract (desiccated) 20 . 0 gm.
Tap water 1,000.0 cc.
1 PovAH, A. H. W., Torrey Bot. Club. Bull, 44: 241, 287, 1917.
This medium is used for the cultivation of Mucorales.
Medium 23
Ammonium Nitrate Lactic Acid Solution^
Ammonium nitrate (NH4NO3) 10.0 gm.
Dipotassium- phosphate (K2HPO4) 5.0 gm.
Magnesium sulphate (MgS04-7H20) 1.0 gm.
Lactic acid (85 per cent) 2.0 gm.
Tap water 1,000.0 cc.
2v. TuBEUF, C, Centr. Bakt., II Abt., 9: 127-135, 1902.
Fifty-cubic centimeter portions of this solution are added
to flasks containing 10 grams of filter paper. This medium is
used for the cultivation of wood-destroying fungi.
16 LABORATORY MANUAL OF MICROBIOLOGY
MEDIA FOR YEASTS
Medium 24
Sucrose Peptone (Hansen's) Solution
Sucrose or maltose 50 . 0 gm.
Peptone 10.0 gm.
Monopotassium phosphate (KH2PO4) 3.0 gm.
Magnesium sulphate (MgS04-7H20) 2 . 0 to 5 . 0 gm.
Distilled water 1,000. 0 cc.
Medium 25
Sucrose — Malt Extract Agar
Agar 15.0 gm.
Sucrose 50 . 0 gm.
Malt extract (desiccated) 10. 0 gm.
Distilled water 1,000.0 cc.
Medium 26
Malt Extract Broth
Malt (finely ground) 250 . 0 gm.
or
Malt extract (desiccated) 15.0 gm.
Water 1,000.0 cc.
Incubate for 1 hour at 65°C. and test for starch; when the
latter is still present, incubate for a longer time. Filter through
a hand press, cook for 2 to 3 hours. Again filter and make up
to 1 liter. For a soUd medium add 1.2 to 1.5 per cent of agar.
Medium 27
Fresh Yeast Infusion
Yeast (fresh-pressed cakes, starch-free) 100.0 gm.
Water 1,000.0 cc.
The 1-pound cake of starch-free pressed yeast will be found
useful. Steam for 3 to 4 hours with occasional stirring. Steri-
lize in deep layers and allow to stand for 1 week. If undisturbed
the yeast cells will settle to the bottom and leave a clear straw-
colored liquid above. This clear infusion should be siphoned
off and the reaction adjusted to pH 6.6 to 6.8. Yeast extract
prepared in this way contains only a trace of fermentable carbo-
CULTURE MEDIA 17
hydrates, a small amount of non-volatile acid, and about 50
milligrams of nitrogen in 100 cubic centimeters. For fermenta-
tion studies, add 1 to 2 per cent of the required carbohydrate.
For a sohd medium add 15 grams of agar.
Medium 28
Dried Yeast Infusion
Yeast (dry, starch-free) 10.0 gm.
Water 1,000.0 cc.
Proceed as described under medium 27.
Medium 29
Dried Yeast-peptone Infusion
Dried yeast 10. 0 gm.
Peptone 10.0 gm.
Water 1,000.0 cc.
Medium 30
Ammonium Sulphate-glucose Solution
Ammonium sulphate ((NH4)2S04) 5 .00 gm.
Dipotassium phosphate (K2HPO4) 0 .75 gm.
Magnesium sulphate (MgS04-7H20) 0 . 10 gm.
Tartaric acid (C4H6O6) 1 .00 gm.
Glucose 100.00 gm.
Distilled water 1,000.00 cc.
Medium 31
Raisin Extract
Raisins 375.0 gm.
Ammonium chloride (NH4CI) 2.0 gm.
Distilled water 1,000.0 cc.
Allow the raisins to stand in 1 Uter of water at ice-box tem-
perature for 1 to 2 days. Mash, add the ammonium chloride,
cook in the steamer for 30 minutes, and filter.
18 LABORATORY MANUAL OF MICROBIOLOGY
Medium 32
Synthetic Solution for Yeast^
Sucrose 50.0 gm.
Ammonium sulphate ((NH4)2S04) 6.0 gm.
Mono-ammonium phosphate (NH4H2PO4) 2.0 gm.
Potassium sulphate (K2SO4) 4.0 gm.
Magnesium sulphate (MgS04-7H20) 2 . 25 gm.
Calcium sulphate (CaS04) 2 . 25 gm.
Tap water 1,000.0 cc.
1 Hayduck, F., U. S. Patent, Reissue 15, 716.
Medium 33
Carrot Extract Agar for Spore Formation of Yeasts
Carrots 1,000.0 gm.
Calcium sulphate (CaS04-2H20) 10.0 gm.
Agar 20.0 gm.
Water 200 to 300 cc.
Wash and grind the carrots in a meat chopper. Add 200 to
300 cubic centimeters of tap water and boil for about 10 minutes.
Filter through cheesecloth. Add the calcium sulphate and agar
and heat in steamer for 30 minutes. Tube for slopes and sterilize
at 15 pounds for 15 to 20 minutes.
MEDIA FOR ACTINOMYCES^
Medium 34
Nitrate-sucrose Agar^
Agar 15.0 gm.
Sodium nitrate (NaNOs) 2.0 gm.
Dipotassium phosphate (K2HPO4) 1.0 gm.
Magnesium sulphate (MgS04-7H20) 0.5 gm.
Potassium chloride (KCl) 0.5 gm.
Ferrous sulphate (FeS04-7H20) 0.01 gm.
Sucrose 30.0 gm.
Water 1,000.0 cc.
Reaction approximately pH 7.0.
2 For the cultivation of acid-resisting actinomyces, the reaction of these
media as well as of medium 17 should be adjusted to pH 4.0. Jensen, H.
L., Soil Sci. 25 : 225, 1928.
3 Waksman, S. a., Soil Sci., 8: 71-215, 1919.
CULTURE MEDIA 19
Medium 35
Sodium Asparaginate-glycerol Agar^
Agar 15.0 gm.
Glycerol 10.0 cc.
Dipotassium phosphate (K2HPO4) 1.0 gm.
Sodium asparaginate 1.0 gm.
Water 1,000.0 cc.
1 Conn, H. J., N. Y. Agr. Expt. Sta., Tech. Bull 60, 1917.
Reaction approximately pH 7.0.
Medium 36
Ammonium Sulphate-starch Agar^
Agar 10.0 gm.
Starch (soluble) 10 . 0 gm.
Ammonium sulphate ((NH4)2S04) 2.0 gm.
Dipotassium phosphate (K2HPO4) 1.0 gm.
Magnesium sulphate (MgS04-7H20) 1.0 gm.
Sodium chloride 1.0 gm.
Calcium carbonate (CaCOs) 3.0 gm.
Tap water 1,000.0 cc.
2 McBeth, I. G., and F. M. Scales, U. S. Dept. Agr., Bur. PI. Ind. Bull.
266, 1913.
Mix the starch with a little cold water and stir well before
adding the salts.
MEDIA FOR PROTOZOA
Medium 37
Nutrient or Beef Extract Agar^
Agar 15.0 gm.
Beef extract 3.0 gm.
Peptone 10.0 gm.
Sodium chloride (NaCl) 5.0 gm.
Distilled water 1,000.0 cc.
3 Sandon, H., and D. W. Cutler, /. Linn. Soc. Zool, 38 : 1, 1924.
Adjust the reaction to pH 7.0.
Medium 38
Hay Infusion
Meadow hay (finely chopped) 50 . 0 gm.
Tap water 1,000.0 cc.
Boil for 2 hours and allow to stand over night. Filter.
20 LABORATORY MANUAL OF MICROBIOLOGY
Medium 39
Ammonium Lactate Solution
Ammonium lactate 0.1 gm.
Ammonium chloride (NH^Cl) 0.3 gm.
Potassium chloride (KCl) 0.3 gm.
Magnesium sulphate (MgS04-7H20) 0.001 gm.
Disodium phosphate (Na2HP04) 0.01 gm.
Calcium chloride (CaCU) 0.2 gm.
Glucose 0.4 gm.
Distilled water 1,000 . 0 cc.
Medium 40
Mannitol Soil Extract
Soil extract (stock) 100 . 0 cc.
Mannitol 10.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Tap water 900 . 0 cc.
Medium 41
Mannitol-phosphate Solution^
Mannitol (C6H8(OH)6) 10.0 gm.
Magnesium sulphate (MgS04-7H20) 0.2 gm.
Monopotassium phosphate (KH2PO4) 0.2 gm.
Sodium chloride (NaCl) 0 . 2 gm.
Calcium sulphate (CaS04-2H20) 0.1 gm.
Calcium carbonate (CaCOs) 5.0 gm.
Distilled water 1,000.0 cc.
1 AsHBY, S. F., J. Agr. Sci., 2: 38, 1907.
Dissolve the phosphate separately in a little water and make
the solution neutral to phenolphthalein with A^/1 NaOH; then
add to the other ingredients. For a solid medium add 15 grams
of agar to each liter.
MEDIA FOR ALG^
Medium 42
Calcium Nitrate Solution (Detmer's)
Calcium nitrate (Ca(N03)2) 10 gm.
Potassium chloride (KCl) 0. 25 gm.
Magnesium sulphate (MgS04-7H20) 0 . 25 gm.
Monopotassium phosphate (KH2PO4) 0 . 25 gm.
Tap water 1,000.0 cc.
CULTURE MEDIA 21
Dilute 1 part of the above medium with 2 parts of tap water
and add 0.01 per cent of ferric chloride (FeCl3'6H20).
Medium 43
Sodium Nitrate Solution (Bristol's) ^
Monopotassium phosphate (KH2PO4) 0.5 gm.
Sodium nitrate (NaNOs) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 0 . 15 gm.
Calcium chloride (CaCl.) 0.05 gm.
Sodium chloride (NaCl) 0.05 gm.
Ferric chloride (FeCh-OH.O) 0 . 01 gm.
Water 1,000.0 cc.
1 Bristol, B. M., Ann. Bot., 34: 35-79, 1920.
Because of the special sensitiveness of the algae to copper,
it is well to avoid distilled water unless from glass.
MEDIA FOR UREA BACTERIA
Medium 44
Urea Citrate Solution^
Urea (CO(NH2)2) 30.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Calcium citrate (Ca3(C6H507)2-4H20) . . . 10.0 gm.
Tap water 1,000 :0 gm.
2S0HNGEN, N. G., Centr. Bakt. II Abt., 23: 91-98, 1909.
Reaction about pH 7.2.
Medium 45
Urea Solution ^
Urea .^ 20.0 gm.
Dipotassium phosphate (K2HPO4) 1-0 gm.
Calcium chloride (CaClo) 0.1 gm.
Magnesium sulphate (MgS04-7H20) 0.3 gm.
Sodium chloride (NaCl) 0.1 gm.
Ferric chloride (FeCla-OHaO) 0.01 gm.
Beef extract 5.0 gm.
Tap water 1,000 . 0 cc.
3 Viehoever, A., Centr. Bakt, II Abt., 39: 209-359, 1913.
22 LABORATORY MANUAL OF MICROBIOLOGY
Medium 46
Urea Soil Extract^
Urea (CO(NH2)2) 50.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Soil extract (stock) 100 . 0 cc.
Tap water 900 . 0 cc.
iLoHNis, F. Centr. BakL, II Abt., 14: 714-723, 1905.
Medium 47
Urea Bouillon Gelatin or Agar
Urea (CO(NH2)2) 20.0 gm.
Gelatin 120 to 150.0 gm.
or
Agar 15.0 gm.
Bouillon 1,000.0 cc.
Reaction about pH 7.5.
MEDIA FOR NITRIFYING BACTERIA
Medium 48
Ammonium Sulphate Solution^
Ammonium sulphate ((NH4)2S04) 1.0 gm.
Dipotassium phosphate (K2HPO4) 1.0 gm.
Sodium chloride (NaCl) 2.0 gm.
Magnesium sulphate (MgS04-7H20) 0.5 gm.
Ferrous sulphate (FeS04-7HoO) trace
Magnesium carbonate (MgCOs) excess
Distilled water 1,000 . 0 cc.
2 WiNOGRADKSY, S. Lafar's "Handb. techn. mylol.,'' 3: 132-181, 1904.
In order to prevent any loss of ammonia, it is desirable to
sterilize the magnesium carbonate separately. When cool add
an excess of the magnesium carbonate to each flask.
Medium 49
Magnesium Ammonium Phosphate Solution
Magnesium ammonium phosphate (MgNH4P04-
6H2O) 2.0 gm.
Dipotassium phosphate (K2HPO4) 1.0 gm.
Sodium chloride (NaCl) 2.0 gm.
Magnesium sulphate (MgS04-7H20) 0.5 gm.
Ferrous sulphate (FeS04 -71120) trace
Magnesium carbonate (MgCOs) 5.0 gm.
Tap water 1,000.0 cc.
CULTURE MEDIA 23
Medium 50
Sodium Ammonium Phosphate Solution
Sodium ammonium phosphate (Na(NH4)HP04-
4H2O) 3.4 gm.
Potassium chloride (KCl) 2.0 gm.
Magnesium sulphate (MgS04-7H20) 0.5 gm.
Ferrous sulphate (FeS04-7H20) trace
Magnesium carbonate (MgCOs) excess
Distilled water 1,000.0 cc.
To prevent a loss of ammonia do not add the MgCOs until
after sterilization
Medium 51
Sodium Nitrite Solution^
Sodium nitrite (NaN02) 1.0 gm.
Sodium carbonate (Na2C03) 1.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Sodium chloride (NaCl) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 0.3 gm.
Ferrous sulphate (FeS04-7H20) trace
Distilled water 1,000 . 0 cc.
1 Fred, E. B., and A. Davenport, Soil Sci., 11: 389-404, 1921.
Medium 52
Medium for Oxidation of Ammonia and Nitrite to Nitrate
Ammonium sulphate ((NH4)2S04) 2.0 gm.
Dipotassium phosphate (K2HPO4) 1.0 gm.
Magnesium sulphate (MgS04-7H20) 0.5 gm.
Sodium chloride (NaCl) 2.0 gm.
Ferrous sulphate (FeS04-7H20) trace
Calcium carbonate 5.0 gm.
Tap water 1,000.0 cc.
Add the calcium carbonate after sterilization.
Medium 53
Washed Agar for Nitrifying Bacteria
Agar (washed)2 25.0 gm.
Water 1,000.0 cc.
2 Allow the agar to wash in running water for 5 to 7 days, and dry at 60°C.
24 LABORATORY MANUAL OF MICROBIOLOGY
Tube the agar in 10-cubic centimeter portions and sterilize.
To this melted agar, at 45°C. in tubes or Petri dishes, add 1-
cubic centimeter portion of the following salts.
1. Dipotassium phosphate (K2HPO4) 1.5 gm.
Water 100.0 cc.
2. Ammonium sulphate ((NH4)2S04) 1.5 gm.
Magnesium sulphate (MgS04-7H20) 0.75 gm.
Ferric sulphate (Fe2(S04)3-9H20) 0.02 gm.
Water 100.0 cc.
3. Sodium chloride (NaCl) 3.0 gm.
Sodium carbonate (Na2C03) 1.5 gm.
Tap water 100.0 cc.
4. Sodium nitrite (NaN02) 1.5 gm.
Sodium carbonate (Na2C03) 1.5 gm.
Water 100.0 cc.
5. Magnesium sulphate (MgS04-7H20) 0.45 gm.
Sodium chlorid (NaCl) 0 . 75 gm.
Ferric sulphate (Fe2(S04)3-9H20) 0.02 gm.
Water 100.0 cc.
For ammonia oxidation add 1 cubic centimeter of the stock
solutions numbers 1, 2, and 3, to 10 cubic centimeters of the
melted agar.
For the nitrite oxidation, add 1 cubic centimeter of stock
solutions numbers 1, 4, and 5.
Agar media prepared in this way should remain clear.
Medium 54
SiLicio AoiD Gel for Nitrite-forming Bacteria
Prepare a series of silica-gel plates, according to the method
outlined on p. 6. To each plate add 2.5 cubic centimeters of
the medium (a), 1 cubic centimeter of medium (6), 1 drop of
(c) and a few drops of (d) to give a milky appearance. Mix
the solutions and place the open plates in incubator, at 60°C.,
to evaporate excess of surface liquid.
(a) Ammonium sulphate ((NH4)2S04) 1.5 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Magnesium sulphate (MgS04-7HoO) 0 . 25 gm.
Distilled water 100.0 cc.
(6) Ferrous sulphate (FeS04-7H20) 1.0 gm.
Distilled water 100 . 0 cc.
(c) Saturated sodium chloride solution.
(d) A suspension of powdered MgC03 in distilled water.
CULTURE MEDIA 25
MEDIA FOR NITRATE-REDUCING AND DENITRIFYING BACTERIA
Medium 55
AsPARAGiN Nitrate-citrate (Giltay's) Solution
(a) Potassium nitrate (KNO3) 1.0 gm.
Asparagin (C4H8N2O3H2O) 1.0 gm.^
Distilled water 250. 0 cc. J
(h) Citric acid (CeHgOT-HsO) 5.0 gm. ^
or neutral sodium citrate 8.5 gm.y
Monopotassium phosphate (KH2PO4) 1.0 gm.— ,
Magnesium sulphate (MgS04-7H20) 1.0 gm.-^^
Calcium chloride (CaCl2-6H20) 0.2 gm. •
Ferric chloride (FeCla-OHoO) trace ~.
Distilled water 250 . 0 cc. -
Neutralize the citric acid solution with a 10 per cent solution
of sodium or potassium hydroxide, using phenolphthalein as an
indicator. Mix the two solutions and add sufficient water to
make 1000.0 cubic centimeters. If the asparagin and potassium
nitrate are dissolved along with the other salts, a decomposition
may occur. This is marked by a browning of the liquid due
to the presence of nitrous acid.
For a solid medium add 15 grams of agar to 1 liter.
Medium 56
Potassium Nitrate-glucose Solution
Potassium nitrate (KNO3) 1.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Calcium chloride (CaCl2-6H20) 0.5 gm.
Glucose 10.0 gm.
Distilled water 1,000.0 cc.
Medium 57
Potassium Nitrate-ethyl alcohol Solution^
Dipotassium phosphate (K2HPO4) 0.5 gm.
Potassium nitrate (KNO3) 10.0 gm.
Ethyl alcohol (C2H5OH) 5.0 cc.
Tap water 1,000 . 0 cc.
1 Beijerinck, M. W., Centr. Bakt., II Abt., 25: 35, 1910.
26 LABORATORY MANUAL OF MICROBIOLOGY
Medium 58
Potassium Nitrate-filter Papers
Dipotassium phosphate (K2HPO4) 0.5 gm.
Potassium nitrate (KNO3) 2.5 gm.
Filter paper in strips 20 . 0 gm.
Tap water 1,000 . 0 cc.
I Van Iterson, Centr. Bakt., II Abt., 11: 689, 1904.
Medium 59
Potassium Nitrate-thiosulphate Solution^
Sodium thiosulphate (Na2S203-5H20) 5.0 gm.
Potassium nitrate (KNO3) 5.0 gm.
Sodium bicarbonate (NaHCOs) 1.0 gm.
Dipotassium phosphate (K2HPO4) 0.2 gm.
Magnesium chloride (MgCU'OHoO) 0.1 gm.
Calcium chloride (CaCl2-6H20) trace
Ferric chloride (FeCl3-6H20) trace
Distilled water 1,000.0 cc.
2L1ESKE, R., Ber. d. Deutsch. Bot. Gessell, 30: 12-22, 1912.
MEDIA FOR THE REDUCTION OF SULPHATES AND OTHER
SULPHUR COMPOUNDS
Medium 60
ASPARAGIN-SODIUM LACTATE SOLUTION^
Asparagin (C4H8N2O3H2O) 1.0 gm.
Sodium lactate (NaC3H503) 5.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 1.0 gm.
Ferrous sulphate (FeS04-7H20) trace
Tap water 1,000.0 cc.
3 Van Delden, A., Centr. Bakt., II Abt., 11: 83, 1904.
Medium 61
Sodium Lactate-ammonium Sulphate Solution
Ammonium sulphate ((NH4)2S04) 2.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Sodium lactate (NaCsHsOs) 5.0 gm.
Ferrous sulphate (FeS04-7H20) trace
Tap water 1,000.0 cc.
To prepare a solid medium add the above ingredients to 15
per cent gelatin. Heat the medium in a steamer until the
CULTURE MEDIA 27
precipitate has settled, and filter. Sterilize at a low temperature,
about 10 pounds' pressure for 15 minutes, or in the steamer for
20 minutes for 3 consecutive days.
Medium 62
Ye AST- WATER S ODIUM SULPHITE AgAR
Agar 15.0 gm.
Ferric chloride (FeCl3-6H20) or strip of iron. ... 0.1 gm.
Sodium sulphite (Na2S03) 1.0 gm.
Yeast water 1 ,000 . 0 cc.
Reaction pH 7.0.
For the culture of certain microorganisms it is desirable to add
3 per cent of sucrose.
Medium 63
ASPARAGIN-SODIUM LACTATE GeLATIN
Asparagin (C4H8N2O3H2O) 1.0 gm.
Sodium lactate (NaCsH.^Os) 5.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 1.0 gm.
Iron-ammonium sulphate (FeS04-(NH4)2
(S04)-6H.O) trace
Gelatin 120.0 to 150.0 gm.
Distilled water 1,000 .0 cc.
Sterilize in the autoclave at 10 pounds' pressure for 15 minutes.
Cool in ice water.
Medium 64
Beef Extract-sodium Sulphite Agar^
Agar 30 . 0 gm.
Beef extract 3.0 gm.
Peptone 5.0 gm.
Glucose 7 10.0 gm.
Sodium sulphite, 20 per cent solution 10 . 0 cc.
Distilled water 1,000.0 cc.
1 Wilson, W. J., and E. Maud McV. Blair, J. Hijg., 24: 111-119, 1925.
Adjust the reaction to pH 6.6 to 7.0.
Add the glucose just before sterilization and fill in small flasks
of 100 cubic centimeters each. To 100 cubic centimeters of this
agar while melted add 1 cubic centimeter of sterile 8 per cent
ferric chloride (FeCl3-6H20) solution, 0.6 cubic centimeter of a
28 LABORATORY MANUAL OF MICROBIOLOGY
10 per cent sodium hydroxide (NaOH) solution and 10.0 cubic
centimeters of a 20 per cent sodiuni sulphite (Na2S03) solu-
tion. Now dilute this agar medium with an equal amount of
a water suspension of the bacteria. Pour deep plates.
MEDIA FOR SULPHUR-OXIDIZING BACTERIA
Medium 65
Thiosulphate Solution^
Sodium thiosulphate (Na2S203-5H20) 5.0 gm.
Ammonium chloride (NH4CI) 0.1 gm.
Sodium bicarbonate (NaHCOs) 1.0 gm.
Disodium phosphate (Na2HP04-2H20) 0 . 2 gm.
Magnesium chloride (MgCl2-6H20) 0.1 gm.
Tap water 1,000.0 cc.
1 Beijerinck, M. W., Centr. Bakt, II Abt., 11: 593-599, 1904.
Sterilize the thiosulphate and acid carbonate separately in a
small amount of water and when cool add to the solution of the
other salts. A trace of ferrous sulphate (sterile solution) should
also be added after sterilization.
Medium 66
Thiosulphate-nitrate Solution-
Sodium thiosulphate (Na2S203-5H20) 2.0 gm.
Potassium nitrate (KNO3) 10 gm.
Ammonium chloride (NH4CI) 0.1 gm.
Sodium bicarbonate (NaHCOs) 1.0 gm.
Magnesium chloride (MgCl2-6H20) 0.1 gm.
Disodium phosphate (Na2HP04-2H20) 0 . 2 gm.
Distilled water 1,000.0 cc.
2TRAUTWEIN, K., Centr. Bakt. II Abt., 53: 513-548, 1921.
Medium 67
Thiosulphate-agar3
Sodium thiosulphate (Na2S203-5H20) 5.0 gm.
Dipotassium phosphate (K2HPO4) 0.1 gm.
Sodium bicarbonate (NaHCOs) 0 . 2 gm.
Ammonium chloride (NH4CI) 0. 1 gm.
Agar 20.0 gm.
Tap water 1,000.0 cc.
3 Beijerinck, M. W., Centr. Bakt., II Abt., 11: 593-599, 1904.
CULTURE MEDIA 29
For certain organisms it is advisable to add an excess of
CaCOs.
Medium 68
Sulphur-phosphate Medium^
Ammonium sulphate ((NH4)2S04) 0.2 gm.
Monopotassium phosphate (KH2PO4) 3.0 gm.
Magnesium sulphate (MgS04-7H20) 0.5 gm.
Calcium chloride (CaCU-GHsO) 0 . 25 gm.
Ferrous sulphate (FeS04-7H20) trace
Sulphur, powdered 10 . 0 gm.
Distilled water 1,000 . 0 cc.
1 Waksman, S. a., Soil Set., 13: 329, 1922.
The sulphur is weighed out into the individual containers,
usually 1-gram portions into 250-cubic centimeter Erlenmeyer
flasks, and 100-cubic centimeter portions of the liquid medium
added. The reaction of the medium is about pH 4.0. The
flasks are sterilized in flowing steam on 3 consecutive days.
Medium 69
SODIUM-THIOSULPHATE AgAR2
Agar 20 . 0 gm.
Sodium thiosulphate (Na2Si03-5H20) 5.0 gm.
Ammonium chloride (NH4CI) 0.1 gm.
Calcium chloride (CaClo-GHzO) 0 . 25 gm.
Magnesium chloride (MgCl2-6H20) 0.1 gm.
Monopotassium phosphate (KH2PO4) 3.0 gm.
Distilled water 1,000 . 0 cc.
2 Waksman, S. A., /. Bad., 7: 605, 1922.
The medium is prepared as usual and sterilized at 15 pounds'
pressure for 15 minutes.
Medium 70
Solution for the Reduction of Hydrogen Sulphide
Ammonium sulphate ((NH4)2S04) 1.5 gm.
Potassium chloride (KCl) 0 . 05 gm.
Magnesium sulphate (MgS04-7H20) 0.05 gm.
Dipotassium phosphate (K2HPO4) 0 . 05 gm.
Calcium nitrate (Ca(N03)2) 0.01 gm.
Calcium carbonate (CaCOs) 10 . 0 gm.
30
LABORATORY MANUAL OF MICROBIOLOGY
Sterilize in small flasks or tubes. After inoculation incubate
under an atmosphere of hydrogen sulphide as shown below. ^ To
obtain a solid medium, the amount of CaCOs in the above solu-
tion is increased to 50 gm. per liter and 0.5 per cent agar is added.
JS^
Fig. 1. — Apparatus for the cultivation of pure cultures of sulfur bacteria using
H2S as a source of energy. A, apparatus for generation of H2S; B, hydrogen
tank; C, wash flask; D, gasometer; E, culture jar; F, manometer. {Bavendamm.)
1 Bavendamm, W., Die farblosen und roten Schwefelbakterien des
Siiss- und Salzwassers. G. Fischer. Jena, 1924.
MEDIA FOR HYDROGEN AND METHANE BACTERIA
Medium 71
Potassium Nitrate Solution for Hydrogen Bacteria^
Potassium nitrate (KNO3) 2.0 gm.
Mono-sodium phosphate (NaH2P04) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 0.2 gm.
Ferric chloride (FeClg-eHzO) trace
Tap water 1,000.0 cc.
^Lebedeff, A. E., (Russian). Odessa, 1910.
Cultures grown in an atmosphere of hydrogen containing 5 to
15 per cent carbon dioxide.
CULTURE MEDIA 31
Medium 72
Ammonium Chloride Solution for Hydrogen Bacteria*
Sodium bicarbonate (NaHCOs) 1.0 gm.
Ammonium chloride (NH4CI) 1.0 gm.
Monopotassium phosphate (KH2PO4) 0.5 gm.
Ferrous carbonate (FeC03-H20) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 0.1 gm.
Sodium chloride (NaCl) 0.1 gm.
Distilled water 1,000.0 cc.
pH = 7.1 - 7.2.
' RuHLAND, W., Jahrh. Wissen. BoL, 63: 321, 1924.
Use freshly prepared NaHCOs. Iron must be added in the
form of ferrous carbonate. A sterile solution of the iron salt in
water containing CO2 is added to the sterile medium. The
culture is grown in an atmosphere containing hydrogen and
carbon dioxide.
Medium 73
Magnesium Ammonium Phosphate Solution for Methane Bacteria^
Magnesium ammonium phosphate (MgNH4-
P04-6HoO) 0.1 gm.
Dipotassium phosphate (K2HPO4) 0 .05 gm.
Calcium sulphate (CaS04-2H20) 0 . 01 gm.
Distilled water 1,000 cc.
2 Sohngen, N. L., Bot. Centr., 105: 371-372, 1907.
The cultures are placed in an atmosphere consisting of 1 part
methane (CH4) and 2 parts air.
Silica gel or washed agar containing the above nutrients can
be conveniently used as solid media.
MEDIA FOR IRON AND MANGANESE OXIDIZING BACTERIA
Medium 74
Iron-ammonium Sulphate Solution^
Ammonium sulphate ((NH4)2S04) 1-5 gm.
Potassium chloride (KCl) 0 . 05 gm.
Magnesium sulphate (MgS04-7H20) 0.05 gm.
Dipotassium phosphate (K2HPO4) 0 .05 gm.
Calcium nitrate (Ca(N03)2) 0.01 gm.
Distilled water 1,000.0 cc.
sLieske, R., Centr. Bakt., II Abt., 49: 413-425, 1919.
32 LABORATORY MANUAL OF MICROBIOLOGY
Sterilize in small flasks in layers about 2 centimeters deep.
Allow the medium to stand for several days so that it becomes
saturated with oxygen and carbon dioxide. After standing, add
to each flask about 0.05 grams of sterilized iron dust and inoculate.
Medium 75
Manganese-ammonium Sulphate Solution
Manganese bicarbonate, saturated solution 100.0 cc.
Sodium bicarbonate (NaHCOa) 0.1 gm.
Ammonium sulphate ((NH4)2S04) 0.1 gm
Dipotassium phosphate (K2HPO4) trace
Magnesium sulphate (MgS04-7H20) trace
Tap water 900.0 cc.
The manganese bicarbonate is prepared by saturating a sus-
pension of manganese carbonate with pure carbon dioxide. The
medium is sterilized in flowing steam for 25 minutes.
Medium 76
Ferrous Carbonate and Potassium Acetate Solution
Potassium acetate (KC2H3O2) 0.5 gm.
Ferrous carbonate (FeCOa) 0.5 gm.
Tap water 1,000.0 cc.
MEDIA FOR NITROGEN FIXING BACTERIA
Medium 77
Nitrogen-free Mannitol Solution
Mannitol 10.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 0.2 gm.
Sodium chloride (NaCl) 0.2 gm.
Manganese sulphate (MnS04-4H20) trace
Ferric chloride (FeCla-OHzO) trace
Distilled water 1,000.0 cc.
For a solid medium add 12.5 to 15 grams of washed agar to
each liter. For the isolation of Azotobacter prepare this liquid
medium in shallow layers and add sterilized CaCOs to each flask.
CULTURE MEDIA 33
Medium 78
Nitrate Mannitol Agar, with and without Indicators
Same as Medium 77 except that 0.5 gram of potassium nitrate
(KNO3) is added to each Uter and an indicator if desired.
Indicators. — A. Brom thymol blue medium is prepared as
follows: add 5 cubic centimeters of 0.5 per cent alcoholic solu-
tion to 1 liter of the medium.
B. Congo Red medium is prepared as follows: add 10 cubic
centimeters of a 1 to 400 aqueous solution to 1 Uter of the
medium.
Medium 79
Yeast Extract-mannitol Agar
Agar 15.0 gm.
Mannitol 10.0 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 0.2 gm.
Sodium chloride (NaCl) 0.1 gm.
Calcium carbonate (CaCOs) 3.0 gm.
Yeast water^ (reaction pH 6.8) 100.0 cc.
Distilled water 900.0 cc.
^ For directions how to make stock yeast extract see Medium 27, page
16.
If this medium is to be used for the isolation of root nodule
bacteria add Congo red as given under B in Medium 78.
Medium 80
Carrot Extract Agar^
Wash the carrots in running water until all soil particles are
removed, then chop (with meat chopper) into small pieces and
prepare as follows.
Carrots (cut) 250. 0 gm.
Tap water 500.0 cc.
Cook in a steamer for 30 minutes and filter. Neutralize to
pH 7.0 to 7.2 with a strong solution of sodium carbonate and
2 Stapp, C. und G. Ruschmann, Arh. a. d. Biol. Reich, f. Landw. u. Forst.,
13 : 314, 1925.
34 LABORATORY MANUAL OF MICROBIOLOGY
make up to 500 cubic centimeters. Now add 500 cubic centi-
meters of 3.6 per cent neutral water solution of agar. Tube and
sterilize. Avoid excessive heating because the carrot extract
becomes more acid after long heating.
Medium 81
Pea Extract Sucrose Solution
Pea seedlings 3 to 5 inches tall (net weight) 300 . 0 gm.
Water 1,000.0 cc.
Heat in the steamer for 4 to 5 hours and then boil for 1 hour
over a free flame. Filter and make up to 1,000 cubic centimeters.
Add sucrose, 2 per cent, and 0.5 per cent of CaCOs. This
medium contains about 50 milligrams of nitrogen in 100 cubic
centimeters.
Medium 82
Navy-bean Seed Extract Sucrose Medium
Bean seed (dry) 250 . 0 gm.
Tap water 2,500 . 0 cc.
Soak the beans in the cold water for 2 to 3 hours. Now add
water to make 2,500 cubic centimeters and heat in the steamer
for 2 hours. Filter and make up to 2,500 cubic centimeters.
Add 1 per cent of sucrose and 0.5 per cent of CaCOs. This
medium contains about 40 milligrams of nitrogen in 100 cubic
centimeters.
Medium 83
Caffein-bean Extract-glucose for Bacteroid Formation^
Agar 15.0 gm.
Cafifein (C8HioN402-H20) 2.0 gm.
Glucose 10.0 gm.
Bean extract2 1,000 . 0 cc.
iZipPEL, H., Centr. Bakt., II Abt., 32: 107-131, 1911.
2 Bean extract : Add to 20 grams of powdered navy-bean seed in a mortar
20 cubic centimeters of A^/1 KOH. Allow this to stand a few minutes, then
add water sufficient to make 1 Hter. This should stand 24 hours. Siphon
off the clear liquid, neutralize with phosphoric acid.
CULTURE MEDIA 35
Medium 84
Peptone-sucrose Solution for Gum
Peptone 1.0 gm.
Monopotassium phosphate (KH2PO4) 2.0 gm.
Magnesium sulphate (MgS04-7H20) 0.1 gm.
Calcium chloride (CaCl2) 0.1 gm.
Sucrose 20 . 0 gm
Distilled water 1,000.0 cc.
MEDIA FOR CELLULOSE-DECOMPOSING ORGANISMS
Medium 85
Ammonium Sulphate-cellulose Solution^-^
Ammonium sulphate ((NH4)2S04) 1.0 gm.
Dipotassium phosphate (K2HPO4) 1.0 gm.
Magnesium sulphate (MgS04-7H20) 0.5 gm.
Calcium carbonate (CaCOs) 2.0 gm.
Sodium chloride (NaCl) trace
Distilled water 1,000 . 0 cc.
^ Ammonium chloride (NH4CI), diammonium phosphate ((NH4)2HP04),
potassium nitrate (KNO3), or sodium nitrate (NaNOs) may also be used.
2OMELIANSKY, W., Centr. Bakt., II Abt., 8: 226, 1902.
Fill large test tubes about one-third full. Add two or three
strips of filter paper to each tube, part of the paper protruding,
above the surface of the medium. When nitrates are used, the
calcium carbonate may be left out.
Medium 86
Silica-gel Cellulose
Prepare a series of silica-gel plates according to Souleyre
method.
Silica Gel according to Souleyre^
(Modified)
To 40 cubic centimeters of 20 per cent tartaric acid, 1 cubic
centimeter of 60 per cent phosphoric acid and 1 cubic centimeter
sulphuric acid (1:1), add 100 cubic centimeters of potassium silicate
solution (specific gravity, 1.057).^ Let stand until clear, then
filter off the clear solution. This is designated as solution A.
3 Souleyre, M., Compt. rend. d. la Soc. Biol., 93: 306, 1925.
^ 7.6 grams potassium silicate made up to 100-cubic centimeters solution
gives specific gravity = 1.057.
36 LABORATORY MANUAL OF MICROBIOLOGY
Solution B is made by mixing two parts of potassium silicate
solution (specific gravity, 1.085)^ with one part of a 1 per cent
KOH solution. This solution will gel on standing. The alkali
solution (1 per cent) should be mixed with the silicate solution
to make Solution B only a short time before it is to be used. The
solution will keep for about 2 weeks to 1 month.
These two solutions are then titrated against each other with
an indicator, which shows the reaction desired.
The solutions can be sterilized. Solution A is sterilized in an
autoclave for 10 minutes at 110°C. Solution B, however, will
solidify if treated in a like manner. It is necessary, therefore, to
put it in a sterile flask and boil for 5 to 10 minutes.
For growth of bacteria, a medium is prepared as follows:
1. 100 cubic centimeters of A, and 50 cubic centimeters
distilled water.
2. 200 cubic centimeters of nutrient liquid (preferably steri-
lized longer than the silicate solution).
3. Amount of B required to bring medium to desired reac-
tion. These three solutions are cooled as rapidly as possible
after sterilization, and are then poured together. Plates can
then be poured without danger of the gel solidifying before
pouring is completed. Time required for setting of the gel is 1
to 6 hours.
Five grams of thoroughly ground cellulose is suspended in
100 cubic centimeters of the following solution.
Diammonium phosphate ((NH4)2HP04) 5.0 gm.
Magnesium sulphate (MgS04-7H20) 1.0 gm.
Ferrous sulphate (FeS04-7H20) 0.02 gm.
Potassium chloride (KCl) 1.0 gm.
Distilled water 100.0 cc.
About 2 cubic centimeters of the suspension of the cellulose in
the medium is poured upon the surface of each sihca plate, in
such a manner as to have the cellulose uniformly distributed
over the surface; some CaCOs is then powdered on over the
whole surface of each plate. The plates are placed in an incu-
bator at 60 to 65°C., until the excess of water has evaporated,
1 11.6 grams potassium silicate made up to 100-cubic centimeters solution
gives specific gravity = 1.085.
CULTURE MEDIA 37
without allowing the plate to become dry. The dishes are kept
until needed for use.
Medium 87
Ammonium Sulphate Cellulose Agar^
(a) Agar 10 . 0 gm.
Ammonium sulphate ((NH4)2S04) 2.0 gm.
Dipotassium phosphate (K2HPO4) 1.0 gm.
Magnesium sulphate (MgS04-7H20) 1.0 gm.
Sodium chloride (NaCl) 1.0 gm.
Calcium carbonate (CaCOs) 2.0 gm.
Tap water 500.0 cc.
(6) Cellulose solution 500. 0 cc.
1 McBeth, I. G., Soil Sci., 1: 438, 1916.
1. Pour 1,000 cubic centimeters of ammonium hydroxide,
specific gravity 0.90, into a glass-stoppered bottle; add 250 cubic
centimeters of distilled water and 75 grams of pure copper
carbonate; shake the solution vigorously until all the copper is
dissolved (about 10 to 15 minutes are ordinarily required).
2. To the copper-ammonium solution add 15 grams of high-
grade sheet filter paper; shake vigorously at intervals of 10
minutes for 3-^ hour. Examine the solution carefully to see that
the paper is completely dissolved. If any particles of paper
remain in the solution, the shaking must be continued until the
solution is perfectly clear. Dilute 250 cubic centimeters of the
ammonium-copper-cellulose solution to 10 liters with tap water;
add slowly, with frequent shaking, a weak hydrochloric acid
solution prepared by adding 500 cubic centimeters of concentrated
acid to 10 liters of tap water. Continue the addition of the
acid until the blue color disappears; add a slight excess of acid,
shake thoroughly, and allow to stand a few minutes. The finely
precipitated cellulose will rise to the top, due to the large quantity
of free hydrogen liberated in the precipitation process. Shake
the solution vigorously at intervals of a few minutes to dislodge
the hydrogen. As soon as the free hydrogen has escaped, the
cellulose will settle rapidly.
3. Wash through repeated changes of water until free from
copper and chlorine. After the washing is complete, bring the
38 LABORATORY MANUAL OF MICROBIOLOGY
cellulose in the solution up to 0.5 per cent by allowing to settle a
few days and siphoning off the clear solution or by evaporating.
Add the nutrient salts desired, together with 1 per cent of
thoroughly washed agar; heat in autoclave or boil until the agar
is dissolved; tube and sterilize in the usual way.
Medium 88
Cellulose-peptone Agar
Agar (washed) 15.0 gm.
Cellulose ! 2.5 gm.
Peptone 0 . 50 gm.
Dipotassium phosphate (K2HPO4) 0.2 gm.
Magnesium sulphate (MgS04-7H20) 0.2 gm.
Potassium carbonate (K2CO3) 0.4 gm.
Calcium chloride (CaCU) fused 0.02 gm.
Ferric sulphate (Fe2(S04)3-9H20) 0 .02 gm.
Sodium chloride (NaCl) 0.02 gm.
Distilled water 1,000.0 cc.
Dissolve the peptone and salts in 100 cubic centimeters of
distilled water and filter. Add 400 cubic centimeters of the
aqueous cellulose suspension and 500 cubic centimeters of 3
per cent aqueous solution of washed agar (see p. 5).
Medium 89
Cellulose-peptone Medium for Thermophilic Bacteria^
Peptone 5.0 gm.
Calcium carbonate (CaCOs) excess
Sodium ammonium phosphate (Na(NH4) HPO4-
4H2O) 2.0 gm.
Monopotassium phosphate (KH2PO4) 10 gm.
Magnesium sulphate (MgS04-7H20) 0.3 gm.
Calcium chloride (CaClo) 0.1 gm.
Ferric chloride (FeCl3-6H20) trace
Water 1,000.0 cc.
1 ViLJOEN, J. A., E. B. Fred, and W. H. Peterson, /. Agr. Set., 16: 1,
1926.
Reaction about pH 7.4. Add about 0.3 gram of paper pulp
or of cut up filter paper to 20 cubic centimeters of liquid in long
test tubes.
CULTURE MEDIA 39
MISCELLANEOUS MEDDl
Medium 90
Litmus or Brom Cresol Purple Milk
Litmus milk is one of the most important culture solutions.
Because of the great variety of organisms which grow in it, and
the decided changes which they produce, litmus milk is very
useful in the classification of bacteria. All organisms should be
inoculated into litmus milk and the changes after various
periods of time recorded. To secure the best results prepare
as follows :
Fresh milk, immediately after milking, should be centrifuged
and this fat-free milk taken directly to the laboratory. Pipette
5- or 10- cubic centimeter portions of the fresh milk into pre-
viously sterilized test tubes, plug with cotton, and sterilize for
10 minutes in the autoclave at 8 to 10 -pounds^ pressure. Cool in
water and repeat sterilization 24 hours later. After the last
sterilization incubate the tubes for a week or more at 28 to 30°C.
and note if there are changes. Now add to each tube of the
bacteria free milk about 2 to 3 drops of a sterihzed saturated
solution of high-grade lime-free blue litmus (litmus 1 gram and
water, 15 cubic centimeters). Litmus milk should give a
lavender color, not too deep, which turns red in the presence of
acids and blue in the presence of alkalies.
Brom cresol purple may be used in place of litmus. For
this purpose, prepare a stock solution as follows: 0.5 gm. of
brom-cresol purple should be ground in a mortar with 14 cubic
centimeters of .V/10 sodium hydroxide and made up to 100 cubic
centimeters with distilled water. Take 10 cubic centimeters of
this 0.5 per cent solution to 1,000 cubic centimeters of milk or
about 2 to 3 drops to 10 cubic centimeters of milk.
Medium 91
Potato
Select large potatoes, wash and scrub well with a stiff brush.
Peel and cut in wedge-shaped blocks about 4 to 6 centimeters
long and 1.5 centimeters wide. The size will depend upon the
40 LABORATORY MANUAL OF MICROBIOLOGY
test tubes. Allow the cut potatoes to stand in running water
for at least 12 hours.
Place a small piece of glass rod or cotton in the bottom of the
test tube and insert the potato wedge into the tube. Fill the
tube with water until the potato is entirely under the liquid.
Plug and sterilize: about 15 minutes at 10 pounds' steam pressure
for 3 consecutive days will usually be found sufficient. Just
before use, pour off the excess water.
Potatoes prepared in this way should retain their white
color.
Medium 92
Mannitol Glycero-phosphate Solution
Mannitol 10.0 gm.
Potassium nitrate (KNO3) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 0.2 gm.
Sodium chloride (NaCl) 0 . 2 gm.
Calcium glycero-phosphate 1.0 gm.
Water 1,000.0 cc.
Adjust reaction to pH 6.5 to 7.0.
Medium 93
Peptone-sucrose-lactose Solution
Sucrose 10.0 gm.
Peptone 20 . 0 gm.
Lactose 10.0 gm.
Sodium chloride (NaCl) 2.0 gm.
Dipotassium phosphate (K2HPO4) 2.0 gm.
Magnesium sulphate (MgS04-7H20) 1.0 gm.
Tap water 1,000.0 cc.
Medium 94
Peptone-sodium Phosphate Solution
Peptone 10.0 gm.
Disodium phosphate (Na2HP04-2H.>0)i 5 .0 gm.
Carbohydrate 10.0 gm.
Tap water 1,000.0 cc.
1 To prepare disodium phosphate (Na2HP04-2H20) take ordinary
disodium phosphate with 12 H2O and spread it out on filter paper and allow
it to remain at room temperature in a dry place for 2 weeks.
CULTURE MEDIA 41
Reaction pH 7.1 to 7.2.
Media 92, 93, 94 are recommended for lactic acid-forming
bacteria of milk and milk products.
Medium 95
Malt Extract-peptone Solution
Malt (finely ground) 30 . 0 gm.
Peptone 10.0 gm.
Lactose 20 to 40 gm.
Calcium carbonate 30 . 0 gm.
Tap water 1,000 . 0 cc.
The malt is extracted in the water by boiling for 1 hour; the
other constituents are then added.
Medium 96
Paraffin Agar for Hydrocarbon-destroying Organisms^
Ammonium chloride (NH4CI) 0.5 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 0.5 gm.
Paraffin oil 10.0 gm.
Agar 20 . 0 gm.
Distilled water 1,000.0 cc.
1 Sohngen, N. L., Centr. Bakt, II Abt., 37: 595-609, 1913.
Medium 97
Solution for Fat Decomposition^
Dipotassium phosphate (K2HPO4) 5.0 gm.
Ammonium phosphate (NH4)3P04) 5.0 gm.
Magnesium sulphate (MgS04-7HoO) 1.0 gm.
Calcium chloride (CaCl2) 1.0 gm.
Ferric chloride (FeCl3-6H20) trace
Sodium chloride (NaCl) trace
Distilled water 1,000.0 cc.
2 Rahn, 0., Centr. Bakt., II Abt., 15 : 423, 1906.
Molten fat is placed in a flask and, on inclining the flask, the
fat is allowed to solidify. The above nutrient solution
is then added. Three-tenth to five-tenth per cent of finely
powdered fat may also be used with the above solution.
42 LABORATORY MANUAL OF MICROBIOLOGY
Medium 98
Peptone-beef Extract Gelatin Agar^
(a) Sodium chloride (NaCl) 5.0 gm.
Monopotassium phosphate (KH2PO4) 0.5 gm.
Dipotassium phosphate (K2HPO4) 1.5 gm.
Water ■ 100.0 ec.
(6) Gelatin 4.0 gm.
Glucose 0 . 05 gm.
Peptone 0.1 gm.
Beef infusion 5.0 cc.
Water 400 . 0 cc.
1 Frazier, W. C., J. Inf. Dis., 39: 302, 1926.
Pour (a) and (h) together, heat in flowing steam and mix with
500 cubic centimeters of a 3 per cent agar (washed) solution.
Adjust pH to 7.0.
Pour duplicate plates and inoculate in the center. After
48 hours or longer at 30°C. make tests.
Flood one plate with 1 per cent tannic acid and the duplicate
with acid HgCl2 solution.^
^HgCU 15 grams, HCl (Cone) 20 cubic centimeters, and 100 cubic
centimeters of water.
If the gelatin has been changed, a clear zone will appear about
the giant colony in the plate to which HgCl2 has been added.
This requires 15 to 30 minutes.
The tannic acid plate will show the amount of decomposition
of the gelatin.
Medium 99
Casein Hydrolysis on Milk Agar
(a) Agar 15.0 gm.
Water 500.0 cc.
(6) Milk 500.0 cc.
Tube separately in portions for plating. Sterihze. Cool to
45°C. and pour plates using one tube of each per plate.
Streak the culture to be tested on the surface of the milk agar.
After incubation examine the plates for clearing along line of
growth. Flood plates with dilute acid (HCl 1:10). If clearing
remains, it denotes true hydrolysis of casein. If not, it is weak
acid clearing.
CULTURE MEDIA 43
Medium 100
Sodium Citrate for the Colon Aerogenes Group^
Sodium chloride (NaCl) 5.0 gm.
Magnesium sulphate (MgS04-7H20) 0.2 gm.
Monoammonium phosphate (NH4H2-POi) 1.0 gm.
Dipotassium phosphate (K2HPO4) 1.0 gm.
Sodium citrate (S^^HaO) 2 . 77 gm.
Distilled water 1,000.0 cc.
1 KosER, S. A., J. Bad., 9: 63, 1924.
Bacterium coli of fecal origin is unable to utilize citrate, while
the B. aerogenes-cloacae types all utilize citrate.
Medium 101
Ferric Ammonium Citrate-nitrate Solution
Ammonium sulphate ((NH4)2S04) 0.5 gm.
Sodium nitrate (NaNOs) 0.5 gm.
Dipotassium phosphate (K2HPO4) 0.5 gm.
Magnesium sulphate (MgS04-7H20) 0.5 gm.
Calcium chloride (CaCla) 0.2 gm.
Ferric ammonium citrate 10.0 gm.
Distilled water 1,000.0 cc.
The ferric ammonium citrate should be sterilized separately
and added to the salt solution when cool. If a solid medium is
wanted, add 1.5 per cent of agar.
The decomposition of the organic radical is accompanied by
the precipitation of the iron; thus in an agar medium brown
zones are formed around the colonies.
Medium 102
Corn Mash
Corn meal (not degerminated) 50 to 80 gm.
Tap water 1,000 cc.
Mix the corn meal with cold water. Steam for 2 to 3 hours
and tube in long test tubes. Sterihze for 2 hours at 120°C.
Medium 103
Sheep- or Beef-brain for Anaerobic Bacteria
1. Boil sheep brain with equal volume of water.
2. Decant water (save) and press brain through a potato ricer.
44 LABORATORY MANUAL OF MICROBIOLOGY
3. Add decanted water, 2 per cent peptone and 0.1 per cent
glucose.
4. Tube by punching through the fiUing funnel with a glass rod.
5. Sterilize intermittently in an Arnold sterilizer.
Five daily runs of 30 minutes each are recommended. Steri-
lization in an autoclave is usually a failure because of the tendency
of the medium to climb on to the plugs. However, if long tubes
are used, this medium may be sterilized in the autoclave. The
finished medium is nearly white with a clear supernatant liquid.
The small amount of glucose is to stimulate growth; excess
would prevent or interfere with putrefaction and blackening.
Peptone stimulates early development and intensifies the blacken-
ing produced by those organisms liberating H2S.
Medium 104
Nitrate Solution for Higher Green Plants
(a) Calcium nitrate (Ca(N03)2-4H,0) 100.0 gm.
Potassium nitrate (KNO3) 25. 0 gm.
Sodium chloride (NaCl) 15. 0 gm.
Distilled water 1,000 . 0 cc.
(h) Monopotassium phosphate (KH2PO4) 25.0 gm.
Distilled water 1,000.0 cc.
(c) Magnesium sulphate (MgS04-7H20) 50 . 0 gm.
Distilled water 1,000. 0 cc.
(d) Ferric chloride (FeCl;r6H20) 5.0 gm.
Distilled water 250 . 0 cc.
Take 10 cubic centimeter portions of solutions (a), (6), and
(c) to 1,000 cubic centimeters of water. Add 1 to 2 drops of
solution (d).
Medium 105
Solution for Growing Higher Plants
(a) Ammonium nitrate (NH4NO3) 32.0 gm.
Distilled water 1,000.0 cc.
(6) Monocalcium phosphate (CaH4(P04) 2. H2O). 10.0 gm.
Distilled water 1,000.0 cc.
(c) Potassium sulphate (K2SO4) 20 . 0 gm.
Distilled water 1,000.0 cc.
(d) Magnesium sulphate (MgS04-7H20) 8.0 gm.
Distilled water 1,000.0 cc.
(e) Ferric chloride (FeCla-OHzO) 0 . 1 gm.
Distilled water 250 .0 cc.
CULTURE MEDIA 45
Prepared with ammonia-free water and chemically pure salts.
Dilute 10-cubic centimeter poritons of (a), (6), (c), and {d)
and 1 cubic centimeter of (e) in 1,000 cubic centimeters of water.
If a nitrogen-free medium is desired, omit (a). Plant food solu-
tions should be renewed at regular intervals of about one week
each.
Medium 106
Modified Crone's Nitrogen-free Solution^
Stock Salt Mixture:
Potassium chloride (KCl) 10 . 0 gm.
Calcium sulphate (CaS04-2H20) 2 . 5 gm.
Magnesium sulphate (MgS04-7H20) 2.5 gm.
Tricalcium phosphate (Ca3(P04)2) 2.5 gm.
Ferric phosphate (FeP04) 2.5 gm.
1 Bryan, 0. C, Soil Sci., 13, 279, 1922.
Mix all of these salts and grind to a fine powder.
Water 1,000.0 cc.
Stock salt mixture 1.5 gm.
If a solid medium is wanted, take 7.5 grams of washed agar to 1
liter. After sterilization, shake until salts are well mixed with
agar and allow to harden.
Medium 107
Preserving Stock Cultures in Soil
Mix equal parts of a fertile soil of neutral reaction with sand.
Fill into test tubes, about 2-inch layers and sterilize without
cotton plugs for 5 to 6 hours at 120°C. Now add sterihzed water
to bring the moisture of the soil to about two-thirds saturation
and plug tubes with cotton. Again sterilize for 2 hours. Add
to a few of the tubes, glucose niitrient broth and incubate for
at least a week. If there are no signs of growth, the soil tubes
are ready for use. To prevent mold growth, through plugs, store
these tubes in a dry place.
Medium 108
Preserving Stock Cultures in Corn Mash
Same as Medium 102. This medium will be found useful for
cultures of lactic-acid-forming bacteria.
46 LABORATORY MANUAL OF MICROBIOLOGY
Medium 109
Preserving Stock Cultures in Meat Infusion, Peptone, Gelatin
Phosphate Agar^
Medium 110
Preservation of Stock Cultures in Vacuo^
111
Preserving Plate Cultures
Washed agar 20 . 0 gm.
Glycerol (C3H5(OH)3) 500.0 cc.
Distilled water 500 . 0 cc.
Dissolve the agar in the water by heating in a steamer, add the
glycerol, and filter through glass wool.
FAVORABLE CONDITIONS FOR THE DEVELOPMENT OF
ANAEROBIC BACTERIA
Living Vegetable Tissue
To an ordinary desiccator or a museum jar with tightly fitting
cover add finely chopped raw potato, carrots, lettuce or similar
vegetable tissue. If potato is used, about 50 grams per liter of
air will be found satisfactory. Now add the tubes or plate
cultures of bacteria and seal the jar. The respiring plant tissue
rapidly absorbs the free oxygen and gives off CO2 thus bringing
about favorable conditions for the growth of anaerobic bacteria.
Sterile milk plus a small amount of methylene blue may be
used as an indicator for anaerobiosis.
Pyrogallic Acid for Absorbing Oxygen
For every 100 cubic centimeters of air space take 1 gram of
pyrogallic acid and 10 cubic centimeters of a 10 per cent solution
of sodium or potassium hydroxide.
Note. — To prepare an anaerobic jar, cover the bottom of
the jar with 3^-inch layer of pyrogalUc acid. Fit the
cover tightly to the jar with vaseline and remove the air with a
suction pump, and when there is a good vacuum, run in 75 to 100
cubic centimeters alkali solution.
1 Ayres, S. H.. and W. T. Johnson, J. BacL, 9: 112, 1924.
2 Brown, J. H., Science, 64: 429, 1926.
PART II
METHODS OF STAINING OF BACTERIA
The general structure of bacteria and other microorganisms
is most easily seen in stained preparations. The process of
staining consists in smearing a suspension of the organisms over
a clean slide, drying at room temperature, fixing to the glass by
passing through a flame two or three times (do not burn) and
staining the film by one of the methods outlined below. After
staining, the film is washed in running water, dried with a blotter,
and examined. To preserve the mount, a small drop of balsam
is placed over the film and the cover glass is pressed down gently
to force out the excess of balsam. Set the slide in a warm place
to dry.
Some of the most important laboratory stains are (1) methy-
lene blue or thionin, (2) crystal violet, (3) fuchsin, (4) erythrosin,
and (5) nigrosin. Stock solutions of methylene blue, thionin,
crystal violet, and fuchsin should be prepared and kept on hand.
For the stocks, make a saturated solution of the dyes in 95 per
cent ethyl alcohol. Filter through paper a small amount of the
dye and dilute as given in the directions.
For quick preparations, not deeply stained, the methylene
blue will be found satisfactory. Fuchsin, on the other hand,
possesses unusual penetrating power for staining bacterial cells
and spores and will be found useful for a great many kinds of
bacteria. If heated these dyes penetrate much more rapidly
and thus give more deeply stained mounts.
Loeffler's Alkaline-methylene Blue
Saturated alcoholic solution of methylene blue 30.0 cc.
Solution of potassium hydroxide in distilled water
(1 : 10,000) 100 . 0 cc.
Ziehl's Carbol-fuchsin
Saturated alcoholic solution of basic fuchsin 10.0 cc.
Carbolic acid, 5 per cent aqueous solution 100.0 cc.
47
48 LABORATORY MANUAL OF MICROBIOLOGY
If used in staining root-nodule bacteria dilute the carbol
fuchsin 1 part of stain to 9 parts of distilled water.
Gram Stain
This is one of the most important methods of staining employed
in a study of bacteria. It is commonly used to distinguish
certain organisms from others and also to show the general
morphology. It has been found that when stained with gentian
or crystal violet, and treated with an iodine solution, some
organisms are easily decolorized while others are much more
difficult to decolorize.. The method follows. Prepare a thin
smear of the culture on a slide. Dry in the air and fix with heat.
Stain 1 minute with the crystal violet dye and then treat with the
iodine solution for 1 minute.
Crystal Violet
Saturated alcoholic solution of crystal violet 10.0 cc.
Ammonium oxalate, 1 per cent aqueous solution 40.0 cc.
Lugol's Iodine Solution
Iodine 1.0 gm.
Potassium iodide 2.0 gm.
Water 300.0 cc.
Decolorize for 30 seconds in 95 per cent alcohol. Wash
between each step. Counter stain for 10 seconds with safranin.
Wash and dry.
Safranin
Saturated alcoholic solution of safranin 10.0 cc.
Distilled water 100.0 cc.
If gram-positive, the cells should retain the crystal violet
stain and thus appear under the microscope as a deep violet.
If gram-negative, the cells are decolorized by the alcohol and
thus show a pale pink color.
Carbol Thionin (Nicolle's)
Prepare a stock saturated alcohoUc solution of thionin in 50
per cent alcohol.
Take 10 cubic centimeters of this saturated solution of thionin
and 100 cubic centimeters of a 2 per cent carbohc acid solution.
METHODS OF STAINING OF BACTERIA 49
This is an excellent bacterial stain.
Erythrosin^
(a) Erythrosin 5.0 gm.
Alcohol (70 per cent) 100 . 0 cc.
(6) Erythrosin 1.0 gm.
Carbolic acid (5 per cent) 100 . 0 cc.
This stain is especially recommended for root-nodule bacteria.
1. Place a drop of the fresh culture on a glass slide, tilt the
slide to allow drop to spread. Dry the film in an oven at 45°C.
and fix in absolute alcohol.
2. After the alcohol evaporates, flood the mount with (a) and
allow to stain for 10 minutes.
3. If the stain is not deep enough, wash off this alcohol ery-
throsin and again stain with (6) for 10 minutes.
1 WiNOGRADSKY, S. Ann. Inst. Pasteur, 39: 325, 1925; Gangulee, N.
Ann. Appl. Biol, 13: 364, 1926.
Capsule Stain Adapted from Dorner Spore Stain
1. Spread a loopful of culture on clean glass slide. Air dry
and do not fix.
2. Stain 1 to 2 minutes with cold carbol fuchsin. Wash
gently in water.
3. Cover with thin layer of nigrosin by spreading a loopful
of saturated aqueous solution of nigrosin. Dry quickly to
prevent decolorization of the cells.
Cells will be red and capsules white against a blue-gray
background.
Capsule Stain. Negative Method with Ink
Place a loopful of liquid culture on a clean glass slide. Add
to it a loopful of ink (small loop carrying about one-fourth as
much liquid). Cover with a cover slip and examine as a wet
mount. The bacteria will be white against a gray background;
their capsules will be unstained and appear as halos around the
denser bacterial cells.
By this method one can demonstrate some capsules which
cannot be stained by any positive staining method.
The ink to be used is Burri's Pelikan Tusche, No. 541 (Griibler).
It is bacteria free when received and must be kept aseptically.
50 LABORATORY MANUAL OF MICROBIOLOGY
Ordinary India ink cannot be used because of the bacteria which
it contains.
Spore Stain (Dorner Modified)
1. Spread a loopful of culture, suspension or liquid culture,
on a clean glass slide ; dry and fix by heat.
2. Cover with Ziehl's carbol fuchsin. Steam over water
bath for 10 to 20 minutes. Do not allow fuchsin to dry on smear.
Add more fuchsin if needed. Wash in water.
3. Spread thinly over the smear a loopful of saturated aqueous
nigrosin solution. Spread either with wire loop or with a glass
slide as for blood smears. If the nigrosin is not well spread the
first time it may be washed off with water and more nigrosin
applied.
4. Dry without heating and examine in oil, or mount in Canada
balsam.
Spore Stain (Lagerberg's Modified)
1. Make film, fix by heat, and cover with small amount of
5 per cent chromic acid.
2. After 30 seconds, add about twice as much concentrated
ammonia as there is chromic acid on the slide. Allow to act
about 2 minutes.
3. Rinse with tap water and steam with carbol fuchsin for 2
or 3 minutes.
4. Rinse and destain with 1 per cent sulphuric acid for 15 to
30 seconds.
5. Rinse again and flood slide with the tap water.
6. Add to this a few drops of Loeffler's methylene blue and
allow to stain for 10 to 30 seconds.
7. Rinse, blot, dry, and examine.
Nigrosin for Negative Mounts
1. Place a loopful of culture on a clean glass sfide, spread, and
allow it to air dry.
2. Spread thinly over the smear a loopful of saturated aqueous
nigrosin solution. Spread either with the wire loop or with a
glass sHde (as for blood smears). Dry and examine in oil, or
mount in Canada balsam.
METHODS OF STAINING OF BACTERIA 51
By the use of nigrosin it is possible to examine organisms
unstained. There are many points in favor of this method, e.g.,
the organisms do not shrink or change their form. Nigrosin may
be used in demonstrating the motihty of bacteria. For this
purpose add a very small amount of nigrosin to a hanging drop.
Congo Red for Negative Mounts
(For differentiating living and dead bacteria)
1. Place a drop of 2 per cent aqueous Congo red solution
(free from bacteria) on a clean glass slide.
2. Mix with it a loopful of the bacterial culture.
3. Allow it to dry thoroughly in air 10 minutes or more.
4. Flood with acid alcohol (1 or 2 per cent HCl). This
changes the color to blue and fixes the film.
5. Dry without washing and examine in oil, with or without
cover glass. Living cells appear unstained-white against blue.
Unless preserved with oil or balsam the preparations fade
rapidly.
This method of preparing negative mounts is recommended
for root-nodule bacteria. The active living cells are negative
while the dead cells are more or less positive.
Erythrosin for Direct Staining of Soil Bacteria
1. Prepare a suspension of soil (about 1 : 10) in a 0.015 per cent
solution of gelatin.
2. Smear a thin film on a slide and dry.
3. Immerse in 40 per cent acetic acid for 1 to 3 minutes, wash
and dry on a water bath.
4. While on the water bath, stain for 1 minute with 1 per cent
aqueous solution of erythrosin Y or rose bengal.
Barlow Stain for Root Nodule Bacteria
(See p. 109)
PART III
QUALITATIVE AND QUANTITATIVE METHODS OF
ANALYSIS
PREPARATION OF REAGENTS
Prepare stock solutions of the indicators in dropping bottles.
Phenolphthalein. — Dissolve 1 gram of pure phenolphthalein in
100 cubic centimeters of 86 per cent alcohol. This indicator
is recommended for the titration of organic and inorganic acids
and strong bases. It should not be used for the titration of
ammonia.
Methyl Orange. — Dissolve 0.02 gram of solid methyl orange
in 100 cubic centimeters of hot water, allow to cool, and, if a
deposit forms, filter. If the sodium salt is used instead of the
acid, take 0.022 gram to 100 cubic centimeters of water. Add
0.67 cubic centimeters of 0.1 N hydrochloric acid, let stand, and
filter. Methyl orange is used for the titration of strong acids.
Litmus. — This indicator is obtained from a species of lichen.
It is widely used in bacteriology, especially in milk. The chief
coloring principle is azolitmin. Dissolve 5 grams of purified
litmus in 100 cubic centimeters of water. Heat in the steamer
for 1 to 2 hours, with occasional shaking. Allow to settle for
several days and then decant the clear liquid.
Cochineal. — Take 6 grams of cochineal to 50 cubic centimeters
of alcohol (95 per cent) and 200 cubic centimeters of distilled
water. Shake the cochineal in the mixture of water and alcohol.
Allow to stand for two days at room temperature. Filter until
clear. The color of this solution should be a deep ruby red; in
the presence of alkali a violet color, and in the presence of acid
a yellowish-red color.
Sodium Alizarine Sulfonate. — Dissolve 1 gram in 100 cubic
centimeters of distilled water by warming. Filter and make
up to volume.
52
QUALITATIVE AND QUANTITATIVE ANALYSIS
53
A list of the color change and pH range of some of the more
common indicators is given below.
The indicators for measuring the hydrogen-ion concentration
are prepared as follows: 0.1 gram of the dry powder is ground in
a mortar with varying amounts of N/20 NaOH and diluted to
25 cubic centimeters with water.
N/20 NaOH
Used, Cubic
0.1 Gram of Indicator Centimeters
Phenol red 5.7
Brom phenol blue 3.0
Cresol red 5.3
Brom cresol purple 3.7
Thymol blue 4.3
Brom thymol blue 3.2
Methyl red 7.4
For colorimetric tests, dilute the stock solution of 25 cubic
centimeters with distilled water to 250 cubic centimeters or a
concentration of 0.04 per cent of the indicator. Take 5 drops of
this dilute solution to 10 cubic centimeters of the liquid to be
tested.
Color Change and pH Range of Certain Indicators
Common name
Color change Range pH
Thymol blue (acid) . . . .
Brom phenol blue
Brom cresol green
Methyl red
Brom cresol purple
Chlor phenol red
Brom thymol blue ....
Phenol red
Cresol red
Thymol blue (alkaline)
Phenolphthalein
Methyl orange
Litmus
Cochineal
Alizarin red
Congo red
Red-yellow
Yellow-blue
Yellow-green
Red-yellow
Yellow-purple
Yellow-red
Yellow-blue
Yellow-red
Yellow-red
Yellow-blue
Colorless-red
Orange red-yellow
Red-blue
Yellow-lilac
Yellow-pink
Blue-red
1.2 to
3.0 to
Oto
.4 to
.2 to
.2 to
6.0 to
6.8to
7.2 to
8.0 to
8.3 to
3.1 to
4.5 to
4.8 to
5.0 to
3.0 to
2.8
4.6
5.6
6.0
6.8
6.8
7.6
8.4
8.8
9.6
10.0
4.4
8.3
6.2
6.8
5.0
54 LABORATORY MANUAL OF MICROBIOLOGY
PREPARATION OF STANDARD SOLUTIONS
A great many methods have been described for the standard-
ization of solutions. Only a brief outline will be given of certain
of these methods; for detailed directions the reader is referred to
some of the recent textbooks on volumetric analysis.
Normal Sulphuric Acid Solution
A normal solution of sulphuric acid is one-half the molecular
weight of H2SO4 in grams, diluted to 1 liter with distilled water.
Since the molecular weight of sulphuric acid is (2 + 32 + 64) 98,
then 49 grams, one-half of 98, is the amount necessary for each liter.
1. In order to secure 49 grams of H2SO4, it requires 49 divided
by 1.80 (specific gravity of concentrated sulfuric acid), or 27.2
cubic centimeters of chemically pure acid. To be sure that
sufficient acid has been used, measure out about 27.5 cubic
centimeters of acid.
2. Place the acid in a 1,000-cubic centimeter Erlenmeyer flask,
containing about 500 cc. of water, mix well and transfer to a
volumetric flask; make up to 1,000 cubic centimeters with water
and mix carefully.
3. From this mixture remove 10-cubic centimeter portions,
accurately measured in a 10-cubic centimeter pipette, and
place in weighing bottles which have been thoroughly cleaned,
dried in an oven, cooled, and weighed.
4. One cubic centimeter of chemically pure ammonia is added
to each weighing bottle to neutralize the sulphuric acid.
5. The water and excess of ammonia is then evaporated in an
oven at 100°C. and the ammonium sulphate remains behind.
If the chemicals are pure, 100 cubic centimeters of the solution
should give 49 grams of sulphuric acid. In 10 cubic centimeters
of the solution there should be 0.49 gram of H2SO4.
H2S04:(NH4)2S04::98:132
49 : X ::98:132
X =0:66
If the solution is exactly normal, there should be 0.6600 gram of
(NH4)2S04 formed from 10 cubic centimeters. In case the
amount of (NH4)2S04 formed is too great, its factor is determined
by dividing 0.6600 into the weight of ammonium sulphate found.
QUALITATIVE AND QUANTITATIVE ANALYSIS 55
If, for instance, the weight of ammonium sulphate is 0.6675, the
factor of the solution is 1.01 13 + . This means that 10 cubic
centimeters of the solution is equivalent to 10.113 cubic centi-
meters of normal solution.
Normal Sodium Hydroxide Solution
To prepare a normal solution of sodium hydroxide, weigh out
roughly 45 grams of caustic soda (wash off the carbonate from
the surface of the sticks of alkali) and dissolve in a little more
than a liter of water. Allow this solution to stand until cool
and titrate against A^/5 acid potassium phthalate. The prepa-
ration of N/5 phthalate follows :
Molecular weight of KHC8H4O4 = 204.14.
Normal phthalate = 204.14 grams in 1,000.0 cubic centimeters
of water.
Fifth normal phthalate = 40.83 grams in 1,000.0 cubic
centimeters of water.
Weigh out 20.414 grams of dried (at 110 to 115°C.) phthalate.
Transfer to a 500-cubic centimeter flask. Dissolve in water
and make up to volume.
To standardize a normal solution of NaOH, measure 50 cubic
centimeters of the N/5 phthalate solution with a pipette and
titrate with phenolphthalein as the indicator; 50 cubic centi-
meters A^/5 phthalate = 10 cubic centimeters of N/1 NaOH.
Example. — 50 cubic centimeters N/5 phthalate = 9.36 cubic centimeters
of the approximately N/1 NaOH. ^
^ = 1.068 factor for iV/1 NaOH.
Make necessary dilution if an exact N/1 solution is desired.
To standardize an N/10 NaOH solution, dilute 20 cubic centimeters of
phthalate solution to 100 cubic centimeters and titrate 25 cubic centimeter
aliquots with the alkali. Calculate the factor as above or dilute if an exact
N/10 solution is desired.
Standard (iV/6) Solution of Oxalic Acid
To prepare a standard, iV/6 solution of oxalic acid, dissolve
63 grams of C.P. oxalic acid [(COOH)2-2H20] in 1 liter of dis-
tilled water by warming. Then dilute to exactly 6 liters with
distilled water. This solution is standardized against the sodium
56 LABORATORY MANUAL OF MICROBIOLOGY
hydroxide solution, using phenolphthalein as an indicator.
One cubic centimeter of this solution is equivalent to exactly 1
milligram of carbon in carbon dioxide determinations.
Standard (A^/6) Barium Hydroxide Solution
To prepare one liter of a A^/6 barium hydroxide solution,
14.28 grams of anhydrous barium hydroxide is theoretically
required. However, in the process of preparation of the solution,
considerable barium carbonate is formed. Hence about one-
fourth to one-sixth more of the hydroxide is taken than is
required theoretically. Add 100 grams of C.P. anhydrous
barium hydroxide to 6 liters of distilled water. Allow to dissolve
for 24 hours with occasional shaking. The residue is allowed to
settle for 7 days, when the supernatant solution is filtered through
a double layer of filter paper. The solution is standardized to
exactly iV/6 against the oxaHc acid, using phenolphthalein as an
indicator.
QUALITATIVE METHODS FOR THE DETERMINATION OF VARI-
OUS FORMS OF INORGANIC NITROGEN
Nessler's Reagent for Ammonia
1. Dissolve 50 grams of potassium iodide in a small quantity
of cold distilled water. (Ammonia, free; about 35 c.c.)
2. Add a saturated solution of mercuric chloride until a slight
precipitate persists.
3. Now add 400 cubic centimeters of a 50 per cent solution of
potassium hydroxide made by dissolving the potassium hydroxide
and allowing it to clarify by sedimentation before using.
4. Dilute to 1,000 cubic centimeters, allow to settle for 1
week, and decant. This solution gives the required color with
ammonia within 5 minutes after addition.
5. Keep the Nessler's solution in a well-stoppered bottle
away from the light.
Test for Ammonia. — Add to a drop of Nessler's solution in a
test plate a loopful of the solution to be tested. A deep golden-
yellow color indicates the presence of ammonia.
The presence of glucose in solution interferes with the ammonia
test. In that case, the ammonia should be removed first by
aeration and then tested (see p. 63).
QUALITATIVE AND QUANTITATIVE ANALYSIS 57
Thomas' Reagent for Ammonia
Prepare a 5 per cent solution of phenol and a sodium hypochlo-
rite solution containing 1 per cent available chlorine: 1 cubic
centimeter of the NaOCl solution will neutraHze 2.86 cubic
centimeters of O.IN sodium thiosulphate.
Dilute the culture, 0.2 to 1.0 cubic centimeter with 8 cubic
centimeters of water and add 1 cubic centimeter of phenol and
1 cubic centimeter of hypochlorite. Let stand for J^ hour. A
blue color indicates ammonia or amines.
Trommsdorf's Reagent for Nitrites
1. Add slowly, with constant stirring, a boiling solution of 20
grams of zinc chloride in 100 cubic centimeters of distilled water
to a mixture of 4 grams of starch in water. Continue heating
until the starch is dissolved as much as possible, and the solution
is nearly clear.
2. Then dilute with water and add 2 grams zinc iodide.
3. Dilute to 1 Hter and filter.
4. Store in well-stoppered bottles in the dark.
Test for Nitrites. — Place 3 drops of Trommsdorf's reagent
in depression on test plate. Add 1 drop of dilute sulphuric acid
(1:3). Remove a loopful of the solution to be tested and touch
to surface of reagent. A blue color indicates the presence of
nitrites.
Reagents for Nitrate, Nitrite, and Hydroxylamine
(a) Dissolve 10.5 grams sulfanilic acid and 6.8 grams sodium
acetate in 1,000 cubic centimeters of 30 per cent (by volume)
acetic acid, by heating on water bath. Boil for 3 minutes and
make up to 1,000 cc.
(b) Boil 5 grams a-naphthylamine in 700 cubic centimeters of
water for 5 minutes, then filter and add 5 cc. of concentrated
HCl, cool and make up to 1,000 cc.
Solutions (a) and (h) are always kept separate.
(c) Dissolve 1.3 grams iodine in 100 cubic centimeters of glacial
acetic acid.
{d) Dissolve 2.5 grams Na2S203-5H20 in 100 cubic centi-
meters of water.
58 LABORATORY MANUAL OF MICROBIOLOGY
(e) Zinc dust is freed from traces of nitrates and nitrites
by treating some of the pure commercial zinc dust with a
dilute solution of acetic acid and heating on a water bath for
1 hour.
The zinc is filtered off by means of suction, washed with dis-
tilled water and dried.
Test for Nitrites. — Place 10 to 15 cubic centimeters of the
solution to be tested into a test tube and add 1 to 2 cubic centi-
meter portions of solution (a) first, then an equal amount of
solution (b). The formation of a red color in solution indicates
the presence of nitrites. This test is sensitive to 1 X 10~^ milli-
gram of N02~ ions in 1 liter of solution. If the solution, which
is to be tested for the presence of nitrite, contains free inorganic
acids, it must be first neutraHzed with a solution of a base. The
same is true of the following three methods.
Test for Nitrates, in the Absence of Nitrites. — This test is
based upon the reduction of nitrates to nitrites by zinc dust.
Place 10 to 15 cubic centimeters of the solution to be tested in a
test tube and add 1 to 2 cubic centimeter portions of reagents
(a) and (h) as in the test for nitrites. Add also a very small
pinch (about 10 to 20 milligrams) of zinc dust (e). The produc-
tion of a red color indicates the presence of nitrates. Sensitivity
of test, 1 milligram NOs" ions in 1 liter of solution.
Test for Hydroxylamine in the Absence of Nitrites. — This
test is based upon the oxidation of hydroxylamine (NH2OH) to
nitrites by iodine.
Place 10 to 15 cubic centimeters of the solution into a test
tube. Add 3 to 5 cubic centimeters of solution (a) and about 5
drops of solution (c). Shake and allow to stand 2 to 3 minutes
in the cold. Add solution (d), drop by drop, until the iodine
is just decolorized. Now add 2 to 3 cubic centimeters of solu-
tion (h). The formation of a red color indicates the presence of
hydroxylamine in the test solution.
Sensitivity of test, 3 X 10-^ milligram of NH2OH in 1 liter.
Test for Nitrates and Hydroxylamine in the Presence of
Nitrites.^ — The destruction of nitrites is first brought about by a
diazo-reaction, which consists in adding the sulfanihc acid reagent
(a) to the test solution and boiling.
1 Blom, J., Ber. deut. chem. Gesell, 59 : 121, 1926.
QUALITATIVE AND QUANTITATIVE ANALYSIS 59
Place 10 to 15 cubic centimeters of the test solution in a test
tube. Add 5 cubic centimeters of reagent (a). Boil over free
flame or heat on water bath for 4 to 5 minutes. To prove that
all nitrites have been destroyed, add to 3 cubic centimeters of
the mixture a few drops of reagents (a) and (b). The lack of
formation of a red color indicates the destruction of all the
nitrites.
The remaining solution containing the reagent (a) is cooled
down and used for the nitrate or hydroxylamine tests.
DiPHENYLAMINE ReAGENT^
1. Dissolve 0.7 gram of diphenylamine in a mixture of 60 cubic
centimeters of concentrated sulphuric acid and 28.8 cubic centi-
meters of distilled water.
2. Cool this mixture and add slowly 11.3 cubic centimeters of
concentrated hydrochloric acid (specific gravity 1.19). After
standing overnight some of the base separates, showing that the
reagent is saturated.
Test for Nitrates. — Place 1 drop of the substance to be tested
in a depression on the test plate. Add 1 drop of diphenylamine
solution and allow the solutions to mix thoroughly. Then add
2 drops of concentrated sulphuric acid. A deep-blue color
indicates nitrates. This test cannot be made in the presence of
nitrites, chloric and selenic acids, ferric chloride, and many other
oxidizing agents. Diphenylbenzidine is recommended as prefer-
able to diphenylamine. 2
Brucine Reagent
Dissolve 1.0 gram of brucine in 10 cubic centimeters of 50 per
cent pure sulphuric acid and make up to 100 cubic centimeters
with distilled water.
Test for Nitrates. — Place 1 drop of the substance to be tested
in a depression on the test plate and add 3 drops of concentrated
sulphuric acid. Now add 1 drop of brucine solution. If nitrates
are present, a red color develops quickly, which changes to orange,
1 Withers and Ray, /. Am. Chem. Soc, 33, 708-711, 1911.
2Snell, F. D., " Colorimetric Analysis," D. Van Nostrand Co., 1921.
60 LABORATORY MANUAL OF MICROBIOLOGY
then slowly to lemon or yellow, and finally becomes a greenish-
yellow.
This test can be used for quantitative determination of
nitrates. The test is based upon the final sulphur yellow color
rather than upon the initial red coloration. If the solution con-
tains much organic matter or ferrous iron, these should be
oxidized by a permanganate solution. The nitrous acid is
thereby also oxidized to nitric acid and if nitrous acid is deter-
mined separately, the results should be subtracted from those
of the nitric acid.
The determinations are made by adding 1 cubic centimeter of
the concentrated solution of brucine and 30 cubic centimeters of
concentrated sulphuric acid to 20 cubic centimeters of the solu-
tion of the unknown. In adding the sulphuric acid, care should
be taken to prevent boiling.
The nitrate is determined in a colorimeter, using for comparison
a standard solution containing 0.1872 gram KNO3 in 1 liter of
water. This standard gives 0.0001 gram N2O5 or 0.00002594
gram nitrate nitrogen per 1 cubic centimeter of solution. A
duplicate blank should always be made.
Phenoldisulphonic Acid^
Dissolve 25 grams of pure white phenol crystals in 150 cubic
centimeters of pure concentrated sulphuric acid, add 75 cubic
centimeters of fuming sulphuric acid (13 per cent SO3), stir well,
and heat for 2 hours at about 100°C. The reagent prepared in
this way should not contain any mono-acids or any tri-acids.
Two cubic centimeters of this reagent give reliable results with
not more than 5 milligrams of nitrate nitrogen.
Since in strongly acid solutions the following reaction takes
place, especially by heating,
3HNO2 = HNO3 + 2N0 + H2O,
a positive test for nitrate will be obtained, by the phenoldi-
sulfonic acid and brucine methods, in solutions containing only
nitrite.
' Chamot, Pratt, and Redfield, /. Am. Chem. Soc, 33, 381-384, 1911.
QUALITATIVE AND QUANTITATIVE ANALYSIS 61
Test for Indol :
Prepare test solutions :
(a) Paradimethylaminobenzaldehyde 1.0 gm.
Absolute alcohol 95 . 0 cc.
Hydrochloric acid (specific gravity 1.2) 20.0 cc.
(6) Potassium persulphate 1.0 gm.
Distilled water 100 . 0 cc.
Place a few drops of solutions (a) and (h) on a piece of absorbent
cotton. Push the cotton into the test tube until the moistened
surface is within about 1 inch of the culture liquid. Now place
the tube in boiling water for 5 to 10 minutes. If indol is present,
a red color will appear on the bottom of the cotton plug.
DETERMINATION OF MOISTURE IN SOIL
Weigh from 5 to 10 grams of soil into a glass or aluminum
dish and dry at 100 to 105°C. until there is no further change in
weight. About 6 to 12 hours are generally sufficient. Cool in a
desiccator and weigh. Determine all percentages of moisture on
the dry basis.
MOISTURE-HOLDING CAPACITY OF SOIL
To determine the amount of moisture which is required to
saturate the particular soil, the following procedure is used:
Round copper cups, about 4 to 5 centimeters in diameter and
about 2 centimeters high, with a perforated copper bottom are
used. Pieces of filter paper are cut to fit exactly upon the bottom
of the cups; the paper is moistened and the cups with the moist-
ened paper weighed. The cups are then filled with the soil in
question and the surface is leveled off carefully with the edge of
the cup. The cups are then weighed again. The cups with soil
are now placed in a dish with water, the water reaching out-
side of the cup to about half its height. After 24 hours the
soil has become saturated with water; the cups are removed and
the surface carefully dried with a cloth to remove water adhering
to outside of cup, and weighed again. The cups are then placed
in a drying oven for 24 hours, at 105°C., until they come to
constant weight. The soil is now carefully and completely
removed from the cup and from paper, and these weighed again,
giving weight of cup and dry paper. The moisture-holding
62 LABORATORY MANUAL OF MICROBIOLOGY
capacity of the soil as well as the moisture content of the par-
ticular soil can be calculated from these data.
QUANTITATIVE METHODS FOR AMMONIA DETERMINATION
1. Direct Distillation :
Sulphuric acid solution N/14
Sodium hydroxide solution N/14
Methyl-red.
Magnesium oxide.
1. Transfer the culture to be analyzed to an 800-cubic centi-
meter Kjeldahl or a copper flask, using about 200 cubic centi-
meters of distilled water.
2. Add 5 grams of magnesium oxide and some shavings of
paraffin to prevent foaming.
3. Connect to a condenser, the lower end of which is in iV/14
acid contained in an Erlenmeyer flask.
The period of distillation will vary with the amount of ammonia
present. As a rule, 1 hour is long enough to drive off all ammonia
nitrogen.
4. If methyl-red is used as an indicator, the distillate should be
boiled for a few minutes, cooled to 15 or 20°C., about 5 drops of
methyl-red added, and the solution titrated.
5. The distillate is titrated with standard alkali, and from the
number of cubic centimeters of standard acid neutralized by
the distillate the weight of nitrogen liberated as ammonia is
calculated.
2. Nesslerization. — Ammonia-free water should be prepared
by adding sodium hydroxide and potassium permanganate to
laboratory water and redistilling. Discard the first portion of
the distillate. After about one-fourth of the water has been
evaporated, the subsequent distillate will be free of ammonia.
Standard Ammonium Chloride Solution. — Dissolve 3.82 grams
of ammonium chloride in 1,000 cubic centimeters of distilled
water; dilute 10 cubic centimeters of this to 1,000 cubic centi-
meters with ammonia-free water. One cubic centimeter of this
solution contains 0.01 milligram of nitrogen.
1. Prepare a series of 16 Nessler's tubes which contain the
following number of cubic centimeters of the standard ammonium
chloride solution, and dilute to 50 cubic centimeters with
QUALITATIVE AND QUANTITATIVE ANALYSIS 63
ammonia-free water, namely: 0.0, 0.1, 0.3, 0.5, 0.7, 1.0, 1.4, 1.7,
2.0, 2.5, 3.0, 3.5, 4.0, 4.5, 5.0, and 6.0.
2. These will contain 0.01 milligram of ammonia nitrogen for
each cubic centimeter of the standard solution used.
3. Nesslerize the standards and also the distillates by adding
approximately 2 cubic centimeters of Nessler's reagent to each
tube.
4. Do not stir the contents of the tubes.
5. After Nesslerizing, allow the tubes to stand for 10 minutes.
6. Compare the color produced in these tubes with that in
the standards by looking vertically downward through them at
a white surface placed at an angle in front of a window, so as to
reflect the light upward.
3. Aeration Method.' — Place in a Kjeldahl flask 100 grams of
soil to be analyzed, 4 grams of sodium carbonate, 0.5 cubic
centimeter paraffin oil, and about 300 cubic centimeters of
ammonia-free water. Apply suction, bubbling the air first
through weak (10 per cent) sulphuric acid solution, to remove
any ammonia that might be present in the air; the air is bubbled
through the solution containing the soil and then is bubbled
through a bottle or long tube containing standard sulphuric acid
0.2N, O.IN, or 0.05N depending on the amount of ammonia
present), where the ammonia is absorbed. The flasks containing
the soil are placed in a water bath, the temperature of which is
maintained at 75 to 80°C. Aeration is continued for 2 to 3
hours. In the case of liquid cultures, 30 to 50 cubic centimeter
portions of the culture solution are placed in Folin aeration tubes,
5 cubic centimeters of 20 per cent sodium carbonate solution,
1 gram NaCl and 0.5 cubic centimeter petroleum oil are added
to each and aeration continued for 3 hours, tubes being placed in
thermostat at 60 to 65°C.
QUANTITATIVE (COLORIMETRIC) METHOD FOR DETERMINING
NITRATES
Evaporate to dryness in a porcelain dish on a water bath a
convenient quantity of unknown nitrate solution, depending
upon the amount of nitrate present.
1 GiBBS, W. M., et at, Soil Sci., 15 : 261-267, 1923.
64 LABORATORY MANUAL OF MICROBIOLOGY
This solution should be alkaline. In the case of soil extracts
which had been treated with CaO in filtration no further treat-
ment is necessary. In other cases, the solution should be treated
with NaOH or limewater to render slightly alkaline before
evaporation.
When evaporated, add 2 cubic centimeters of phenoldisulphonic
acid and stir with the rounded end of a glass rod for about 10
minutes so as to loosen the residue.
Note. — Equations for the action of phenoldisulphonic acid on a
nitrate :
H2SO4 + 2KNO3 = 2HNO3 + K,S04.
C6H3(OH)(S03H)2 + HNO3 = C6H2(OH)(S03H)2(N02) + H2O.
C6H2(OH)(S03H)2(N02) + 3NH4OH =
C6H2(ONH4)(S020NH4)2N02 + 3H2O.
Dilute with water and add ammonia solution (strong ammo-
nium hydroxide diluted with an equal volume of water) until
alkaline; a yellow color is formed. This is then diluted to a
known volume and compared with the standard.
Example. — Take 500 cubic centimeters of water to 100 grams of soil, and
in order to clarify add about 2 grams of calcium oxide. To secure a fair
sample, mix by rubbing in a mortar or by shaking in a wide-mouthed bottle.
Filter through folded filter paper until clear. Take a convenient volume,
for example, 25 cubic centimeters, and determine the nitrate present.
This is equal to 5 grams of soil. Use the colorimeter to compare the
standard solution with the unknown.
Formula for calculating results:
^ 100 *S , K ,,
^=W-A^'U-^
Where X = Number of milligrams of N as NO3 per 100 grams dry soil.
W = Weight of dry soil.
S = Cubic centimeters of water added to W.
A = Aliquot taken for evaporation.
d = Number of cubic centimeters to which A was diluted.
K = Reading on scale of standard solution.
U = Reading on scale of unknown solution.
M = Milligrams of N as NO3 in 1 cubic centimeter of standard
solution as diluted for reading.
Standard Nitrate Solution. — Dissolve 0.722 gram of pure dry
potassium nitrate in 1,000 cubic . centimeters of water. Of this
strong solution dilute 10 to 100 cubic centimeters, and from this
QUALITATIVE AND QUANTITATIVE ANALYSIS 65
take 10 cubic centimeters for a standard. Evaporate to dryness
in a porcelain dish on a water bath and treat as described on p.
64. Make up volume to 100 cubic centimeters. This standard
is equal to 0,1 milligram of nitrogen as nitrate.
Determination of Nitrates by the Reduction Method
Prepare an aqueous extract of the soil, peat or decomposing
organic matter which is to be analyzed. Extract 50 or 100
grams of the material three to six times with water and filter
through a folded paper, so as to collect 250 cubic centimeters of
solution. This solution is collected into a 500-cubic centimeter
Kjeldahl flask.
Add 5 cubic centimeters of a 50 per cent solution of sodium
hydroxide and distill over an open flame, until about 100 to 150
cubic centimeters have been driven over.
Replace the water driven off in heating.
When cool, add 2 grams of finely divided Devarda's alloy
(about 60-mesh) and connect flask again with distilling apparatus.
Boil carefully for 30 to 60 minutes and collect the distillate in
a definite volume of standard acid solution.
Titrate the solution with standard alkali and calculate the
nitrate-nitrogen distilled over as ammonia.
By this method one can determine both ammonia and nitrate
in soil, in peat, in manure, or in other organic composts. For
this purpose the original extract is made with 4 per cent KCl
solution instead of water. The first distillate will give the
ammonia present (collected in standard acid solution) and the
second distillate the nitrate.
QUANTITATIVE METHODS FOR DETERMINATION OF TOTAL
NITROGEN
Sulphuric acid A''/14
Sodium hydroxide ^ A^/14
Sulphuric acid (concentrated)
Potassium or sodium sulphate
Mercuric oxide or metallic mercury
Copper sulphate
Pumice powder or zinc powder free of nitrogen
Sodium hydroxide (40 per cent)
Place a 10-gram sample of soil in a dry Kjeldahl flask of 800-
cubic centimeters capacity.
66 LABORATORY MANUAL OF MICROBIOLOGY
Add 0.7 gram of mercuric oxide, 0.1 to 0.3 grams of crystalline
copper sulphate, and 10 grams of anhydrous sodium sulphate or
powdered potassium sulphate.
Add 30 to 35 cubic centimeters of concentrated sulphuric acid.
Thoroughly moisten the whole sample with acid before beginning
to heat.
Place the flask on the digestion shelf and heat slowly until
frothing ceases. Avoid a very high flame; do not aUow the flame
to touch the flask above the part occupied by the liquid.
Now raise the heat (avoid a very hot flame) until the acid boils
rapidly.
Digest for some hours after the liquid clears — until the cool
liquid is no longer yellow but blue-white in color.
In case the contents of the flask are likely to become solid
before digestion is complete, cool, and add 10 cubic centimeters
more of sulphuric acid.
When digestion is complete, cool, and add about 300 cubic
centimeters of water. Shake until the mixture is thoroughly in
solution. Be sure that none of the digested material remains
caked to the sides of the Kjeldahl flask.
Recool, add a teaspoonful of powdered pumice or a small
amount of zinc powder (0.5 gram) to prevent bumping.
Add 25 cubic centimeters of an 8 per cent sodium thiosulphate
solution in order to precipitate the mercury.
Add 80 to 100 cubic centimeters of the concentrated sodium
hydroxide. (The stock solution of alkali should be prepared 2
days or more before it is to be used in order that the sodium car-
bonate may precipitate out. Avoid the deposit in the bottom of
the alkali.) Enough alkali should be added to the acid digest to
make the solution react strongly alkaline. A few strips of litmus
paper may be added in order to test the reaction. The alkali
should be poured slowly down the sides of the flask, connected
at once to the condenser, and shaken.
See that the rubber stopper flts snugly in the flask. Now mix
the contents thoroughly by shaking.
Just before connecting the flask have a very low flame burning
on the distillation shelf. After the alkaU and acid mixture are
well mixed, raise the flame.
QUALITATIVE AND QUANTITATIVE ANALYSIS 67
The proper amount of standard acid should be measured
into flasks connected to the distillation shelf prior to adding
the alkali.
Distill slowly. After the first 15 minutes the flame may be
raised, but never so high that the distillate collects in the con-
densing bulbs. Generally the first two-thirds of the original
volume recovered as distillate will contain all the ammonia.
The distillate is now titrated with standard alkali, using
methyl-red or any other convenient indicator. From the cubic
centimeters of standard acid neutralized by the distillate the
weight of nitrogen liberated as ammonia is calculated.
Methyl-red Indicator. — Dissolve 1 gram of methyl red in 50
cubic centimeters of 95 per cent alcohol and dilute to 100 cubic
centimeters with water. Filter if the solution is turbid.
The percentage of nitrogen should be reported on the dry basis
of the soil.
A. Quantitative Methods for Determination of Total
Nitrogen Including Nitrates
Follow the same method as described under Quantitative
Methods for Total Nitrogen Determination, except for the use of
salicylic acid and sodium thiosulphate during digestion.
Add to the substance to be analyzed in a Kjeldahl flask, 10
cubic centimeters of sulphuric acid with salicylic acid (1 gram in
30 cubic centimeters of sulphuric acid) ; shake until thoroughly
mixed and allow to stand 5 or 10 minutes, with frequent shaking.
Now add 10 grams of sodium thiosulphate and heat the solu-
tion gently for 5 minutes, then bring to boiling for 5 minutes;
cool; add 0.5 gram of copper sulphate and mercuric oxide and
boil. This reduces the danger of foaming. Heat gently until
SO3 fumes begin to come off. Cool. Add 15 cubic centimeters of
the salicylic — sulphuric acid reagent, making in all 25 cubic centi-
meters of acid in each flask.
Heat very gently until foaming ceases, then heat strongly until
colorless. Continue boiling for 2 hours after substance is color-
less. The entire process requires 5 to 6 hours.
Follow the directions as given in Quantitative Methods for
Total Nitrogen Determination.
68
LABORATORY MANUAL OF MICROBIOLOGY
B. Total Nitrogen Including Nitrates in Aqueous Solu-
tions (Davisson-Parsons Method) 1
In addition to the usual apparatus for Kjeldahl analysis, pre-
pare absorbing towers as shown in Fig. 2. Place 35 cubic centi-
meters of sulphuric acid in each glass bead tower (concentrated
H2SO4 4 parts and water 1 part).
Take 100 cubic centimeters of the liquid to be analyzed, e.g.,
soil extract (if rich in nitrogen a suitable
aliquot may be used). Place the 100
cubic centimeters in an 800-cubic centi-
meter Kjeldahl flask and add sufficient
strong sodium hydroxide to make the
solution approximately 0.1 N NaOH.
Now add 4 drops of paraffin oil and
one gram of Devarda's alloy (60-mesh,
made free of ammonia by heating to
about 200°C. for 30 minutes) and im-
mediately connect to the glass bead
tower. Heat with a rose top burner
(high flame) until boiling, boil gently
for 20 minutes (during this time the acid
in the tower should about reach the
boiling temperature). Remove the
flame, allow the acid to suck back into
the flask, and again bring the solution
in Kjeldahl flask to the boiling tempera-
ture. Boil for a few minutes. Wash
the tower with small quantities of dis-
tilled water and allow the water to suck
back into the flask. About four wash-
ings of 25 cubic centimeters each will
be found sufficient to remove all of the
ammonia from the tower. Place on Kjeldahl digestion shelf and
heat with a low flame until the organic matter begins to char.
Now add 5 grams of potassium sulphate and continue digestion
1 Davisson, B. S. and J. T. Parsons, /. Ind. Eng. Chetn., 11: 306-311,
1919. Jacob, K. D. and W. J. Geldard, /. hid. Eng. Chem., 14: 1045-
1047, 1922.
S
Fig. 2. — Apparatus for
measuring total nitrogen in
the presence of nitrates.
QUALITATIVE AND QUANTITATIVE ANALYSIS 69
for about 2 hours after the liquid becomes clear. When com-
pleted, cool, add water, pumice stone, and strong alkali carrying
potassium or sodium sulfide (0.4 per cent) and distill as in the
official Kjeldahl method.
DETERMINATION OF AMINO NITROGEN
Amino acids and other amino compounds can be determined
gravimetrically or colorimetrically, either by the Van Slyke
method,^ which is based upon the interaction of nitrous acid with
the amiino group, giving gaseous nitrogen; by the Sorensen for-
malin titration method, ^ and by the Folin colorimetric method.^
QUANTITATIVE DETERMINATION OF CARBOHYDRATES
1. Free Celluloses. — True celluloses are characterized by the
fact that they are insoluble in dilute acids (2 to 4 per cent), but
they are soluble in concentrated acids, such as 42 per cent hydro-
chloric or 72 per cent sulphuric. The quantitative determination
of pure cellulose in soil or in culture is based upon its solu-
bility in ammoniacal copper solution, from which it is reprecipi-
tated by alcohol.
Preparation of Schweitzer^ s Reagent. — Two hundred grams of
copper sulphate are dissolved in hot water and precipitated with a
calculated amount of ammonia (95 cubic centimeters of ammonia,
specific gravity 0.90). The excess of ammonia is then neutral-
ized with sulphuric acid. The precipitate is washed by decanta-
tion in a large bottle three or four times and is then transferred
to a Biichner funnel and filtered through hardened filter paper
by the use of suction. With the aid of a porcelain spoon the
excess of water is pressed out from the copper hydroxide. It is
then removed in the form of a hardened paste from the filter
paper and introduced into a bottle containing ammonia water
and shaken in a shaking machine for 4 to 5 hours. An undissolved
part of the copper hydroxide should remain at the bottom of the
flask. The Schweitzer's reagent prepared in this way should
contain 1.5 grams of copper per 100 cubic centimeters of solution.
1 Van Slyke, D. D., J. Biol. Chem., 10: 15-55, 1911, 12: 275-284, 1912;
16: 539-547, 1913.
2 Sorensen, S. P. L., Biochem. Z., 7: 45, 1907-1908.
3 Folin, 0., J. Biol. Chem., 51: 377, 1922.
70 LABORATORY MANUAL OF MICROBIOLOGY
To test the strength of the reagent, 5 cubic centimeters are
placed in a crucible of constant weight, near a dish of H2SO4
under a bell jar. As soon as all the ammonia is absorbed, the
Cu(0H)2 is dried and heated to constant weight and weighed as
CuO.
Cellulose Determination.^ — Cellulose is added to the soil either in
the form of finely cut or well ground filter paper. After the soil
is properly mixed, a 20-gram sample is obtained from moist, or
preferably air-dried, soil. The sample is placed in a 250-cubic
centimeter sampling bottle, 100 cubic centimeters of Schweit-
zer's reagent is added; the bottle is then stoppered with a
rubber stopper and shaken for an hour in a shaking machine.
After settling, somewhat more than 50 cubic centimeters of the
liquid is filtered through a Gooch crucible by the use of suction.
Fifty cubic centimeters of the filtrate are then precipitated with
200 cubic centimeters of 80 per cent alcohol and the precipitate
is filtered through a Gooch crucible and washed as follows: (1)
dilute 1 per cent HCl, (2) warm distilled water, (3) dilute 2 per
cent KOH to get rid of humic acids — washing with KOH is
continued until all brown color disappears, (4) warm distilled
water, (5) dilute 1 per cent HCl to get rid of free alkali, (6) warm
distilled water until free from chlorides, (7) alcohol — after
cooling the crucible, (8) ether.
Dry to constant weight at 110°C., weigh, burn off and weigh
again. The difference between the two weights gives the
quantity of cellulose for 10 grams of the sample.
2. Celluloses in Plant Tissues. — Treat fresh or decomposed
material with 2 per cent solution of hydrochloric acid for 5 hours
under reflux condenser. Filter, and wash residue. Take aliquot
portion of residue and treat with 10 volumes of 80 per cent solu-
tion of H2SO4, then proceed as outlined on page 77. Calculate
cellulose content in original material.
3. Pentosans. — The determination of pentosans is based upon
their transformation into furfural when boiled with 12 per cent
hydrochloric acid : 2 grams of material are placed in a 500-cubic
centimeter flask provided with a separatory funnel and an
1 Charpentier, C. a. G. Thesis, Helsingfors, 1921; Waksman, S. A., and
O. Heukelekian, Soil Sci., 17: 275-292, 1924; Barthel, Chr., Abder-
halden's Handbuch, p. 754, 1927.
QUALITATIVE AND QUANTITATIVE ANALYSIS 71
outlet tube connected with condenser. One hundred cubic
centimeters of 12 per cent hydrochloric acid (specific gravity
1.06) is added from separatory funnel, and 30 cubic centimeters
distilled over in 10 minutes, the distillate passing through small
filter paper into receiver. As soon as 30 cubic centimeters of the
distillate are collected, 30 cubic centimeters more HCl is added to
flask and distillation continued. This is repeated until 270 cubic
centimeters of distillate are collected. Forty cubic centimeters of
filtered phloroglucide solution (11 grams of phloroglucinol
dissolved in 300 cubic centimeters of 12 per cent hydrochloric
acid) are added to the distillate and made up to 400 cubic centi-
meters with hydrochloric acid. After standing 16 hours, the
precipitate is filtered off using a weighed Gooch crucible with
thick asbestos mat. Wash with 150 cubic centimeters of water,
dry at 100 to 105°C. for 4 hours, and weigh. If a is the weight
of the precipitate, the pentosan content is, when less than 0.03
gram
(a - 0.0052) X 0.8949.
When the precipitate weighs 0.03 to 0.3 gram, the pentosan
content is
(a - 0.0052) X 0.8866.
4. Starches. — Starches are soluble in hot water and are readily
hydrolized by dilute acids and by diastatic enzymes giving the
reducing sugars maltose and glucose.
A definite amount of material (4 to 5 grams), previously
extracted with ether and alcohol, is treated with 50 cubic centi-
meters of water, in a double boiler and brought to the boiling
point; it is kept at that point for 15 minutes, until all the starch
has gelatinized. The mixture is then cooled to 50°C. and treated
with 10 cubic centimeters of malt extract. The temperature
(50°C.) is maintained for 30 to 60 minutes. The mixture is
again heated, cooled to 50°C., again treated with 10 cubic centi-
meters of the malt extract, and incubated for 30 to 60 minutes,
until no blue color is given with iodine solution. The mixture is
made up to 250 cubic centimeters and filtered through paper;
200 cubic centimeters of the filtrate is treated with 20 cubic
centimeters of hydrochloric acid solution (specific gravity
1.125) and heated, under a reflux condenser, for 2 hours.
72 LABORATORY MANUAL OF MICROBIOLOGY
The solution is cooled, neutralized with sodium hydroxide solu-
tion, and made up to 500 cubic centimeters. The solution is
filtered and reducing sugar (glucose) determined in an aliquot
portion. A blank determination of 20 cubic centimeters malt
extract boiled with acid as before outHned is subtracted. The
amount of glucose calculated is multiplied by 0.9 to give the
weight of starch.
5. Reducing Sugars. — Reducing sugars may be determined con-
veniently by the Bertrand or the modified Shaffer and Hart-
mann^ method or by any other convenient method.
Micro Method. — Prepare the following reagents:
(a) Combined micro reagent:
Grams per Liter
CuSOrSHzO 5.0
Tartaric acid 7.5
NazCOa (anhydrous) 40 . 0
KI 10.0
KIO3 0.7
Potassium oxalate 18.4
The sodium carbonate is dissolved in about 400 cubic centi-
meters of warm water, and into this, with stirring, is poured the
copper sulphate and tartaric acid dissolved in about 150 cubic
centimeters of water. The iodate, iodide, and oxalate are
dissolved in about 250 cubic centimeters of water, rinsed into
the alkaline copper solution, cooled and diluted to a liter. It is
usually more convenient to make up a larger quantity based on
the same proportion of reagents.
(5) lA^ H2SO4 — 27 cubic centimeters of concentrated H2SO4
are poured into a quantity of water and diluted to a liter. The
resulting solution is approximately normal but can be readily
adjusted by titration.
(c) O.IA^ thiosulphate — 25 grams of pure sodium thiosulphate
and 1 gram of NaOH are dissolved in water and diluted to a liter.
This should give a solution a trifle stronger than O.liV. It can
be readily standardized with a solution of O.IA^ K2Cr207.
To prepare this solution pure K2Cr207 is dried at 110°C., and
4.9033 grams are dissolved in water and diluted to 1 liter.
Twenty-five cubic centimeters of the standard dichromate are
1 Stiles, H. R., W. H. Peterson, and E. B. Fred, /. Bad., 12: 427-439,
1926.
QUALITATIVE AND QUANTITATIVE ANALYSIS 73
transferred to a large beaker containing about 3 grams KI and
10 cubic centimeters strong HCl in aqueous solution. The
contents are diluted to 500 or 600 cubic centimeters and titrated
with the thiosulphate. Starch paste is added toward the end
of the reaction, and at the end point the solution turns from a
blue to a light green. If the thiosulphate used is pure, the
volume will be a little less than 25 cubic centimeters, and the
solution can be readily adjusted to O.IN by dilution with water.
Thus if the titration is 24.7 cubic centimeters thiosulphate, every
24.7 cubic centimeters of thiosulphate solution should have added
to it 0.3 cubic centimeters water, or to 914 cubic centimeters of the
914
remaining solution add ^-^ X 0.3 or 11.1 cubic centimeters of
water. The resulting solution should be O.liV and can be
readily checked against the O.IA^ dichromate. The O.IA^ thio-
sulphate solution thus prepared will keep its strength for more
than a year. To make 0.005iV thiosulphate, 25 cubic centimeters
of the O.liV solution are diluted to 500 cubic centimeters in a
volumetric flask and mixed. This solution keeps for only a few
days and is best prepared anew for each set of determinations.
(d) Basic lead acetate, Home reagent — A 33 per cent solution
is used.
(e) Phosphate solution — For removing excess lead, a 10 per
cent solution of Na2HP04'12H20 is used. Three cubic centi-
meters are required for every cubic centimeter of the lead acetate
solution used. Add phenolphthalein and if alkaline or acid
neutralize.
Place 10 or 25 cubic centimeters of culture, depending on the
percentage of sugar, in a 50-cubic centimeter volumetric flask,
add a few drops of phenolphthalein and neutralize with sodium
hydroxide. Add 1 cubic centimeter of lead acetate solution,
shake, and then add 3 cubic centimeters of phosphate solution.
If alkaline or acid, neutralize, dilute to exactly 50 cubic centi-
meters and mix thoroughly by inverting. Let stand for 3
minutes and then remove an aliquot for analysis by means of a
pipette. If chlorides or other compounds precipitable by lead
are present, a little more of the lead acetate solution can be used.
An excess of lead acetate is to be avoided for some of the sugar
will be carried down with the precipitate.
74
LABORATORY MANUAL OF MICROBIOLOGY
Micro Sugar Table — Glucose Corresponding to Difference
IN Titration between Control and Sample
0.005 N
Thiosul-
phate,
Cubic
Centimeters
0.005 N
0.005 N
0.005 N
Glucose,
Milli-
gram
Thiosul-
phate,
Cubic
Centi-
meters
Glucose,
Milli-
grams
Thiosul-
phate.
Cubic
Centi-
meters
Glucose,
Milli-
grams
Thiosul-
phate,
Cubic
Centi-
meters
Glucose,
Milli-
grams
0.3
0.067
4.1
0.622
8.1
1.159
12.1
1.649
0.4
0.086
4.2
0.634
8.2
1.173
12.2
1.662
0.5
0.105
4.3
0.647
8.3
1.186
12.3
1.674
0.6
0.125
4.4
0.660
8.4
1.198
12.4
1.687
0.7
0.142
4.5
0.672
8.5
1.211
12.5
1.700
0.8
0.157
4.6
0.685
8.6
1.224
12.6
1.713
0.9
0.173
4.7
0.698
8.7
1.237
12.7
1.728
1.0
0.191
4.8
0.713
8.8
1.249
12.8
1.742
4.9
0.729
8.9
1.262
12.9
1.756
1.1
0.210
5.0
0.745
9.0
1.275
13.0
1.770
1.2
0.229
1.3
0.247
5.1
0.759
9.1
1.288
13.1
1.785
1.4
0.263
5.2
0.772
9.2
1.300
13.2
1.800
1.5
0.279
5.3
0.784
9.3
1.313
13.3
1.813
1.6
0.294
5.4
0.797
9.4
1.326
13.4
1.827
1.7
0.306
5.5
0.810
9.5
1.339
13.5
1.842
1.8
0.319
5.6
0.822
9.6
1.354
13.6
1.856
1.9
0.332
5.7
0.837
9.7
1.368
13.7
1.871
2.0
0.344
5.8
0.852
9.8
1.382
13.8
1.885
5.9
0.868
9.9
1.397
13.9
1.899
2.1
0.357
6.0
0.882
10.0
1.411
14.0
1.913
2.2
0.370
2.3
0.382
6.1
0.892
10.1
1.424
14.1
1.928
2.4
0.395
6.2
0.902
10.2
1.435
14.2
1.942
2.5
0.408
6.3
0.911
10.3
1.446
14.3
1.956
2.6
0.421
6.4
0.926
10.4
1.457
14.4
1.971
2.7
0.434
6.5
0.940
10.5
1.469
14.5
1.984
2.8
0.446
6.6
0.955
10.6
1.480
14.6
1.997
2.9
0.461
6.7
0.969
10.7
1.491
14.7
2.010
3.0
0.477
6.8
0.983
10.8
1.502
14.8
2.022
6.9
0.997
10.9
1.513
14.9
2.035
3.1
0.493
7.0
1.010
11.0
1.524
15.0
2.048
3.2
0.507
3.3
0.520
7.1
1.023
11.1
1.535
3.4
0.532
7.2
1.036
11.2
1.547
3.5
0.545
7.3
1.048
11.3
1.558
3.6
0.558
7.4
1.061
11.4
1.569
3.7
0.571
7.5
1.074
11.5
1.580
3.8
0.583
7.6
1.088
11.6
1.591
3.9
0.596
7.7
1.102
11.7
1.602
4.0
0.609
7.8
1.116
11.8
1.613
7.9
1.130
11.9
1.624
8.0
1.145
12.0
1.636
QUALITATIVE AND QUANTITATIVE ANALYSIS 75
Place 5 cubic centimeters of the micro reagent in a 50-cubic
centimeter Pyrex test tube and to this add from 1 to 5 cubic
centimeters, depending upon the quantity of sugar present, of
the clarified sample. If less than 5 cubic centimeters of sample is
taken, add sufficient water to make the total volume 10 cubic
centimeters. At the same time make up a blank with 5 cubic
centimeters of water and 5 cubic centimeters of reagent. Stopper
the test tubes with loose-fitting corks, to prevent oxidation from
the air, and heat for 15 minutes in a boiling water bath. Cool
in running water, add 5 cubic centimeters of IN H2SO4, shake
well, let stand 1 minute and titrate with O.OOSiV thiosulphate
and starch paste as an indicator. Add the starch solution when
the solution has turned a light straw color which indicates that
only a trace of iodine remains. Continue the titration with
thiosulphate until the blue color of the starch iodine compound
completely disappears. The end point is very sharp — within
1 to 2 drops of thiosulphate.
Calculation may be made directly from the micro-sugar table.
The titration of the sample in cubic centimeters of O.OOSA^
thiosulphate is subtracted from the titration of the blank,
the difference found in the table and read in milligrams of
glucose. The glucose per cubic centimeter of sample is then
calculated from the glucose found in the aliquot taken.
When 1 cubic centimeter of culture is taken for the direct deter-
mination, this method will accommodate samples containing up to
0.20 per cent glucose. If the sugar percentage is greater, the
sample must be diluted. For a sugar concentration of about 0.50
per cent, a convenient dilution would be 10 cubic centimeters made
up to 50 cubic centimeters and a 1 cubic centimeter aUquot
taken.
6. Lignins. — ^Lignins are prepared and determined by treating
natural or decomposed organic tnaterials with 42 per cent hy-
drochloric acid,^ 72 per cent sulphuric acid or a mixture of
hydrochloric and sulphuric acids; the acids decompose all the
celluloses, pentosans and proteins, leaving the lignins unattacked.
Four 2-gram portions of organic material, such as straw, ground
wood shavings, etc., are treated with ether to remove the fats
1 WiLLSTATTER, R., and L. Zeichmeister, Ber. deut. Chem. GeselL, 46:
2401, 1913. ScHWALLEE, H., Papi&rfabr., 23: 174-177, 1925.
76 LABORATORY MANUAL OF MICROBIOLOGY
and waxes. After the ether has been removed, the residue is
placed in a glass-stoppered flask or bottle and covered with 10
cubic centimeters of 18 per cent hydrochloric acid solution and
50 cubic centimeters of 72 per cent sulphuric acid solution.
The flask is stoppered, shaken, and immediately immersed in cold
water. The reacting mixture is allowed to stand for 2 to 3
hours, then transferred with 300 cubic centimeters of water to
500-cubic centimeter flasks and boiled for 30 minutes. The
residue is filtered off upon dried and weighed paper or Gooch
crucible, and washed with an excess of water. The residues
are then dried at 70°C. to constant weight. Two portions are
used for ashing and two for determination of total nitrogen by
the Kjeldahl method. (The weight of the dry residue) — (the
weight of ash + weight of nitrogen X 6.25) = amount of lignin.
COMPLETE ANALYSIS OF NATURAL OR DECOMPOSED PLANT
MATERIAL!
An approximately complete analysis of a natural organic
material is carried out as follows:
1. Moisture is determined on two 5-gram portions of material.
2. Total nitrogen is determined on two 2-gram portions of
material.
3. Ash on two 2-gram portions.
4. Pentosans on two 2-gram portions.
5. Two 5-gram portions are analyzed as follows:
(a) Treat for 12 hours with ether in Soxhlets. The ether
extract is evaporated to a small volume, then transferred to
weighing bottles and dried to constant weight, giving ether
soluble fraction.
(6) Residue from ether treatment is extracted for 24 hours
with cold water. Extract is made up to volume and divided into
four portions, one to be used for total nitrogen determination,
one for determination of total soluble organic matter by evaporat-
ing in silica dishes, one for determination of reducing sugar,
and one for the determination of ammonia or nitrate.
(c) Residue from cold water extraction is treated with 100 cubic
centimeters hot water, boiling to be continued for 30 to 60
minutes. The solution is analyzed as in (b).
1 Waksman, S. A., and F. G. Tenney, Soil Sci., 24: 317-341 (1927) 26:
113-137 (1928).
QUALITATIVE AND QUANTITATIVE ANALYSIS 77
(d) The residue from the hot water extraction is now treated
two or three times with boihng 95 per cent alcohol. The alco-
holic solution is evaporated in a weighing bottle and dried to
constant weight.
(e) The residue from the alcohol extraction is treated with 100
cubic centimeters of a 2 per cent hydrochloric acid solution and
boiled under a reflux condenser for 5 to 6 hours. The solution is
filtered off, through dried and weighed filter papers, and the
residue is washed with dilute acid, then with distilled water,
until free from acid. The filtrate and washings are now analyzed
for reducing sugars, by the Bertrand method, and for total
nitrogen. The amount of reducing sugar multiplied by 0.9 gives
the hemicellulose content of the material.
(/) The washed residue from the hydrochloric acid extraction
is dried to constant weight. Two 1-gram portions of the dry
material are placed in 300-cubic centimeter Erlenmeyer flasks
and treated with 10 cubic centimeters of an 80 per cent sulphuric
acid solution (if the residue is compacted, it should be first well
ground; if it is horny, it should have been washed with alcohol
and ether, before drying), for 2 hours, in the cold. The acid
must be brought in contact with all particles of the material.
After 2 hours, 150 cubic centimeters of distilled water is added to
each flask and contents autoclaved for 1 hour at 120°C., or
boiled for 2 to 3 hours under the reflux condenser. The con-
tents are then filtered through small dried and weighed filter
papers or through weighed Gooch crucibles. The residue is
well washed with water to wash out traces of sulphuric acid.
The combined solution and filtrate are analyzed for reducing
sugar. The amount of glucose found multiplied by 0.9 gives the
cellulose content of the material. Of the four residues for each
original material, two are used for ash and two for nitrogen
determinations. Weight of -(residue) — (ash + nitrogen X
6.25) = lignin content. The cellulose and lignin found in 1
gram of residue left from the 2 per cent HCl extraction are
now multiplied by the number of grams in this residue to
give the cellulose and lignin content in the original 5 grams
of material.
78 LABORATORY MANUAL OF MICROBIOLOGY
HUMUS DETERMINATION 1
Place six 50-gram samples of soil, previously well mixed and
sieved through a 1-millimeter sieve, into six 500-cubic centimeter
beakers. Add to each 50-cubic centimeters of 2.5 per cent NaOH
solution. Place beakers in autoclave and heat for 30 minutes
at 15 pounds pressure. Add 50-cubic centimeter portions of
cold distilled water to each beaker, and filter the dark-colored
solution through folded filter paper. After draining off all the
dark solution from the soil, add again fresh portions of 50-cubic
centimeters of 2.5 per cent NaOH solution to the same soils
previously extracted, and heat again as before. Add water and
filter upon the same papers. Wash soil with 50-cubic centimeter
portions of hot distilled water.
The combined filtrates are collected in 500-cubic centimeter
Erlenmeyer flasks and treated with a hot solution of hydrochloric
acid 1:1, until a heavy precipitate is formed; add half as much
more acid as was required to form the precipitate; warm and
shake well. Filter off precipitates (a-fraction of soil organic
matter or ''humic acid") through filter papers, previously
weighed and dried, or through Gooch crucibles. Wash with
warm 10 per cent hydrochloric acid, then a number of times with
distilled water. Dry the precipitates at 65 to 70°C. for 24 hours
and weigh. Three portions are used for ash determination and
three for total nitrogen determinations. If the precipitation
and washing with acid have been thorough, the ash content should
not exceed 1 to 2 per cent.
The filtrate from the acid precipitate is now treated with a 5
per cent solution of NaOH, until just neutral to litmus or until
solution has reached a pH of 4.8 to 5.0. A heavy precipitate
will be formed in case of mineral soils. The precipitate (j8-
fraction) is filtered off through a series of fresh papers, previously
dried and weighed, or through Gooch crucibles. These precipi-
tates are washed thoroughly with distilled water, dried and
weighed. Three portions are used again for ash and three for
nitrogen determinations. The nitrogen content of an aliquot
portion of the filtrate from the second precipitate is also
determined.
Tabulate results.
1 Waksman, S. a., Soil Sci., 22: 221-232, 1926; Springer, U., Zischr.
Pflanzenern, Dung. Bodenk., IIA: 313-359, 1928.
QUALITATIVE AND QUANTITATIVE ANALYSIS
79
CARBON DIOXIDE EVOLUTION
Evolution of carbon dioxide from soil, to which no fresh
quantities of undecomposed organic matter are added, is best
determined using 1-kilogram quantities of soil. Fresh, sieved
soil is placed in small unglazed porcelain pots holding just about
1 kilogram of soil. The pots are placed upon wooden bases and
covered with bell jars, which are sealed down to the stand with
r
D£
Fig. 3.-
-Apparatus for the study of the influence of plant growth upon the
evolution of CO 2 from soil. {Neller.)
paraffin. Air, freed from carbon dioxide by passing through soda-
lime and through bottles containing 10 per cent sulphuric acid, is
drawn through the bell jars, by means of glass tubes passed
through the wooden stands. The air is then drawn through
absorption towers containing 50 cubic centimeters of barium
hydroxide solution.
When the decomposition of fresh organic matter in soil is
studied, 100-gram quantities of soil may be used. Long-necked,
80 LABORATORY MANUAL OF MICROBIOLOGY
flat-bottomed flasks of 300-cubic centimeter capacity are used
for this purpose. The soil and the proper amount of organic
matter are introduced into the flasks; the moisture of the soil is
brought to the desired concentration and the flasks connected
with the respiration apparatus. The air, freed from CO2 as
before, is passed through the flasks over the surface of the soil.
The CO2 is absorbed in 100-cubic centimeter heavy-walled test-
tubes containing 25 to 50 cubic centimeters of the standard
barium hydroxide solution. The excess barium hydroxide is
then titrated back with standard oxalic acid solution (see also
Fig. 19, Exercise 58).
DETERMINATION OF TOTAL CARBON
Total carbon can be determined by (1) the various dry com-
bustion methods, including the bomb method (with Na202), and
(2) wet combustion methods, using a mixture of chromic and
sulphuric orchromic and phosphoric acids or permanganate and
sulphuric acid. The following method was found to give very
good results, especially with liquid cultures (see Fig. 4).
A definite portion of liquid culture or soil, containing not
more than 100 to 120 milligrams of carbon, is placed in a 200- to
500-cubic centimeter round-bottomed Pyrex flask {B of Fig. 4).
In case of liquid materials, the water should be evaporated on a
steam bath before beginning the analysis. The flask is then
attached to the condenser D, and 10 cubic centimeters of an
oxidizing solution (85 grams chromic anhydride in 100 cubic
centimeters of water made up to 250 cubic centimeters with 85
per cent phosphoric acid) is introduced through the separatory
funnel C. Gentle suction is then applied and 25 to 40 cubic
centimeters of a mixture of equal parts of concentrated phos-
phoric and sulphuric acids are added. The stopcock in C is closed
and by the use of a low flame, flask B is heated as rapidly as
possible without developing pressure within. The gas is drawn
out through condenser D and through a U-tube E, which con-
tains a saturated solution of Ag2S04 and 5 per cent H2SO4 in
20-mesh pumice on the left side 1, and boiled concentrated H2SO4
in 20-mesh pumice on the right side 2. The Ag2S04 serves to
remove the chlorine and the H2SO4 the SO3 fumes. The gas is
then drawn through the absorption flask F and absorbed in a
measured quantity of O.bN NaOH solution. After flask B has
QUALITATIVE AND QUANTITATIVE ANALYSIS
81
been boiled gently for 15 to 20 minutes, flask F is removed, the
solution washed down into the flask, the carbon precipitated
by the addition of an excess of 2 iV neutral BaCU solution,
and the excess alkah titrated with 0.5A^ HCl solution using
phenolphthalein as an indicator.
Fig. 4.' — Apparatus for determination of total carbon in soil or in solution.
The absorption flask F is made of a 300-cubic centimeter
Erlenmeyer flask, a 1.5 X 25 cm. test tube, and a few glass beads.
The bottom of the test tube is drawn out and while still hot the
upper part of the constriction is flattened so that two beads will
lie over the opening instead of one, so as to facilitate washing out.
82 LABORATORY MANUAL OF MICROBIOLOGY
Suction Flask. — The suction flask H is provided with a constant
pressure valve / which is made from small glass T-tube in which
a rubber band holds a small rubber disk over the lower end. By
the aid of this valve and the screw cock G, the flow of air can be
easily regulated and should not exceed about 120 bubbles per
minute.
1 cubic centimeter 0.5A^ NaOH =c= 3.0 milligrams of carbon.
Cubic centimeters NaOH added— cubic centimeters HCl used X
3 = milligrams carbon.
SEED STERILIZATION
Although a great number of methods employing various agents
have been recommended for removing microorganisms from seed,
only a few of the more promising ones will be given. Where it is
not necessary to render the seeds free of bacteria, but merely to
destroy the majority of the flora, alcohol may be used.
Among the chemicals that have proved satisfactory for steriliz-
ing seed, mercuric chloride, hypochlorite of lime, and silver
nitrate are the most commonly used. The effectiveness of these
substances depends on many factors: strength of solution, time
of exposure, temperature, pressure, and nature of the seed coat.
Sterilization hy Mercuric Chloride in Vacuum. — Select a large
heavy walled desiccator and connect to vacuum pump.
Fill test tubes or flasks about one-half full of seed, cover with
0.25 per cent solution of mercuric chloride and place in the
desiccator.
Exhaust for 3 to 5 minutes depending upon the kind of seed.
This should remove the air particles from around the seed coats
and allow the disinfectant to come in direct contact with the
seed.
At the end of this time remove the mercuric chloride solution
and run in a small amount of sterile water, shake vigorously,
empty, and repeat this process three or four times.
Remove some of the seed to sterile Petri dishes and pour over
them a layer of nutrient agar.
After the agar hardens, invert and place in the incubator at
20 to 25°C. In 2 or 3 days the seed should germinate. If
bacteria or molds are present, they may be readily noted on
the agar.
QUALITATIVE AND QUANTITATIVE ANALYSIS 83
Sterilization by Calcium Hypochlorite.^ — 1. Add 10 grams of
commercial chloride of lime (titrating 28 per cent chlorine) to
140 cubic centimeters of water.
2. Allow the mixture to settle for 5 or 10 minutes and decant
the supernatant liquid. This solution should contain about 2 per
cent of chlorine.
3. For seed sterilization the solution may be diluted or used
full strength. The volume of the liquid should be about five
times that of the seed.
4. Place the seed in a sterile test tube and cover with a 1 per
cent chlorine solution (original solution diluted one-half).
5. The time required for sterilizing varies with the different
seeds, about 6 hours for alfalfa, 8 hours for corn, and 15 hours for
wheat.
1 Wilson, J. K, Am. J. BoL, 2: 420-427. 1915.
PART IV
EXERCISES
THE STUDY OF MICROORGANISMS IN THE SOIL
A. Suggested Arrangement of Class Exercises in Soil Micro-
biology for Course of Five Credits for one Semester
of 18 Weeks
Because of the time required for the incubation of the different
groups of microorganisms, it is suggested that the exercises be
given in the order listed below.
Introduction
General Characteristics of the Soil Population:
1. Bacteria.
2. Fungi.
3. Algse.
4. Protozoa.
5. Invertebrate population of soil, non-protozoan in nature.
6. Approximate number of microorganisms in soil.
A. Microscopic examination of microorganisms:
1. Examination of living microorganisms in hanging drops.
2. Examination of bacteria in Congo red or Nigrosin preparations; of
yeast and yeast spores in Erythrosin or Rose Bengal preparations.
B. Methods for counting numbers of microorganisms :
3. Number of algae in soil according to dilution method.
4. Number of protozoa according to the dilution method.
5. Number of fungi according to the plate method.
6. Number of bacteria according to the plate method (aerobic).
7. Number of anaerobic bacteria in the soil.
8. Number of spore-forming bacteria in the soil.
9. Number of thermophilic bacteria.
10. Effect of season of the year on number of microorganisms.
11. Effect of plant roots on number Of microorganisms.
12. Effect of depth of soil on number of microorganisms.
13. Effect of manures on number of microorganisms.
14. Dilution method for determining the number of specific physiological
groups of bacteria.
15. Direct microscopic examination of soil.
16. Winogradsky's method of microscopic analysis of soil.
17. Determination of numbers of nematodes (and other worms) and
insects in soil.
87
88 LABORATORY MANUAL OF MICROBIOLOGY
C. Nitrogen-fixing bacteria and nitrogen fixation in soil :
18. The isolation of bacteria from the root nodules of various leguminous
plants.
19. Cultural characteristics of root-nodule bacteria.
20. The formation of root nodules in agar and sand cultures.
21. Artificial cultures for the inoculation of leguminous plants.
22. The structure of root nodules of leguminous plants.
23. Effect of root-nodule bacteria on the growth and the nitrogen content
of alfalfa.
24. Direct method for demonstrating the occurrence of Azotobacter in
soil.
25. Isolation of Azotobacter from various soils.
26. Use of silica-gel plate for the isolation of specific bacteria.
27. Nitrogen fixation by pure cultures of Azotobacter.
28. Effect of variation in carbohydrates on the growth of Azotobacter.
29. Anaerobic nitrogen fixation (Clostridium pasteurianum and related
organisms). Spore stain- Dorner modified).
30. Isolation of anaerobic nitrogen-fixing organisms.
31. Nitrogen-fixing capacity of soil (Winogradsky).
32. Influence of nitrate on the fixation of nitrogen.
33. Effect of a soluble carbohydrate on nitrogen assimilation.
D. Denitrifying bacteria:
34. Isolation of denitrifying bacteria.
35. Denitrification by pure cultures of bacteria.
36. Denitrification in soil.
E. Nitrification experiments:
37. Nitrification in impure cultures.
38. Nitrification in liquid cultures (quantitative).
39. Isolation of nitrifying organisms.
40. Nitrification of various substances.
F. Urea and protein decomposition:
41. The decomposition of urea with the production of ammonia.
42. Isolation of urea-decomposing organisms.
43. Ammonia production from various substances in soil.
44. Decomposition of an amino acid and a protein by Bac. cereus and
Bad. fluorescens.
G. Sulphate-reducing and sulphur-oxidizing bacteria.
45. Reduction of sulphates with the formation of hydrogen sulphide.
46. Isolation of hydrogen sulphide-forming microorganisms.
47. Crude cultures of higher sulphur bacteria.
48. Oxidation of sulphur and the dissolving of rock phosphate.
49. Isolation of pure cultures of higher sulphur bacteria.
50. Growth and isolation of Thiobacillus thioparus.
51. Growth of Thiobacillus thiooxidans in liquid medium.
H. Iron bacteria:
52. Iron-precipitating bacteria.
63. Iron bacteria from drinking water.
THE STUDY OF MICROORGANISMS IN THE SOIL 89
/. Cellulose-decomposing bacteria:
54. Anaerobic cellulose decomposition in impure cultures (liquid).
55. Number of aerobic cellulose-decomposing bacteria in soil.
56. The thermophilic fermentation of cellulose.
57. Isolation of cellulose-decomposing bacteria.
58. The evolution of carbon dioxide from soil.
A SUGGESTED LIST OF APPARATUS FOR ONE STUDENT
The following apparatus should be in each desk. Any omission must be
reported to the instructor at once.
1 Bunsen burner and tubing
4 Wire baskets
2 Metal cups
2 Funnels
5 Erlenmeyer flasks (150 cubic centimeters)
5 Glass tumblers
10 Pipettes (1 cubic centimeter)
2 Pipettes (10 cubic centimeters)
1 Graduated pipette (5 cubic centimeters)
1 Thermometer
2 Platinum needles
20 Object slides (not returnable)
1 Hanging-drop slide
50 Cover glasses (not returnable)
1 Aluminum weighing dish
6 Evaporating dishes
1 Test plate
1 Wash bottle
Filter paper (8-inch)
1 Forceps (steel)
1 Spatula
1 SHde box
1 Test-tube brush
1 Towel
1 Box of matches
1 Box of labels
1 Wax pencil
1 Microscope No
(No. ...^.
Objectives -j No
I No
1 Microscope lamp
LABORATORY RULES
Read carefully the following rules:
Before pouring plates or making transfers, wash off the desk
with a 1 : 1,000 mercuric chloride solution.
90 LABORATORY MANUAL OF MICROBIOLOGY
Mercuric Chloride. — A stock solution is prepared and diluted
to the desired strength.
Add 1 part of mercuric chloride to 2.5 parts of commercial
hydrochloric acid (40 per cent HgCU in HCl). To prepare
a 1:1,000 solution, take 2.5 cubic centimeters of the stock solu-
tion and dilute to 1,000 cubic centimeters. Color with dye and
label ''poison."
Transferring Cultures. — Hold the test-tube cultures to be
transferred as nearly in a horizontal position as possible. Avoid
opening cultures in a current of air.
All cultures are to be grown in the incubator at 28°C. unless
otherwise stated.
Care of Apparatus. — After using balances, always return
weights to their proper places. Do not leave any dust or dirt
on balances.
All solid material, as soil, agar, cotton or filter paper, must be
emptied into waste jars and not into the sinks.
Soil should not be sieved in the laboratory. The greenhouse
or potting room may be used for this purpose.
At the end of the laboratory period return all stock bottles
and chemicals to their proper places on the shelves. See that
all apparatus is replaced in the lockers and that all gas burners
are shut off. Wipe off the table top before leaving.
Anything left on the desk will be collected after the laboratory
period and returned to the store room.
Cleaning Glassware. — All glassware must be thoroughly cleaned
before it is ready to use. Test tubes, Petri dishes, flasks, and
similar glassware should be boiled in a 5-per cent soda solution or
washed in hot soapsuds until free from organic matter. When it
is desirable to use very clean glassware, immerse for 10 minutes
or longer if possible in the dichromate solution.
Potassium (K2Cr207) or sodium dichromate (Na2Cr2-
Ot) 40 gm.
Water 150 cc.
Sulphuric acid (H2SO4) 230 cc.
Note. — Dissolve the dichromate in warm water and, when cool, add
slowly concentrated sulphuric acid. If properly prepared, the liquid should
be thick, with small crystals. It may be used repeatedly, provided the
crystals are present.
THE STUDY OF MICROORGANISMS IN THE SOIL 91
After removing glassware from the cleaning solution rinse
thoroughly in distilled water.
Dirty cover glasses and slides may be treated in the same
manner. Drop these, one at a time, into the dichromate mixture
and allow to remain for several hours. Remove from this solu-
tion, wash, and wipe with a soft, clean cloth.
A simple and more rapid method, suitable for general work,
is to rub the shdes with moist Bon Ami, and when dry pohsh
them with a clean cloth.
In order to remove fat, pass the cover slips through a flame.
Where it is desirable to have very clean slides and cover slips,
it is well to heat them in water and then in 50 per cent sulphuric
acid. After rinsing in distilled water, wash in alcohol and wipe
with a clean cloth. These should be kept in a clean, covered dish.
A Black Finish for Table Tops
Method 'I
The following solutions are required:
A. Copper sulphate 125 gm.
Potassium chlorate 125 gm.
Water 1,000 cc.
Boil until salts are dissolved.
B. Aniline hydrochloride 150 gm.
Water 1,000 cc.
Or, if more readily procurable:
Aniline oil 120 gm.
Hydrochloric acid 180 gm.
Water 1,000 cc.
By means of a brush apply two coats of solution A while hot,
the second coat as soon as the first is dry. Then apply two
coats of solution B and allow the wood to thoroughly dry. A
coat of raw linseed oil is next apphed. It is best to use a cloth
instead of a brush so as to get only a very thin coat of oil. The
desired amount of polish is now given the wood by rubbing in
the oil. In the treatment with the oil the deep black color is
partially brought out, although this does not uniformly appear
until the table has been thoroughly washed with hot soap suds.
This takes out the superfluous chemicals.
The finish thus secured is an ebony black which is permanent
and very highly resistant to the action of chemicals, such as
acids and alkahes, even concentrated sulphuric acid having httle
or no effect if quickly washed off.
92 LABORATORY MANUAL OF MICROBIOLOGY
Method II
The following solutions are required:
A. Aniline 120 gm.
Hydrochloric acid (commercial) 180 gm.
Water 1,000 cc.
B. Sodium dichromate 120 gm.
Hydrochloric acid 100 gm.
Water 1,000 cc.
Solution A should be applied with a brush to the fresh smooth
surface and allowed to dry overnight. The color will turn
bright yellow. Solution B should then be spread on the wood,
which will turn dark and be very streaky at first. After this
second coat dries the surface should be rubbed with vaseline,
motor oil, or paraffin. Vaseline seems to be preferable.
GENERAL CHARACTERISTICS OF THE SOIL POPULATION
Every particle of soil harbors numerous microorganisms, which
vary in kind, nature of nutrition and, therefore in the biochemical
processes which they bring about. Although one set of condi-
tions may favor the activities of one group of organisms in prefer-
ence to others, no soil is known in which only a single species of one
organism is found. The actual number of cells may vary from a
few hundred thousands per gram in certain very poor sandy soils
and in bog soil to many billions in good garden, field, and green-
house soils. The number of species varies from a few types of
fungi and bacteria in marshy lands or in very acid forest soils to
thousands of species in the garden and field soils.
The soil population can be readily divided into the following 5
groups :
1. Bacteria. — This group is usually the most numerous in the
soil, both in the total number of cells and in the number of species.
Morphologically, they comprise the cocci, non-spore forming
and spore-forming bacteria and spirilla. Some bacteria, forming
thin, long, flexuous, undulating cells are often referred to as Spiro-
chsetes, but these soil forms belong to the bacteria rather than
to the protozoa. These numerous soil bacteria vary consider-
ably in their nutrition and in their response to environmen-
tal conditions. We have aerobic and anaerobic forms, auto-
trophic and heterotrophic bacteria, those that are resistant to
acids and those that do not grow at a pH less than 6.0. The
THE STUDY CF MICROORGANISMS IN THE SOIL 93
nature and abundance of the various types of bacteria thus
depends both upon the available nutrients present in the soil and
upon the soil environmental conditions, such as reaction, aeration,
moisture content, abundance of organic matter, etc. The plate
method of counting gives only a fraction of the total number of
bacteria present in a given soil. This is due to the fact that
the many types of bacteria such as the autotrophic, anaerobic,
nitrogen-fixing ones, etc., do not develop on the common media.
2. Fungi. — Although the bacteria from the largest group of
organisms in most soils, the fungi may be present in considerably
greater bulk in some soils, such as acid forest soils or heavily
manured, acid garden and field soils. This is due to the consid-
erable difference in size of a piece of fungus mycelium and a
bacterial cell. The abundance of fungi in the soil can be deter-
mined both by the plate and the microscopic methods. A colony
may develop either from a spore or from a piece of mycelium.
Spore fungi, like most Hyphomycetes, form a mycelium which
readily breaks up into fragments. Others, like the Phycomycetes
form a unicellular mycelium. In the case of these organisms, the
number of colonies developing on a plate may be far from repre-
senting their actual abundance. The fungi are represented in the
soil by (a) the filamentous fungi or molds, (b) the actinomyces, or
ray fungi, and (c) the yeasts. In acid forest soils, there is an
extensive development of fungus mycelium, belonging to Basidio-
mycetes and other fungi, which do not readily form any fruiting
bodies. Many of these produce '^mycorrhiza" with the roots of
various trees, and a number of other perennial and annual plants.
The relative abundance of these fungi can be obtained only by the
aid of the microscope.
3. Algae. — The chlorophyll-bearing microscopic plants do not
need any carbon compounds as sources of energy. They obtain
from the soil only nitrogen and minerals while the carbon is
derived from the CO2 of the atmosphere, using photosynthetic
energy. The algae are represented in the soil by the (a) Cyano-
phycese, or blue-greens, (h) Chlorophycese, or grass-greens, (c) and
Diatomaceae, or diatoms. Many of the algae are able to live at
lower depths of soil and are capable of obtaining their carbon
and energy from complex organic compounds. The number
of algae in the soil is best determined by the dilution method.
94
LABORATORY MANUAL OF MICROBIOLOGY
4. Protozoa. — Many of these organisms feed upon bacteria,
and their development depends upon the abundance of bacteria.
Some of the protozoa, hke the ciUates and amoebae, are large in
size, and they may form a considerable quantity of living or
recently living protoplasm in the soil. The protozoa are repre-
sented in the soil by the (a) flagellates, (b) amoebde, and (c) ciliates
which are usually less numerous than the other two groups.
5. Invertebrate Population of Soil, Non-protozoan in Nature. —
This group is represented in the soil by the (a) Nematodes, or
round worms, which may be present in hundreds of millions per
acre of soil, (b) Rotifers or wheel animalcules, (c) Turbellarians,
or flat worms, (d) Trematodes, or flukes, (e) AnneUds or seg-
mented worms, including the earthworms, (/) Arachnids, includ-
ing the mites, ticks, and spiders, (g) Myriapodes, (h) insects, etc.
The relative abundance of these organisms per gram of soil
can be shown as follows :
Approximate
Number of
Microorganisms in Soil
Numbers,
plate
method in
1 gram
Numbers,
microscopic
method in 1
gram
Approximate size,
in microns
Bacteria :
Minimum
100,000
100,000,000
1,000
20,000,000
0
40,000,000
a few
500,000
10,000
2,000,000
500,000,000
10,000,000
10,000,000,000
a few
50,000,000
0.3-2X0. 4-10
Maximum
Fungi :
Minimum
3-10 X 3-100
Maximum
Actinomyces :
A/Tinirniim
0.5-2.0X0. 5-50
Maximum
AlgjE:
Minimum
1.5 X 2-50?
Maximum
Protozoa :
A/Tini mnm
2-10 X 5-200
Maximum
Invertebrates (non-pro-
tozoan) per acre, up to . .
100 to 2,000
THE STUDY OF MICROORGANISMS IN THE SOIL 95
The microorganisms are responsible for the various biochemical
processes going on constantly in the soil and which lead to the
liberation of nutrients essential for the growth of higher plants.
These nutrients include the carbon, as carbon dioxide, the
nitrogen as ammonia and nitrate, the minerals, especially
phosphorus and potassium. The following exercises are planned
to show how the microorganisms liberate those nutrients, as well
as the nature of the organisms themselves and methods of
study.
MICROSCOPIC EXAMINATIONS OF MICROORGANISMS
Exercise 1
Examination of Living Microorganisms in Hanging Drops
Place a small drop of the liquid to be examined in the center
of a cover glass without spreading. Smear vaseline around
the concavity of a hollow-ground slide and invert this slide over
the cover glass with the drop so that the cover glass adheres
to the slide. Turn over the sHde and see that the drop does not
touch the sHde. Examine the hanging drop with a low- and then
with a high-power objective. To secure the best results reduce
the illumination. The addition of a small amount of nigrosin
(saturated aqueous solution of nigrosin B) to the hanging drop
makes the bacteria stand out as clear spots in a dark background.
It is advisable to focus on the edge of this drop.
While this method of examination is primarily for detecting
motihty and Brownian movement, it is useful in a study of general
morphology.
Examine: 1. Hay infusion.
2. Pure culture of yeasts and bacteria.
Exercise 2
A. Examination of Bacteria in Congo Red or Nigrosin
Preparations
This method of examination as described below has many
advantages; it is rapid and does not alter the size of the organisms.
It is especially useful for liquid cultures. The bacteria appear as
clear spots in a dark background. If Congo red is used the dead
cells appear as faintly colored, the living as colorless. Because
96 LABORATORY MANUAL OF MICROBIOLOGY
of contamination which may be present in the dye solutions it is
always well to mount a drop of the uninoculated dye on the same
slide.
Examine: Cultures of bacteria from hay infusion, B. suhtilis
or B. mesentericiis.
1. Congo Red for Negative Mounts :
(For differentiating living and dead bacteria.)
(a) Place a drop of 2-per cent aqueous Congo red solution (free
of bacteria) on a clean glass slide.
(6) Mix with it a loopful of the bacterial culture.
(c) Allow it to dry thoroughly in air 10 minutes or more.
(d) Flood with acid-alcohol (1- or 2-per cent HCl). This
changes the color to blue and fixes the film.
(e) Dry without washing and examine in oil, with or without
cover glass. Living cells appear unstained- white against blue.
Unless preserved with oil or balsam the preparations fade rapidly.
This method of preparing negative mounts is recommended for
root-nodule bacteria. The active living cells are negative while
the dead cells are more or less positive.
2. Nigrosin for Negative Mounts :
(a) Place a loopful of culture on a clean glass slide, spread, and
allow it to air dry.
(b) Spread thinly over the smear a loopful of saturated aqueous
solution of nigrosin B. Spread either with the wire loop or with
a glass slide (as for blood smears). Dry and examine in oil, or
mount in Canada balsam.
By the use of nigrosin it is possible to examine organisms
unstained. There are many points in favor of this method,
e.g., the organisms do not shrink or change their form.
B. Examination of Yeasts and Yeast Spores in Erythrosin or Rose
Bengal Preparations
In a drop of sterile water on a glass slide, mix a small amount of
the yeast culture.
Add a large loop of Erythrosin or Rose Bengal (0.5 gram in 100
cubic centimeters of water). The dead cells stain a deep pink
while the living cells remain colorless.
To prevent evaporation make a ring of vaseline around the
edge of the drop culture and place cover slip over the top.
THE STUDY OF MICROORGANISMS IN THE SOIL 97
Examine under high-dry and oil-immersion lenses. Make
drawings.
METHODS FOR COUNTING NUMBERS OF MICROORGANISMS
Directions for Drawing Soil Samples. — Samples from the sur-
face to 1 foot deep may be taken as follows: Remove the coarse
surface debris and sink a metal cylinder or soil sampler to the
desired depth. Samples of surface soil may be taken with a
sterile spatula. Draw several samples and empty into sterilized
paper bags or other vessels. Mix and pulverize the sample. This
may be done with a sterile spatula upon a large piece of sterile
paper. From the well-mixed sample remove a representative
portion for dilution, and at the same time make a moisture
determination.
When it is necessary to secure accurate samples from various
depths, it is well to dig a ditch to the desired depth. By means
of a sterile trowel, representative samples may be drawn from
the sides of the ditch. In this way outside contamination is
largely prevented.
In order to reduce the error common to determinations of this
character, it is well to use a large sample of soil. Balances
sensitive to 10 milligrams are satisfactory for this work.
Exercise 3
NUMBEK OF AlG^ IN SoiL ACCORDING TO DILUTION METHOD
Add 50 grams of soil to 500 cubic centimeters of sterile
water and shake vigorously for 5 minutes.
After the coarse particles have settled, dilute as follows:
(a) Add 10 cubic centimeters of soil suspension to 90 cubic
centimeters of sterile water, giving a dilution of 1 : 100.
(5) Add 10 cubic centimeters of dilution (a) to 90 cubic centi-
meters of sterile water, giving a dilution of 1:1,000. If the soil
is rich it is well to increase the dilutions.
Inoculate one flask of algal medium (Medium 42 and 43) with
1 cubic centimeter of each of the above dilutions.
The cultures should be kept near a window. Note when green
color appears, usually 30 to 90 days. Examine in wet mounts.
Calculate the number of algae in 1 gram of soil.
98 LABORATORY MANUAL OF MICROBIOLOGY
Exercise 4
Number of Protozoa according to the Dilution Method
Add 50 grams of surface soil to 500 cubic centimeters of sterile
water, as directed in the preceding exercise.
After the coarse particles have settled, dilute as follows:
(a) Add 10 cubic centimeters of the soil suspension to 90 cubic
centimeters of sterile water; dilution 1:100.
(6) Add 10 cubic centimeters of dilution (a) to 90 cubic centi-
meters of sterile water; dilution 1:1,000.
(c) Add 10 cubic centimeters of dilution (b) to 90 cubic centi-
meters of sterile water; dilution 1:10,000.
(d) Add 10 cubic centimeters of dilution (c) to 90 cubic centi-
meters of sterile water; dilution 1:100,000.
Inoculate duplicate tubes containing liquid media, using hay-
extract (Medium 38), soil extract (Medium 40), or any medium
adapted to protozoa. Nutrient agar placed in dishes may also
be used.
Incubate the protozoan cultures at room temperature.
At intervals of 2 days each, make a microscopic examination
of the cultures. Since the protozoa are usually larger than bac-
teria— the 16 millimeters, or two-thirds, and 4 millimeters, or
one-sixth objectives will be found desirable.
By means of a large-mouthed pipette or loop, transfer a small
portion of the protozoan culture to a slide and examine. A wet
or hanging-drop mount may be used. In certain cases the small
flagellates become so numerous that it is difficult to distinguish
between the bacteria and protozoa.
If it is desired to differentiate between the number of vege-
tative cells of protozoa and number of cysts, another 50 grams of
soil maybe treated with 200 cubic centimeters of a 2 per cent solu-
tion of hydrochloric acid and allowed to stand over night. The
next morning 300 cubic centimeters of water are added and dilu-
tions made as before. The treatment with acid results in the
destruction of the vegetative cells and thus the number of
protozoan cysts may be obtained.
Exercise 5
Number of Fungi according to the Plate Method
Add 50 grams of soil to 500 cubic centimeters of sterile water.
Shake vigorously for 5 minutes.
THE STUDY OF MICROORGANISMS IN THE SOIL 99
After the coarse particles have settled, dilute as follows:
(a) Add 10 cubic centimeters of soil suspension to 90 cubic
centimeters of sterile water; dilution 1:100.
(6) Add 10 cubic centimeters of dilution (a) to 90 cubic centi-
meters of sterile water; dilution 1:1,000.
(c) Add 10 cubic centimeters of dilution (b) to 90 cubic centi-
meters of sterile water; dilution 1:10,000.
{d) Add 10 cubic centimeters of dilution (c) to 40 cubic centi-
meters of water; dilution 1:50,000.
From dilutions (6), (c), and (d), of 1:1,000, 1:10,000 and
1:50,000, pour plates in triplicate.
Add to each plate about 10 cubic centimeters of the peptone-
sucrose-agar, Medium 18, melted and cooled to 40°C.
Incubate at 28°C. and count after 2 and 4 days.
Exercise 6
Number of Bacteria according to the Plate Method (.erobic)
Weigh 50 to 100 grams of soil on a piece of paper or scoop.
Transfer the soil to a 500- or 1,000-cubic centimeters sterile
water blank.
Five hundred cubic centimeters of water in a 750-cubic centi-
meter Erlenmeyer flask allows ample space for shaking (tap.
water may be used). For ordinary work, provided the blanks are
not stored for a long time, sterilization of the water blanks for
30 minutes in the steamer will be sufficient for use in soil counts.
Some prefer sterilization in the autoclave for 15 minutes at 15
pounds' pressure.
Shake the soil suspension vigorously for at least 5 minutes and
allow the coarse particles to settle.
Add 10 cubic centimeters of this first dilution (equivalent to 1
gram of soil) to a 90-cubic centimeter sterile water blank.
After shaking, add 1 cubic centimeter to a 99-cubic centimeter
sterile water blank (dilution 1:10,000).
Transfer 1 cubic centimeter of the above to a 9-cubic centi-
meter sterile water blank. As a rule, this dilution, which
represents 1 : 100,000 of a gram of soil to each cubic centimeter,
is the one from which to pour plates. If the soil is very poor,
use a dilution of 1:10,000; if very rich, 1:1,000,000. The
number of dilutions will depend on the type of soil. Garden or
100 LABORATORY MANUAL OF MICROBIOLOGY
well-cultivated soil rich in organic matter requires a higher
dilution than poor, sandy soil.
According to the total number of bacteria pour plates from
the dilutions 1:10,000, 1:100,000 and 1:1,000,000. In general
dilutions of about 1:100,000 are satisfactory. About 5 to 10
parallel plates for each dilution should be poured.
Add to each plate about 10 cubic centimeters of the agar
medium (Mediums 4 or 5), melted and cooled to about 45°C.
A blank plate or control should be poured with each series. In
case the medium is turbid, heat slowly, allowing the deposit to
Fig. 5. — Colonies of bacteria and fungi on Petri dish. (Lohnis and Fred.)
settle. Use only the clear portion of the medium for pouring
plates.
Immediately after adding the culture medium, rotate each
plate to secure a uniform mixture. Allow agar plates to harden
on a level surface for at least 30 minutes.
Agar plates should be inverted and incubated under a moist
chamber at 28°C. The time of incubation will depend upon the
microorganisms and the medium. After 5 to 10 days count the
number of colonies on each plate. If the colonies are not too
thick, it is well to dot each one with a pen and ink. When the
colonies are too thick to count easily, use a hand lens and count-
ing plate.
Reduce all results to number of bacteria in 1 gram of dry soil.
Exercise 7
Number of An^robic Bacteria in the Soil (Optional)
Prepare a series of dilutions of soil in sterile tap water as
previously outhned. Dilutions equal to 1:1,000 and 1:10,000.
THE STUDY OF MICROORGANISMS IN THE SOIL 101
One-cubic centimeter portions of the final dilutions are added
to 15 cubic centimeters of a sterile specific agar medium {e.g.,
glucose agar for Clostridium pasteurianum, Medium 11 or 12).
To remove oxygen from the medium steam for about 30 minutes
just before it is to be used. Now cool to 45°C. and add the
dilutions of soil, mixing well the soil suspension with the agar by
rotating the tube. The deep agar tube will produce anaerobic
conditions favorable for the development of the specific
organisms.
The colonies may be counted in the tube, after a definite
period of incubation (7 to 14 days).
Liquid media, incubated under anaerobic conditions may be
employed.
If it is desired to determine the number of anaerobic bacteria
present in the form of spores and vegetative cells, one portion of
soil is suspended in water and heated at 80°C. for 10 minutes,
which results in the destruction of the vegetative cells; the
further dilutions are then prepared from the first suspension.
If the colonies are to be isolated for pure culture study, allow
the agar medium to solidify and then pour on the surface of the
agar in the tubes or plates a layer of agar containing 0.05 per
cent of HgCl2, about 3^ centimeter in thickness.
Exercise 8
Number of Spore-forming Bacteria in the Soil
Prepare water suspensions of the soil as described in Exercise
6, making dilutions of 1 : 1,000, 1 : 10,000, and 1 : 100,000.
Transfer 10 cubic centimeters of each suspension to a sterile
test tube. Heat these inoculated tubes for 10 minutes at 80°C.
Cool quickly under the tap.
Place in Petri dishes 1 cubic centimeter portions of each of the
unheated and heated dilutions. Use three plates for each dilution.
Pour plates of sodium caseinat^ or nutrose agar (Medium 4)
and incubate at 28°C.
After 7 to 10 days count the number of colonies.
Exercise 9
Number of Thermophilic Bacteria (Optional)
Incubate several samples of soil and some fresh stable manure
at 60°C.
102 LABORATORY MANUAL OF MICROBIOLOGY
In order to prevent evaporation all samples must be kept
in a moist chamber. A large glass beaker or metal container may
be used. Avoid glass bell jars, unless of Pyrex glass, since the
high temperature may cause them to crack.
After 1 week in the incubator, prepare agar plates from the
different samples. Pour from dilutions 1:1,000 and 1:10,000.
The plates must be incubated at 60°C.
Determine the number of thermophilic bacteria in 1 gram of
soil.
If desirable, a study may be carried on of the bacteria growing
at low temperatures.
Exercise 10
Effect of Season of Year on Number of Bacteria
Collect samples of soil from an alfalfa field, clover field, blue-
grass field, orchard sod, etc. If the soil is frozen, it will be
necessary to use a pick or hatchet in securing samples.
Prepare dilution plates as soon as possible after the samples
reach the laboratory. Unless the soil is very rich use dilutions
of 1 : 10,000 and 1 : 100,000, about four or five parallel plates for
each dilution.
At the same time plates are poured, make moisture determina-
tions of the soil.
Record outside temperature and also soil temperature at the
time samples are drawn. Read the directions given in Exercise 6.
Exercise 11
Effect of Plant Roots on Number of Microorganisms
Collect soil samples from the immediate vicinity of the roots
of various plants (alfalfa, clover, etc.), and similar samples 1 or
2 feet away from the plants.
Determine the number of bacteria by means of plate counts.
Exercise 12
Effect of Depth of Soil on Number of Microorganisms
The samples for this exercise should be drawn from virgin soil
well removed from any source of contamination. The type of
soil will determine to a certain degree the number of organisms at
different depths.
Bacteria
Fungi
100,000
1 : 10,000
10,000 and
1:1,000
100,000
1,000 and
1:100
10,000
100 and
1:10
1,000
THE STUDY OF MICROORGANISMS IN THE SOIL 103
Divide the sample of soil, taking one portion for plate count,
the other for moisture determination. For virgin field soil the
following dilutions have been found satisfactory.
Take soil samples and plate as follows:
(a) Surface soil 1
(6) Soil 1 foot deep 1
1
(c) Soil 2 feet deep 1
1
(d) Soil 4 feet deep 1
1
Follow the method given in previous exercises.
Tabulate results.
Exercise 13
Effect of Manures on Number of Microorganisms
Prepare five tumblers or beakers with 100 grams each of field
soil.
Arrange as follows:
(a) Control. No treatment.
(6) Treat with 1 per cent of finely chopped green clover,
(c) Treat with 1 per cent of well-rotted stable manure.
(d) Treat with 1 per cent of ground wheat or rye straw.
(e) Treat with 1 per cent of ground wheat or rye straw and 0.1
per cent (NH4)2HP04.
Since these substances contain great numbers of bacteria,
especially the stable manure, plate counts should be made of the
manures at the time the soils are treated. For this purpose shake
5-gram portions of the manures with 5 cubic centimeters of
sterile water. Dilute as given in the previous exercises. Pour
plates from the dilutions 1:100,000 and 1:1,000,000 for bacteria
and 1 : 10,000 for fungi.
After mixing thoroughly the soil and manure in tumblers,
raise the moisture to two-thirds saturation.
Cover the soil with Petri dishes and incubate at room
temperature.
Determine the number of bacteria and fungi after 1 and 3
weeks.
104 LABORATORY MANUAL OF MICROBIOLOGY
Before drawing the sample for counts mix the contents of the
tumblers thoroughly. This may be done with a sterile spatula.
In the case of treated soils plate from the dilutions 1 '- 100,000 and
1:1,000,000 for bacteria, and 1:10,000 and 1:100,000 for fungi.
Exercise 14
Dilution Method for Determining the Number of Specific
Physiological Groups of Bacteria
This method is based upon the growth of different groups of
organisms upon specific substrates.
Prepare a series of dilutions of soil in sterile tap water, as 1 : 10,
1:100, 1:10,000, etc., and add two 1-cubic centimeter portions
of the final dilutions to flasks or tubes containing the desired
liquid medium or to plates containing the desired agar, gelatin
or silica-gel medium.
For example: For the determination of the number of Azoto-
bacter in soil, a medium free from nitrogen, containing mannitol
as a source of energy and of a pH 7.0 to 9.0 is used. For the
determination of the number of urea bacteria, a medium contain-
ing 2 to 5 per cent of urea as the only source of energy and nitro-
gen is used. To determine the number of nitrite forming
bacteria, a medium containing an ammonium salt and CaCOs
or MgCOs is employed.
Exercise 15
Direct Microscopic Examination of Soil^
Make a suspension of soil in nine times its weight of a 0.015
per cent solution of gelatin.
Smear a drop of this in a thin film on a slide and dry. (Both
the amount used and the size of the smear must be measured if
the number of bacteria is to be counted.)
Immerse for 1 to 3 minutes in a 40 per cent solution of acetic
acid or a O.liV solution of hydrochloric acid. Wash off the excess
acid quickly, and dry on a flat surface over a boiling water bath.
While still on the water bath cover the fllm with a 1 per cent
aqueous solution of Rose Bengal, or Erythrosin, and allow it to
stain for about 1 minute.
1 Conn, H. J., " Stain Technology," 1: 125, 1926; Soil Sci., 26: 257, 1928.
THE STUDY OF MICROORGANISMS IN THE SOIL 105
Exercise 16
Winogradsky's Method of Microscopic Analysis of Soil
Take several soil samples, mix carefully, and powder. Add 1
gram of soil (on dry basis) to 4 cubic centimeters of distilled water
and shake vigorously for 5 minutes.
Allow to settle for 30 seconds, then pour off suspension into
small tube of hand centrifuge.
Add twice 3-cubic centimeter portions of water to residue,
shake 1 minute, allow to settle for 30 seconds and pour into same
tube of centrifuge. These manipulations take about 10 minutes.
The combined extract (10 cubic centimeters) has in the meanwhile
formed another sediment.
Pipette off 5 cubic centimeters of the supernatant liquid
into another centrifuge tube and again centrifuge a few minutes;
a third sediment is formed.
Stained preparations are then made of the three sediments,
of the centrifuged and non-centrifuged suspensions.
A drop is placed upon a slide, to cover 1 square centimeter.
Dry in oven and cover with a small portion of 1 per cent warm
agar solution or 0.1 per cent cold agar solution.
When the agar has dried, a few drops of absolute alcohol are
added to fix the preparations.
This is followed by staining with a 1 per cent solution of
Erythrosin in 5 per cent phenol solution, allowing the dye to act
for 5 to 15 minutes in the cold or slight warming, then washing
a few seconds in water. Attempt to differentiate between short
rods and cocci, bacilli, Azotobacter cells, spores and filaments of
fungi, actinomyces filaments, and protozoan cysts.
Exercise 17
Determination of Numbers of Nematodes and Other Worms and
Insects in Soil^
Obtain several soil sampling tubes consisting of open cylinders
(made of tin or galvanized iron), 72.1 millimeters of internal
diameter, with the rim of one end sharpened and the other rein-
forced. The area of the internal cross-section of the tube is
1 Cobb, N. A., Bur. PI. Ind., U. S. Dept. Agr., Agr. Tech. Circ, 1, 1918.
106 LABORATORY MANUAL OF MICROBIOLOGY
just one-millionth of an acre. The tubes are usually 6 to 9
inches long.
By means of this tube, several samples are obtained from
different parts of the field. The soil of the various samples is
then well mixed and sifted.
An aliquot portion of soil is then suspended in 10 to 20 vol-
umes of water and well shaken.
The heavy particles of soil are allowed to settle for 5 seconds
and the supernatant liquid immediately poured into another
vessel.
The residue is washed two or three times with clean water and
the washings added to original liquid.
The liquid is allowed to run through a series of superimposed
sieves, ranging from 16- to 200-mesh per inch and the residual
material is washed with a little water. When the liquid flowing
through the finest sieve shows no animals, it is discarded.
The animals are washed away from each sieve with a small
amount of water and washings with animals placed in a series of
tubes.
The animals settle more readily to the bottom than the clay.
The suspended liquid can be removed after 30 minutes' settling
and replaced by clean water. The mixture of inorganic particles
and organisms is then examined under the microscope.
NITROGEN-FIXING BACTERIA
Exercise 18
The Isolation of Bacteria from the Root Nodules of Various
Leguminous Plants
Thoroughly wash the roots and nodules of several leguminous
plants {e.g., red clover, alfalfa or sweet clover, pea or vetch,
and soybean) under the tap.
Compare the number, size, color and position of the nodules on
the roots of these different leguminous plants.
Select a large and firm nodule, cut off, and immerse for 3 to 5
minutes in mercuric chloride solution (1:1,000), or in 70 per
cent alcohol. Remove alcohol or mercuric chloride by washing in
sterilized water and place the nodule on a sterile surface (flamed
slide or Petri dish).
THE STUDY OF MICROORGANISMS IN THE SOIL 107
If the nodule is small, crush, if the nodule is large, cut open
with a sterile knife and press out some of the inner contents into
a drop of sterile water.
Make two or more loop transfers from the first drop of water in
a Petri dish to a second containing a few drops of water.
Repeat these dilutions to a third and fourth Petri dish. Pour
Congo red mannitol agar plates (1 liter of Medium 79 + 10 cubic
centimeters of a solution of 1 gram of Congo red in 400 cubic
centimeters water), agitate until thoroughly mixed and incubate
at 28°C.
Fig. 6.-
-Bacteria from the nodules of pea ( X 1,500), stained with dilute carbol-
fuchsin.
Instead of diluting in the melted agar, a suspension of the
bacteria may be spread over the surface of hardened agar.
After 5, 10 and 20 days examine the plates. Fish off pure cul-
tures and transfer to slopes of Brom thymol blue mannitol agar.
Select about 5 pure cultures for each plant. The root nodule
bacteria usually form raised moist surface colonies, with round
edges, at first glistening, later changing to an opaque white.
Prepare stained mounts of the root nodule bacteria.
1. Fuchsin (carbol, according to Ziehl). Dilute with distilled
water, 1 part of stain and 9 parts of water.
108 LABORATORY MANUAL OF MICROBIOLOGY
2. Eryihrosin
(a) Erythrosin 5.0 gm.
Alcohol (70 per cent) 100 . 0 cc.
(6) Erythrosin 1.0 gm.
Carbolic acid (5 per cent) 100 . 0 cc.
This stain is especially recommended for root nodule bacteria.
1. Place a drop of the fresh culture on a glass slide, tilt the
slide to allow drop to spread. Dry the film in an oven at 45°C.
and fix in absolute alcohol.
2. After the alcohol evaporates, flood the mount with (a) and
allow to stain for 10 minutes.
3. If the stain is not deep enough, wash off this alcohol Ery-
throsin and stain with (6) for 10 minutes.
Exercise 19
Cultural Characteristics of Root Nodule Bacteria
Select three cultures from the Brom thymol blue slope cultures
of Exercise 18, and make subcultures on the various kinds of
culture media given below.
(a) Inoculate into litmus milk (Medium 90).
(6) Inoculate into bean extract in shallow layers (Medium 82).
(c) Inoculate on potato slopes (Medium 91).
{d) Inoculate on slopes of xylose, glucose, and sucrose-nitrate-
mineral salts agar to which Brom thymol blue has been added
(Medium 78).
After 1, 2, and 3 weeks, at 28°C., record growth and reaction
changes. In the milk tubes, note the effect on reaction and
serum zone formation. In the bean extract, note gum produc-
tion. After 4 to 5 weeks at 28°C., test for gum. Add 10 cubic
centimeters of alcohol (95 per cent) or 5 cubic centimeters of
acetone to 2.5-cubic centimeter portions of the bean extract
culture. Note the thickness of the layer of gum on top of the
liquid.
On potato, note growth and pigment formation. In the case of
Brom thymol blue agar, note the changes in reaction. Alfalfa
bacteria produce a strong acid reaction with all of the carbo-
hydrates listed. Clover bacteria produce a strong acid reaction
from glucose, little or no acid from sucrose and xylose. Pea
THE STUDY OF MICROORGANISMS IN THE SOIL 109
bacteria produce a strong acid reaction from glucose, medium
acid from sucrose and little or no acid from xylose. Soybean
and cowpea bacteria produce a strong alkaline reaction in these
culture media.
Prepare the following stain:
Barlow Stain for Root Nodule Bacteria
Glucose 50 . 0 gm.
Glycerol 50 . 0 cc.
Distilled water 50 . 0 cc.
Gentian violet 3.0 gm.
Dissolve the glucose in the glycerol-water
solution by heating and then add the
Gentian violet. Bring this mixture to a
boil and allow to cool.
Negative Stain. — Place a loopful of the
gum from an agar slant culture on a clean
slide and rapidly whip out into long thin
streaks. Dry in the air.
Flood with the Barlow stain, let stand for
30 seconds to 1 minute and wash off in run-
ning water. Blot off the excess water and
dry quickly with gentle heat.
Positive Stain. — Prepare the slide as
above.
Flood the slide with water and run the
stain into this water at one end of the slide.
As soon as the stain has diffused through
the water, wash in running water and dry
quickly.
Exercise 20
The Formation of Root Nodules in
Agar and Sand Cultures
Wash thoroughly the seed of alfalfa,
clover, and similar plants, which are to be
grown in agar under aseptic conditions, in
, ,. . ^ . 1, ., Fig. 7.— Growing plants
water and immerse m mercuric chloride free of bacteria. A Pyrex
solution (1:500) for 2 to 3 minutes. To ^l^^^ cylinder 4 inches in
,^ , , 1, i , ,1 1 .1 diameter and 24 inches tall
secure the best results, treat the seeds with covered with a beaker.
110 LABORATORY MANUAL OF MICROBIOLOGY
mercuric chloride in a partial vacuum. Rinse in sterile
water.
In the case of peas, cowpeas, soybeans, etc., which are to be
grown in pots of sterilized sand, the seed may be dipped in hot
water (65°C.) for 4 minutes to kill the nodule bacteria. Spread
out and dry quickly in an atmosphere free from dust.
Place three to five seeds each of alfalfa or red clover into a
large tube containing agar Medium 106.
Plant the peas, soybeans, and related plants, four or five
seeds, in half-gallon pots of sand which has been sterilized 4
hours in an autoclave at about 110 to 115°C. Cover the seed
with about 1 inch of sand and add sterile distilled water.
To inoculate, prepare a water suspension of a young culture of
the desired organism and from this take 1 cubic centimeter for
each tube or pot.
Peas or Soybeans (Pots) Alfalfa or Clover (Tubes)
(a) 1 and 2 uninoculated. 1, 2, and 3, uninoculated.
(6) 3 and 4 inoculated with known 4, 5, and 6, inoculated with known
culture. culture.
(c) 5 and 6 inoculated with an un- 7, 8, and 9 inoculated with an un-
known culture. known culture.
{d) 7 and 8 inoculated with an un- 10, 11, and 12, inoculated with an
known culture. unknown culture.
Under favorable conditions, nodules will begin to form on the
clover and peas in 10 to 15 days. With alfalfa and soybeans it
requires about 21 to 28 days.
Keep cultures in a greenhouse, free from dirt and dust.
Plants in open pots should be watered from time to time
with sterile distilled water. About once a week, add about 30
to 40 cubic centimeters of the nitrogen-free modified Crone's
solution.
After 4 to 6 weeks examine for nodules; note the number,
size, shape, and location.
Exercise 21
Artificial Cultures for the Inoculation of Legumes
Prepare two 6-ounce Signet bottles of mannitol agar (Medium
77) about 30 cubic centimeters of agar in each. After steriHza-
THE STUDY OF MICROORGANISMS IN THE SOIL 111
tion allow the bottles with the melted medium to cool in a sloped
position. When the agar has hardened the bottles are ready for
inoculation with the proper bacteria.
Select young cultures of root-nodule bacteria of alfalfa, or
clover and cowpea, or soybean from Exercise 13. Inoculate the
surface of the agar with three long streaks. Use a large loop
which has been bent at right angles.
Incubate the cultures at 28°C. or room temperature, for 4 to
10 days. Examine the cultures microscopically (Barlow stain)
and pack for shipping.
Exercise 22
The Structure of Root Nodules
A nodule consists mainly of a mass of large, thin-walled
nucleated cells often completely filled with bacteria, i.e., bac-
teroidal tissue. The action of the bacteria on the plant cells
as well as the infection threads may be seen from free hand
sections or better from paraffin embedded sections cut about 3
to 5 microns- thick.
A. Free-hand Sections. — Cut a cross-section through a young
pea nodule and root.
Mount in a drop of water and examine.
To secure a clear section, immerse it for 2 to 5 minutes in a
dilute sodium hypochlorite solution and stain with safranin.
B. Embedded Sections. — Select young nodules from the
desired plant grown in quartz sand or on filter-paper pads.
Fix by immersing for 24 hours in a formol-alcohol-acetic acid
mixture.
Acetic acid (glacial) 2 . 5 cc.
Formalin 6 . 5 cc.
Alcohol (50 per cent) 100 . 0 cc.
Transfer to 70 per cent alcohol and change the liquid until
there is no odor of acetic acid.
Dehydrate in 80, 90, and 100 per cent alcohol, clear in chloro-
form, embed in paraffin, and cut microtome sections 3 to 5 microns
thick. In order to stain the bacteria and the infection threads
use method A or B.
112
LABORATORY MANUAL OF MICROBIOLOGY
Method A. Flemming^s Triple Stain. — For general cytology
of the nodule. Vascular system, bacteroid area, shape and
position of infection threads, starch grains.
Remove paraffin from the sections in xylol 1 minute; rinse in
95 per cent alcohol 1 minute.
Stain 2 minutes in safranin (saturated solution in equal parts
of 95 per cent alcohol and anilin water). Rinse in water.
Fig. 8.-
-Cross-section of alfalfa nodule showing infection threads,
triple stain.
riemming'a
Stain 3 minutes in crystal violet (saturated aqueous solution).
Rinse in water.
Treat with Orange G (saturated aqueous solution) from a
medicine dropper. Time should be as short as possible. Rinse
with absolute alcohol. Destain with clove oil until the desired
differentiation of infection threads and bacteroid tissue is reached;
observe under microscope. Rinse quickly in absolute alcohol.
Clear in xylol 5 to 10 minutes.
Mount in Canada balsam.
Method B. Heidenhain's Haematoxylin. — For bacteria within
infection threads, internal structure of bacteroids.
THE STUDY OF MICROORGANISMS IN THE SOIL 113
Remove paraffin from the sections in xylol 1 minute; rinse in
95 per cent alcohol 1 minute.
Immerse in iron alum mordant (2 per cent aqueous solution)
4 to 24 hours. Over night is convenient. Rinse in water.
Stain in haematoxylin^ 1 to
6 hours.
Destain in the mordant until
desired differentiation is
reached; observe under
microscope.
Wash in running water 20
minutes to remove all iron alum
and thus prevent fading.
Counterstain in Licht grim
(saturated solution in 95 per
cent alcohol).
Dehydrate in absolute alco-
hol, clear in xylol, and mount
in balsam.
Exercise 23
Effect of Root Nodule Bacteria
ON the Growth and the Nitro-
gen Content of Alfalfa
2 -gallon jars
Prepare eight
of clean sand.
Plant to peas as follows :
(a) 1, 2, 3, 4, sand uninocul-
ated.
(h) 5, 6, 7, and 8, sand
inoculated.
One week after seeds begin to Fig. 9.— Effect of bacteria on growth
1 1 -.nrK L-- ,. of sweet peas. Jar on left not inocu-
germmate add 100 cubic centl- i^ted. Jar on right inoculated.
meters per jar of plant food
minus nitrogen (Crone's Medium 106). This nutrient solution
should be added at intervals of every 2 weeks or whenever needed.
1 Heidenhain's Haematoxylin. Dissolve 5 grams of haematoxylin
crystals in 400 cubic centimeters of absolute alcohol using heat if necessary.
Add 600 cubic centimeters of distilled water and allow the solution to ripen
a month or more.
114 LABORATORY MANUAL OF MICROBIOLOGY
After 6 to 8 weeks examine for nodules.
When mature, remove and analyze the tissue for total nitrogen.
Exercise 24
Direct Method for Demonstrating the Occurrence of Azotobacter
IN Soil
A. Macroscopic Method (Winogradsky).^ — To 100 grams of
freshly sieved soil, add 5 grams of pulverized starch. Mix the
starch thoroughly with the soil.
Transfer the starch-soil mixture to a shallow dish and add
water in sufficient amount to make a thick paste of the soil.
Avoid too much water.
Divide the soil paste between two Petri dishes. Press the soil
well into the Petri dishes, and smooth off the surface with a
wet glass slide. Make the top as smooth and even as possible.
Incubate the plates at 28°C. for 48 hours. If Azotobacter is
present in the soil, small limpid colonies will be observed on the
surface of the plate. Stain these colonies of organisms for 2 to
3 minutes with thionin or erythrosin. Pure cultures may be
obtained from these colonies (see Exercises 25 and 26).
B. Microscopic Method. — Thoroughly mix 0.5 gram of manni-
tol with 50 grams of soil. Place the mixture in a Petri dish, and
moisten the soil with water.
Incubate the plate at 28°C. for 48 hours.
Make an erythrosin or thionin stain of the soil. Azotobacter
cells will frequently be observed.
Anaerobic nitrogen fixing organisms (CI. pasteurianum) can
be cultivated by adding 0.5 gram of glucose to 50 grams of soil
and mixing thoroughly. Divide the soil-glucose mixture between
three large test tubes and saturate the soil with water.
Incubate the tubes at 28°C. and examine microscopically after
48 hours. Examine portions of the soil taken from different
depths in the tube.
Exercise 26
Isolation of Azotobacter from Various Soils
Prepare four small Erlenmeyer flasks of mannitol liquid
medium (Medium 77), about 20 cubic centimeters in each.
Avoid deep layers.
1 WiNOGRADSKY, S., Ann. Inst. Past., 40: 455 (1926).
THE STUDY OF MICROORGANISMS IN THE SOIL 115
Inoculate with 1 or 2 grams of soil and incubate at 28°C. and
note changes occurring in cultures, film formation, and color of
film.
After 2 days, examine the films in hanging-drop or wet mount
and note the predominating type of organism. Also examine
some of the surface film in a drop of water mixed with a drop of
Meissner's or Gram's iodin solution. Prepare stained mounts
Fig. 10. — Colonies of Azotobacter on mannitol agar, natural size.
with thionin and carbol erythrosin. The Erythrosin stain is
especially good for young cultures.
Dilute two loops of surface film in a 100-cubic centimeter
sterile water blank containing 50 grams of clean sand.
Shake vigorously, and transfer 1 cubic centimeter to a second
100-cubic centimeter blank, and so on to a third.
From the second and third dilution pour plates, using 1 cubic
centimeter for each. Because of the aerobic nature of Azotobacter
116 LABORATORY MANUAL OF MICROBIOLOGY
dilutions on the surface of the agar offer an easy and rapid method
of isolation. Sometimes it is difficult to separate Azotobacter
from a small organism known as Bacterium radiobacter.
After 4 to 6 days examine plates. The Azotobacter colonies
are raised, convex, smooth, white, semi-opaque, moist, viscid,
often 4 to 8 millimeters in diameter. Make transfers to mannitol
agar slants. Stain with carbolated rose bengal or erythrosin.
Note. — Instead of the enrichment liquid culture described above,
isolations may be made directly from the colonies on soil. Exercise 24.
Exercise 26
Use of Silica Gel Plate for Isolation of Specific Bacteria
Silica-gel plates are prepared as described under culture media
and dialyzed free from chlorides. A sterile solution containing
the necessary mineral salts and the specific substrate is placed
upon the surface of the gel. The uncovered plates are then
placed in a warm place (at 60°C.) to allow the evaporation of
excess liquid, care being taken that medium does not become too
dry.
The silica gel is then inoculated with small particles of soil.
After a few days incubation, growth will take place around each
particle of soil. The nature of the organism developing will
depend upon the nature of the medium added to the gel.
A mannitol medium free from combined nitrogen will allow the
development of Azotobacter under aerobic and of CI. pasteur-
ianum under anaerobic conditions.
A medium containing cellulose and an inorganic source of
nitrogen will allow the development of Spirochseta and other
cellulose decomposing bacteria.
When the colonies have developed sufficiently, they can be
transferred to specific nutrient media.
Exercise 27
Nitrogen Fixation by Pure Cultures of Azotobacter
Prepare four 1-liter Erlenmeyer flasks with 100 cubic centi-
meters each of mannitol agar (Medium 77). In place of the
THE STUDY OF MICROORGANISMS IN THE SOIL 111
flasks large pans or moist chambers may be used. The object
is to use a vessel that will give a large surface exposure.
After sterilization, inoculate the agar films with a pure culture
of Azotobacter. This may be accomplished by using 1-cubic
centimeter transfers from a suspension in sterile water.
Immediately after inoculation remove half of the cultures for
control analysis. These may be treated
with sulphuric acid or sterilized.
A few days after inoculation, add 5 cubic
centimeters of sterile water to each culture.
Incubate the cultures in such a position
that only a portion of the surface will be
covered with water, and from day to day
rotate. In this way it is possible to get an
f,, .1 ,. J. Fig. 11. — Azotobacter
even film over the entire surface. chroococcum, young cui-
About 28°C. is a favorable temperature tures. {Krzemieniewski.)
for growth.
After 21 days, analyze all of the cultures for total nitrogen.
Exercise 28
Effect of Variation in Carbohydrate on the Growth of Azotobacter
Prepare six tubes of culture Medium 77, without mannitol.
(a) 1 and 2, control (without mannitol) .
(6) 3 and 4, add 1 per cent of mannitol.
(c) 5 and 6, add 1 per cent of lactose.
Inoculate all cultures from a water
suspension of a young culture of Azoto-
bacter. Incubate tubes 1, 3, and 5 in
the open and tubes 2, 4, and 6 in the
anaerobic jar. Examine every 2 or 3
days for a period of 14 days.
Record the growth and pigment for-
mation of Azotobacter on the various
culture media. If the tubes kept under
anaerobic conditions fail to show a brown to black pigment
after 2 weeks, open the jar, incubate again and note change
in the color.
Fig. 12. — Azotobacter agile
showing flagella, X 660.
{Beijerinck.)
118 LABORATORY MANUAL OF MICROBIOLOGY
Exercise 29
An^robic Nitrogen Fixation (Clostridium pasteurianum and
Related Organisms)
Prepare four small flasks (or 6-ounce Signet bottles) with 150
cubic centimeters of Winogradsky's solution (Medium 11).
Arrange to have the liquid high in the necks of the flasks (or
bottles).
As soon as possible after sterilization inoculate all of the flasks
with a pasteurized soil extract. Heat 50 grams of soil with 200
cubic centimeters of water for 15 minutes at 80°C.
Allow the coarse particles to settle and pipette 5-cubic centi-
meter portions into each of the four flasks.
Arrange as follows:
(a) 1 and 2, controls (sterilize immediately, or add 5 cubic
centimeters of sulphuric acid. In order to avoid too rapid evolu-
tion of carbon dioxide, the sulphuric acid should be added slowly).
(6) 3 and 4, cultures of anaerobic bacteria. Incubate all flasks
at 28°C.
One week after incubation examine under the microscope a
drop taken from the lower layers of the liquid. This may be done
by inserting a pipette. Make a wet mount using Meissner's
solution. What forms are seen? Note the color. Make a
permanent spore stain, Dorner method.
After 21 days transfer the entire contents to a Kjeldahl flask
and analyze for total nitrogen.
Exercise 30
Isolation of An^robic Nitrogen-fixing Organisms (Clostridium
pasteurianum and Related Forms)
From the impure cultures of Exercise 26 or 29 make isolation
plates on agar medium (Medium 12).
From the bottom of an actively fermenting culture take 1 or 2
drops and spread on the surface of the glucose peptone agar in
Petri dish. Dilute by spreading over the surface of at least three
plates. To secure the best results use Petri dishes with porous
covers.
THE STUDY OF MICROORGANISMS IN THE SOIL 119
Incubate the plates under anaerobic conditions (potato jar)
and pick colonies into Winogradsky's liquid medium or corn
mash.
Fig. 13. — Dorner spore stain of anaerobic nitrogen-fixing bacteria.
Exercise 31
Nitrogen-fixing Capacity of Soil^
Large Petri dishes (20 centimeters in diameter) are filled with
the selective silica gel medium (see Exercise 26) containing 2
grams of mannitol.
The surface of the gel in the plate is inoculated with 1 gram of
soil (on dry basis).
Plates are incubated at 30°C. for 7 days.
The gel in the plates is now dried and contents transferred to
Kjeldahl flasks and total nitrogen determined.
1 WiNOGRADSKY, S., and J. ZiEMiECKA, Ann. Inst. Past., 42: 35, 1928.
120 LABORATORY MANUAL OF MICROBIOLOGY
Exercise 32
Influence of Nitrate on the Fixation of Nitrogen
Prepare eight large silica plates containing 2 grams of manni-
tol and medium for fixation of nitrogen. Arrange as follows:
(a) 1 and 2 no nitrate.
(6) 3 and 4 1.0 mgm. of nitrate nitrogen.
(c) 5 and 6 5.0 mgm. of nitrate nitrogen.
(d) 7 and 8 25 . 0 mgm. of nitrate nitrogen.
Inoculate each with 1 gram of good garden soil.
Incubate for 7 days at 30°C.
Dry gel and determine total nitrogen.
Tabulate results.
Exercise 33
Effect of a Soluble Carbohydrate on Nitrogen Assimilation
Weigh out two 400- to 500-gram portions of field or garden
soil in deep soup plates or shallow earthenware jars. The soil
should be thoroughly mixed and sieved.
(a) Control — untreated.
(6) Add 2 per cent of mannitol, glucose, sucrose, lactose, or
other carbohydrates.
Mix the carbohydrate thoroughly with the soil by means of a
spatula and then add tap water until the moisture content of the
soil is about two-thirds saturation. Do not attempt to mix soil
and carbohydrate immediately after the water is added. At
intervals of about 2 days add more water, sufficient to replace
the loss by evaporation and incubate at 28°C. for from 14 to 21
days. At the end of this time the carbohydrate should be
entirely destroyed.
Prepare the soil for analysis. When dry, pass it through a 20-
mesh sieve, mix thoroughly, and draw a small sample for analysis;
about 100 to 150 grams is enough. This smaller sample should
be pounded in a mortar until the entire mass passes through a
100-mesh sieve. Weigh out from three to six portions of 10
grams from each jar into 800-cubic centimeter Kjeldahl flasks
and analyze according to the Kjeldahl method not to include
nitrates (p. 65).
THE STUDY OF MICROORGANISMS IN THE SOIL 121
Make moisture determinations on the soil at the time samples
are taken for nitrogen analysis. Tabulate results.
DENITRIFYING BACTERIA
Exercise 34
Isolation of Denitrifying Bacteria
Fill three test tubes about two-thirds full of asparagin nitrate
solution (Medium 55).
Inoculate as follows:
(a) Control — no inoculation.
(h) Inoculate with approximately 0.1 gram of garden soil.
(c) Inoculate with approximately 0.1 gram of fresh manure.
Incubate at 28°C. until all nitrates have disappeared. The
destruction of nitrates is generally indicated by foaming.
At regular intervals, daily if possible, make qualitative tests
(spot plate) for the presence of nitrates, nitrites, and ammonia.
As soon as the nitrates are destroyed, transfer a loopful of the
old culture to a new tube of asparagin nitrate solution. This
may be repeated several times, although a pure culture is readily
isolated from the second transfer.
Follow the same method of isolation as given in the previous
exercises. It is well to make a series of not less than four dilu-
tions. Pour plates of asparagin nitrate agar and incubate
until there is a good growth.
Now pick off several isolated colonies, making transfers into
tubes of asparagin nitrate solution.
From the pure culture showing the most vigorous destruction
of nitrates make a transfer to the agar medium. Preserve this
pure culture for later study.
Exercise 35
Denitrification by Pure Cultures of Bacteria
Prepare four bottles, 125 cubic centimeters each, of asparagin
nitrate solution (Medium 55). Because of the total nitrogen
analyses to be made at the end of this experiment it is well to
prepare the medium with great care. Exact amounts of the
122 LABORATORY MANUAL OF MICROBIOLOGY
nitrate and asparagin salt, previously analyzed for total nitrogen
should be used.
(a) Control — not inoculated.
(6) Pure culture of unknown organism capable of bringing
about denitrification.
(c) Pure culture of unknown organism capable of bringing
about denitrification.
{d) B. pyocyajieus, B. fluorescens liquefaciens, or B. hartlehii.
After 2 to 3 days incubation at 28°C. make qualitative tests of
each culture for ammonia, nitrites, and nitrates. After 7 to 10
days incubation make quantitative determinations of nitrates
and total nitrogen.
Use the modified Kjeldahl method to include nitrates. For total
nitrogen analysis take portions of 50 cubic centimeters each of
the cultures.
Nitrates. — Take 10-cubic centimeter portions of the control,
dilute with 100 cubic centimeters of distilled water, and of this
evaporate 10-cubic centimeter portions to dryness.
In the case of the inoculated cultures with nitrates present,
take 2 samples and proceed as follows: (a) Evaporate 10 cubic
centimeters to dryness, and (b) dilute 10 cubic centimeters to 100
cubic centimeters and evaporate 10 cubic centimeters of this
to dryness.
Exercise 36
Denitrification in Soil
Prepare eight 100-gram samples of field soil in tumblers.
Add to each sample of soil 60 milligrams of nitrogen in the
form of potassium nitrate.
Treat the series as follows:
(a) 1 and 2, control untreated.
(6) 3 and 4, add 2.5 grams of glucose.
(c) 5 and 6, control untreated.
(d) 7 and 8, add 2.5 grams of glucose.
Mix these materials thoroughly by means of a spatula.
To soil portions 1 to 4, add sterile water to bring the moisture
content to about one-half saturation.
To soil portions 5 to 8 add sterile water to bring moisture up to
total saturation.
THE STUDY OF MICROORGANISMS IN THE SOIL 123
Incubate for two weeks at 28°C.
At the end of this time remove a sample for nitrate determina-
tion and dry the remainder for total nitrogen analysis. Use the
modified Kjeldahl method to include nitrates (see p. 67).
From these results calculate the percentage of the nitrogen
denitrified, and note the effect of excessive moisture and excessive
organic matter on the loss of nitrogen.
Tabulate results.
NITRIFICATION
Exercise 37
Nitrification in Impure Cultures
A. Nitrite Formation (Qualitative)
(a) Prepare five 150-cubic centimeter Erlenmeyer flasks with
20-cubic centimeter portions each of "ammonia" solution
(Medium 48).
(5) Inoculate two of the flasks and incubate at 28°C.
1. Add approximately 0.1 gram of field soil.
2. Add approximately 0.1 gram of garden soil.
(c) At regular intervals of 7 to 10 days remove, with a sterilized
platinum needle or pipette, 1 drop of the solution from each
flask and test as follows :
1. Nitrites — Trommsdorf's reagent.
Place 3 drops of Trommsdorf's reagent in a depression of a
spot plate. Add 1 drop of dilute sulphuric acid (1:3). Remove
a loopful of the solution to be tested and touch to surface of
reagent. A blue color indicates the presence of nitrites.
2. Ammonia — Nessler's reagent.
Place in a depression on a spot plate 1 drop of Nessler's solu-
tion. Remove a loopful of the solution to be tested and touch to
surface of reagent. Do not stir. A yellow-brown color indicates
ammonia.
(d) As soon as the culture shows the presence of nitrites and
absence of ammonia, make subinoculations into a sterile flask of
the same medium. If it is desired to study the nitrite bacteria
in enrichment cultures, repeated subinoculations may be made.
B. Nitrate Formation (Qualitative). — Prepare five 150-cubic
centimeter Erlenmeyer flasks with 20-cubic centimeter por-
tions each of nitrite solution (Medium 51).
124 LABORATORY MANAUL OF MICROBIOLOGY
Inoculate two of the flasks and incubate at 28°C.
1. Add approximately 0.1 gram of field soil.
2. Add approximately 0.1 gram of garden soil.
At regular intervals of 7 to 10 days remove, with a sterilized
platinum needle, 1 drop of the solution from each flask and test
as follows:
1. Absence of nitrites — Trommsdorf's reagent.
2. Presence of nitrates — Diphenylamine reagent.
Place in a depression on a spot plate a drop of concentrated
sulphuric acid plus diphenylamine. Touch with a drop of the
solution to be tested. A deep-blue color indicates nitrates.
This test cannot be made in the presence of nitrites, chloric, and
selenic acids, ferric chloride, and many other oxidizing agents.
If the test for nitrites (Trommsdorf) is positive do not make this
test.
As soon as the culture shows the presence of nitrates and
absence of nitrites, make loop subinoculations into a sterile flask
of the same medium. If it is desirable to study the nitrate bac-
teria in enrichment cultures, repeated subinoculations may be
made.
Exercise 38
Nitrification in Liquid Cultures (Quantitative)
Prepare two flasks (750-cubic centimeter capacity) with
100 cubic centimeters each of Medium 52.
Inoculate each flask with 1 cubic centimeter of enrichment
culture — ammonia oxidizing — and 1 cubic centimeter of nitrite
oxidizing culture.
Incubate the flasks at 28°C.
Once a week test the solution for the oxidation of ammonia and
nitrite. If this process is complete, add 1 cubic centimeter of a
10 per cent solution of ammonium sulphate. Repeat until the
oxidation of the ammonium sulphate ceases. If necessary, add a
small amount of magnesium carbonate to the culture.
After the cultures cease to oxidize, make qualitative tests for
ammonia, nitrite, and nitrate. If nitrates are present, make
quantitative analyses by the phenoldisulphonic method as given
on page 63.
\
THE STUDY OF MICROORGANISMS IN THE SOIL^ 125
Exercise 39
Isolation of Nitrifying Organisms
Prepare eight tubes of acid sodium-potassium silicate.
Dilute the second enrichment cultures of the nitrite and
nitrate organisms until 1 cubic centimeter represents 1:1,000,
1:10,000, 1:100,000, and 1:1,000,000 of the original culture.
Pour the acid silicate into a sterile Petri dish with the culture
dilutions; add the nutrient salts and enough sodium carbonate
to harden the silicate.
Fig. 14. — Nitrobacter in liquid culture.
When hard, invert the plates and incubate under a moist
bell jar for 3 to 6 weeks. The cultures may also be streaked on
the surface of silica gel.
Examine at weekly intervals, using the low-power objective.
As soon as small colonies appear, make transfers to sterile nitrite
or nitrate solution.
The nitrifying organisms may be also grown on washed agar.
Exercise 40
Nitrification of Various Substances
Prepare two portions of soil, 500 to 700 grams each, in small
jars. Mix and sieve the soil well before using,
(a) Control — no treatment.
126 LABORATORY MANUAL OF MICROBIOLOGY
(b) Add 30 milligrams of nitrogen per 100 grams of soil from
(NH4)2S04, blood meal, casein, or peptone.
Mix these substances thoroughly with the soil, add sterile
water to give one-half to two-thirds saturation, record weight
and incubate at 28°C.
Each week, weigh and restore loss of water by evaporation.
After 10 and 20 days determine the nitrate nitrogen.
Express the results as milligrams of nitrate nitrogen in 100
grams of soil; also as percentages of the original substance
nitrified.
Tabulate results.
UREA AND PROTEIN DECOMPOSITION
Exercise 41
Decomposition of Urea
Prepare three Erlenmeyer flasks or bottles, 200-cubic centi-
meter capacity, with 50 cubic centimeters each of urea solution
(Medium 45).
After sterilizing, arrange as follows:
(a) Control — not inoculated.
(h) Inoculate with 1 gram of soil.
(c) Inoculate with 1 gram of fresh manure.
It is not necessary to weigh accurately the soil or manure.
Incubate the cultures at 28°C. and after 2 days remove from
each flask 5-cubic centimeter portions of the solution, with a
sterile pipette, to a small Erlenmeyer flask. Add about 50 cubic
centimeters of distilled water to the urea solution and a few drops
of methyl red. Determine the ammonia production by titrating
with N/14: acid.
From the results of the titrations, calculate the amount of
ammonia nitrogen in 100 cubic centimeters of the different urea
solutions. In order to find the amount of ammonia formed by
bacterial action, subtract the untreated from the treated series.
Determine the percentage of urea nitrogen transformed into
ammonia nitrogen.
Similar samples may be drawn after 3 or 4 days and the
amount of ammonia determined. Tabulate the results.
THE STUDY OF MICROORGANISMS IN THE SOIL 127
Exercise 42
Isolation of Specific Urea-decomposing Organisms
Pour isolation plates of several dilutions of the enrichment
cultures of Exercise 41. Use urea agar or gelatin Medium 47.
Incubate the plates at 28°C.
Examine the plates every 48 hours for a period of 10 days.
The urea organisms are often characterized by a distinct halo
around the colonies. Under the low power of the microscope the
halo is composed of crystals.
Make transfers to tubes of urea solution Medium 44 or 45
and incubate for two days.
Now test the ammonia-producing power of the pure cultures
by inserting sterilized strips of Nessler's paper or tumeric paper
in the upper part of the tube.
Prepare a stained mount of these organisms.
Exercise 43
Ammonia Production from Various Substances in Soil
Prepare 500-gram portions of soil in small jars. The soil
should be mixed thoroughly.
1. Control — no treatment.
2. Soil plus 0.5 per cent of blood meal, casein, peptone, alfalfa
meal, soybean meal, or similar substances.
After these substances are mixed with the soil bring the mois-
ture content of the soil to two-thirds saturation. In order to
secure the proper moisture content it is necessary to take into
account the water-holding capacity of the added substances.
Incubate at room temperature and, after 2, 5, and 7 days,
determine the ammonia nitrogen.
Ammonia may be estimated in a number of ways. The
aeration method is usually considered the most accurate although
good results may be obtained by the leaching of the soil with a
strong chloride solution and distilling with magnesium oxide.
Determination of Ammonia Nitrogen. ^ — Weigh out 50 grams
of soil in a 750-cubic centimeter Pyrex Erlenmeyer flask. Add
500 cubic centimeters of a 10 per cent KCl solution (in case of neu-
tral or alkaline soils use a 20 per cent KCl solution) ; shake for
1 Harper, H. J., Soil Sci., 18: 409, 1924.
128 LABORATORY MANUAL OF MICROBIOLOGY
30 minutes in a mechanical shaker; allow to settle 10 minutes
and filter on a large folded filter. Transfer 400 cubic centi-
meters of filtrate to a Kjeldahl flask, add a small piece of paraffin
to prevent foaming and a little 20-mesh limestone to prevent
bumping. Distill with 1 gram of MgO.
Exercise 44
Decomposition of an Amino Acid and a Protein by Bac. oereus and
Bact. fluorescens
Prepare 1 liter of a mineral salt solution.
Weigh out 3 grams of K2HPO4, 0.2 gram KCl, 0.2 gram MgS04,
0.2 gram NaCl, 0.1 gram CaS04 and 0.01 gram FeS04, in 1,000.0
cubic centimeters of water.
To 500 cubic centimeters add 5 grams of glycocoll and mark
solution '' Glycocoll medium;" the other 500 cubic centimeters is
used for the casein solution.
Weigh out into a beaker 5 grams of casein, purified (Hammer-
sten). Add about 50 cubic centimeters of the medium and sus-
pend the casein in it. Then add 4 cubic centimeters of IN.
NaOH solution, warm until casein has dissolved. Filter. Add
casein solution to the rest of the salt solution, make up to 500
cubic centimeters and mark it ''Casein solution."
Distribute the two solutions in 50-cubic centimeter portions in
250-cubic centimeter Erlenmeyer flasks, plug with cotton, and
sterilize.
Inoculate two flasks of each medium with a pure culture of
Bacillus cereus and two flasks of each medium with a pure cul-
ture of Bacterium fluorescens. Leave 1 flask of each as control.
Incubate 14 days.
Examine cultures for growth and determine in each flask the
amino-acid-nitrogen, ammonia, and residual casein.
Tabulate results.
SULPHATE-REDUCING AND SULPHUR-OXIDIZING BACTERIA
Exercise 45
Reduction of Sulphates by Bacteria
Prepare three small bottles of sulphate medium (Medium 60),
and treat as follows:
(a) Control — uninoculated.
THE STUDY OF MICROORGANISMS IN THE SOIL 129
(b) 1 gram of rich garden soil.
(c) 1 cubic centimeter of sewage.
Stopper tightly with paraffined corks and incubate at 28°C.
for 2 or 4 weeks.
At the end of this time remove bottles from incubator and note
the change in color and odor.
Hold over the open mouth of the bottle a small piece of filter
paper saturated with a solution of lead acetate. A blackening
of the paper indicates the presence of hydrogen sulphide.
Remove a few cubic centimeters with a pipette to a test tube
or small Erlenmeyer flask and add a few drops of BaCU solution.
Compare the amount of white precipitate in the inoculated
cultures with that in the uninoculated control.
The amount of hydrogen sulphide may be determined quanti-
tatively by titrating with iodine and sodium thiosulphate.
Exercise 46
Isolation of Hydrogen Sulphide-forming Microorganisms
Prepare five deep tubes of Medium 62.
Make shake cultures, deep tubes, from varying dilutions of the
enrichment cultures of Exercise 45 (reduction of sulphates with
the formation of H2S).
After the inoculated tubes have hardened, cover with glycerol
agar to a depth of about 2.5 centimeters. Incubate at 28°C.
and note the formation of colonies. After 5 to 10 days these
colonies become brown to black in color. Pure cultures may be
obtained from these deep colonies by cutting the tubes and
transferring the black colonies to fresh tubes of Medium 62.
Glycerol Agar^
Agar, washed 20 . 0 gm.
Glycerol (C3H5(OH)3) 500.0 cc.
Distilled water 500 . 0 cc.
^ Dissolve the agar in the water by heating in a steamer, add the glycerol
and filter.
130 LABORATORY MANUAL OF MICROBIOLOGY
Exercise 47
Crude Cultures of Higher Sulphur Bacteria
Place some mud from ditches or other bodies of water in a large
flask or cylinder and cover to a depth of 3 to 6 inches with fresh
or salt water.
Add 5 grams of magnesium sulphate and 2 grams of calcium
carbonate per liter of water. This furnishes a supply of H2S
due to the action of sulphate-reducing bacteria present in the
mud.
Inoculate the solution with material that contains the sulphur
bacteria, namely pond scum, submerged plant material, organic
matter from sulphur springs, etc. H2S may also be used in the
form of a gas.
Incubate in the dark for colorless sulphur bacteria, while, for
colored forms, the cultures are exposed to transmitted light.
Growth may appear on surface of liquid, along walls of container,
or on surface of mud.
Concentration of H2S, amount of mud added to flasks, purity
of water used, amount and nature of inoculum will all influence
the type of organism developing. The red forms develop better
under higher partial pressures of H2S than the colorless forms.
Exercise 48
Isolation of Pure Cultures of Higher Sulphur Bacteria
100-cubic centimeter portions of Medium 70 are placed in
250-cubic centimeter Erlenmeyer flasks and steriUzed, the
ammonium sulphate being steriUzed separately in stock solutions,
then added to the rest of the medium, after sterilization.
Inoculate flasks from enrichment cultures obtained in previous
experiment, using as little foreign material as possible.
The flasks are then placed under jar (e) into which hydrogen
sulphide is introduced, as shown in Fig. 1 (p. 30).
The sulphide is generated by the action of HCl upon FeS in
(a).
Hydrogen, CO2, oxygen, or air may be introduced from com-
pressed gases (5) in regulated amounts controlled by manometer
attachment (/).
THE STUDY OF MICROORGANISMS IN THE SOIL 131
Agar media containing CaCOa may be employed for isola-
tion purposes.
Frequent transfer of the cultures and incubation in H2S
atmosphere will finally lead to isolation of pure cultures.
Exercise 49
Oxidation of Sulphur and the Dissolving of Rock Phosphate
Prepare a mixture of 60 parts good garden soil, 30 parts of
finely ground rock phosphate, and 10 parts of powdered sulphur.
Place in pots or tumblers.
Add enough water to bring to 60 per cent saturation.
Incubate at 25°C.
At the end of every week, mix mixture well and add enough
water to keep at optimum.
After 2, 7, 21, 42, 70, and 100 days, determine pH value,
amount of water soluble sulphate, and phosphate. Draw curves.
After 2 and 60 days, determine also number of bacteria and
fungi by ordinary plate method.
Exercise 50
Growth and Isolation of Thiobacillus thioparus
Prepare 600 cubic centimeters of Medium 65. Distribute,
in 100 cubic centimeter portions, into 250 cubic centimeter
Erlenmeyer flasks. Do not sterilize.
Inoculate two flasks with 1-gram portions of garden soil, two
with fresh manure. Incubate flasks.
Test every 3 days for disappearance of thiosulphate, using a
dilute iodine solution.
The flasks where the thiosulphate has first disappeared are used
for inoculation of two flasks with fresh medium.
After the thiosulfate has disappeared in the second lot of flasks,
examine culture microscopically.
Note bacteria and granules of precipitated sulphur.
Culture is now plated upon thiosulphate agar.
Incubate.
Isolate from the sulphur-yellow colonies into flasks with sterile
liquid medium, in which the nitrogen salt and carbonate have
been sterilized separately and mixed aseptically.
132 LABORATORY MANUAL OF MICROBIOLOGY
Exercise 61
Growth of Thiobacillus Thiooxidans in Liquid Medium
Prepare six flasks of medium (Medium 68).
Inoculate two flasks with 1-gram portions of compost from
Exercise 49 (100 days old), two flasks with 0.1 gram of same
compost. Two flasks are left as controls.
Incubate flasks at 25°C.
After 7, 14, and 30 days, determine pH of flasks.
Fig. 15. — Thiobacillus thiooxidans (X 1,000).
If there is an increase in acidity in the inoculated flasks above
the control, determine sulphates in solution in all flasks, and
examine culture microscopically.
Repeated transfers of the culture upon fresh sterile medium will
result in a highly enriched culture of the organism. Final isola-
tion can be made by the use of the agar plate. ^
iWaksman, S. a., Jour. Bact., 7: 605-608, 1922.
IRON BACTERIA
Exercise 52
Iron-precipitating Bacteria (Optional)
Shake 20 grams of field soil with 200 cubic centimeters of
water. Dilute until 1 cubic centimeter equals 1 : 100,000.
Pour plates of Medium 101 or similar formula.
Incubate the plates for several weeks at 28°C.
Note the precipitation of iron compounds around certain
colonies.
THE STUDY OF MICROORGANISMS IN THE SOIL 133
Exercise 63
Iron Bacteria from Drinking Water
Clean and sterilize a Berkefeld filter.
Connect the filter to the city water-supply
and allow the water to run slowly for 24 to
48 hours.
After the metal cap is removed from the
filter, place the filter in a large beaker of
iron solution (Medium 76).
Incubate in the ice-box or at about 15 to
20°C.
At regular 2-day intervals examine the
deposit on the sides of the filter.
If bacteria are found, test for iron. Add
a few drops of a 5 per cent hydrochloric acid
solution and a 4 per cent potassium ferro-
cyanide solution. In the presence of ferric
salts an intense blue color is formed.
In order to stain the higher forms of iron
bacteria it is well to remove the deposit of
iron by treating with a 5 per cent hydro-
chloric acid solution.
CELLULOSE-DECOMPOSING BACTERIA
Exercise 54
An^robic Cellulose Decomposition in
Impure Cultures
Fill four large test tubes or wide-mouthed
deep-form bottles about three-fourths full
of Omeliansky's solution (Medium 85).
Add four strips of filter paper to each.
Treat as follows: (1) uninoculated, (2)
stable manure, (3) leaf mold, (4) garden
soil.
Incubate at 28°C.
Examine the cultures at regular intervals,
taking note of the change in color and the
structure of the filter paper.
Fig. 16. — Fermenta-
tion of filter paper in
Omeliansky's solution.
134 LABORATORY MANUAL OF MICROBIOLOGY
When the filter paper shows evidences of disintegration, make
transfers to new tubes of Omeliansky's medium (enrichment
cultures).
Exercise 65
Number of Aerobic Cellulose-decomposing Bacteria in Soil^
Prepare a series of medium-sized test tubes containing 5 cubic
centimeters of Medium 85 (the CaCOa may be left out and a
trace of FeS04 introduced) and a strip of
paper. Part of the paper should protrude
above the surface of medium.
Prepare a series of soil dilutions (1 : 10,
1:100, 1:1,000) and inoculate 1 cubic
centimeter of each dilution into one tube
of medium.
Take out, with a sterile pipette, 1 cubic
centimeter from the tube inoculated with
cytlphaga^'^'^luUnTol 1 ' l^O^O dilution and add to a fresh tube of
and Clayton.) sterile medium, to give a dilution of
1:5,000.
If further dilutions are desired the last step is repeated.
The tubes are incubated at 25 to 30°C. and examined daily.
Presence of cellulose-decomposing bacteria will be shown by the
decomposition of the paper just at the surface of the liquid.
Examine bacteria by Nigrosin method.
Exercise 56
Themophilic Fermentation of Cellulose
Prepare four long tubes (about 25 centimeters each) with
20 cubic centimeters each of the Cellulose peptone medium
(Medium 89).
Treat as follows:
(a) Control — not inoculated.
(6) About 1 gram of fresh horse manure.
(c) About 1 gram of well-rotted leaf mold.
(d) About 1 cubic centimeter of a culture of Clostridium ther-
mocellum.
1 DuBos, R. J., J. BacL, 15 : 223, 1928.
THE STUDY OF MICROORGANISMS IN THE SOIL 135
Cover the tubes with tinfoil and incubate at 60 to 65°C.
As soon as the cultures show active fermentation make 1 cubic
centimeter transfers to tubes of fresh media. This enrichment
process may be repeated several times.
Examine young and old cultures under the microscope.
Exercise 57
Isolation of Cellulose Decomposin-g Bacteria
Select samples of soil from plots which have received applica-
tions of straw or materials rich in cellulose. From these samples
prepare dilutions of about 1 : 10,000 and 1 : 100,000. Instead
of soil, enrichment cultures may be used (see Ex. 54, 55). If
Fig. 18. — Bacillus cellulosae dissolvens attacking filter paper. (Khouvine.)
enrichment cultures are used the dilutions should be greater than
those from soil.
Pour plates of the dilutions from soil or enrichment cultures
with the cellulose agar medium (Medium 88).
Because of the long period of incubation, 2 to 4 weeks, it is
desirable to keep the plates in a moist chamber, under bell jar
at 28°C. At regular intervals, 5, 10, 20, and 30 days, examine
the plates. Note change in color and clear zones throughout
the opaque agar.
Make microscopic mounts — nigrosin and carbol fuchsin.
136
LABORATORY MANUAL OF MICROBIOLOGY
Silica gel containing macerated cellulose on surface may also
be used for the isolation of cellulose-decomposing bacteria.
Exercise 58
The Evolution of Carbon Dioxide from Soil
Weigh out 100 grams of well mixed soil into 500-cubic centi-
meter Erlenmeyer flasks or other suitable vessels. Arrange as
follows :
1. Control, soil alone. No treatment.
2. Soil plus 1 per cent of glucose, starch, cellulose, alfalfa meal,
blood meal or similar substances in the powdered form.
Bring the moisture content of the soil to two-thirds saturation.
Fig. 19. — Apparatus for measuring carbon dioxide evolution from soil.
After the test substance is well mixed with the soil set up the
apparatus in such a way that the CO2 evolved will be absorbed
in 25 cubic centimeters of 0.2 N Ba(0H)2 solution in a large test
tube. The large tower A of Fig. 19 is filled with soda-lime and
connected to B with a glass tube. The end of this tube carries a
gas washing tube. B is filled about two-thirds full of water and
connected to C. In this way moist CO2 free air is carried to C,
In flask C is the soil sample. £' is a test tube containing the
Ba(0H)2 solution. The gas is dispersed into fine bubbles by
means of a Rose end tube. D is a trap to catch the alkali solu-
tion in E if there is any back pressure, and the alkali in E sucks
back. As soon as possible, after the soil is mixed with the
various substances, begin a slow aeration at the rate of one
THE STUDY OF MICROORGANISMS IN THE SOIL 137
bubble per second. Every 24 hours, titrate the Ba(0H)2 solu-
tion to the phenolphthalein end-point with 0.1 iV oxalic acid
(6.3 grams per liter). Continue the absorption from day to
day until the CO 2 evolution declines to nearly a constant amount.
Barium hydroxide reacts with CO2 according to the following
equation :
Ba(0H)2 + CO2 = BaCOs + H2O. O.IA^ Ba(0H)2 = O.IN
CO2 = 2.2 milligrams.
The excess Ba(0H)2 is titrated with oxalic acid. The acid
neutralizes only the Ba(0H)2. Ba(0H)2 + (C00H)2 = Ba-
(C00)2 + 2H2O.
The BaCO.s remains unchanged. The difference between the
excess of Ba(0H)2, as determined by oxalic acid titration, and
the Ba(0H)2 taken at the beginning, will give the amount of
Ba(0H)2 acted upon by the CO2.
An example of the calculation follows :
Ba(0H)2, O.IA^, factor 1.06 50 cubic centimeters
Oxalic acid, O.IA^ 12.6 cubic centimeters
50 X 1.06 53.0 cubic centimeters O.liV
Back titration (oxalic) 12.6
CO2 evolved 40.4 cubic centimeters O.liV
Since 1 cubic centimeter O.IA^
Ba(0H)2 takes up 0.0022 gram CO2
40.4 X .0022 0.0889 gram CO2 per 100
grams of soil.
When the CO2 evolution has declined to nearly a constant
amount, plot a curve showing the rate of evolution of the gas.
Plot milligrams of CO2 as ordinates, and days as abscissae.
LITERATURE
The following list includes some of tEe more important books that treat of
bacteriology :
A. General Bacteriology:
Benecke, W. : " Bau und Leben der Bakterien," 2d Ed., Leipzig, 1924.
Buchanan, E. D. and R. E. Buchanan: "Bacteriology for Students in
General and Household Science," rev. Ed. 560 pp., 360 figs., New York,
1926.
Buchanan, R. E., and Fulmer, E. I.: " Physiology and Biochemistry of
Bacteria," 516 pp., Baltimore, 1928.
138 LABORATORY MANUAL OF MICROBIOLOGY
Conn, H. J. and W. H. Conn: "Bacteriology," 3d Ed., Baltimore, 1926.
Ford, W. W.: "Textbook of Bacteriology," 1,069 pp., Philadelphia, 1927.
Greaves, J. E.: "Elementary Bacteriology," 506 pp., 129 ills. Phila-
delphia, 1928.
Jordan, E. O.: "General Bacteriology," 9th Ed., 778 pp., 191 ills., Phila-
delphia, 1928.
Jordan, E. O. and I. S. Falk: "The Newer Knowledge of Bacteriology
and Immunology," 1,196 pp., Chicago, 1928.
Kelser, R. a.: "Manual of Veterinary Bacteriology," 525 pp., Baltimore,
1927.
Kendall, A. I.: "Bacteriology, General, Pathological and Intestinal,"
2d Ed., 680 pp., Philadelphia, 1921.
Kruse, W.: "Allgemeine Mikrobiologie," 1184 pp., Leipzig, 1910.
LiESKE, R. : "Allgemeine Bakterienkunde," 338 pp., 118 figs., Berlin, 1926.
Park, W. H., A. W. Williams, and C. Krumwiede: "Pathogenic Micro-
organisms," 8th Ed., 811 pp., 211 figs., 9 plates, Philadelphia, 1924.
RippEL, A.: "Vorlesungen iiber Theoretische Mikrobiologie," 171 pp.,
Berlin, 1927.
Zinsser, H. and S. S. Tyzzer: "A Textbook of Bacteriology," 1,193 pp.
6th Ed., New York, 1927.
B. Agricultural Bacteriology:
FuHRMANN, F. : "Einfiihrung in die Grundlagen der technischen Myko-
logie," 2d Ed., 554 pp., Jena, 1926.
Greaves, J. E.: "Agricultural Bacteriology," Philadelphia, 437 pp. 1922.
Greaves, J. E. and E. O. Greaves: "Bacteria in Relation to Soil Fertil-
ity," 239 pp., New York, 1925.
Kayser, E.: " Microbiologie Agricole," 4th Ed., 1921.
LoHNis, F. and E. B. Fred: "Agricultural Bacteriology," 283 pp., 66 figs..
New York, 1923.
Marshall, C. E.: "Microbiology," 3d Ed. 1,043 pp., 200 ills., Phila-
delphia, 1922.
Russell, E. J., et al. "The Microorganisms of the Soil," 188pp., 23 figs.,
London, 1923.
Russell, H. L. and E. G. Hastings: "Agricultural Bacteriology," 368
pp., 63 figs.. New York, 1921.
Waksman, S. a.: "Principles of Soil Microbiology," 897 pp., 77 figs., 19
plates, Baltimore, 1927.
C. Reference Books in Bacteriology:
Henneberg, W. : "Handbuch der Garungsbakteriologie," Bd. 1 and 2,
Berlin, 1926.
de Rossi, G.: " Microbiologia Agraria e Tecnica," 1410 pp., Torino,
1921-1927.
Lafar, F.: "Handbuch der technischen Mykologie," Bd. 3, 503 pp., 10
plates, 90 figs., Jena, 1904-1907.
LoHNis, F. : "Handbuch der landwirtschaftlichen Bakteriologie," Berlin,
907 pp., 1910.
Smith, E. F.: "Bacteria in Relation to Plant Diseases," Vols. 1, 1905; 2,
1911; 3, 1914, Washington.
THE STUDY OF MICROORGANISMS IN THE SOIL 139
Stoklasa, J. and E. G. Doerell: "Handbuch der biophysikalischen und
biochemischen Durchforschung des Bodens," 812 pp., Berlin, 1926.
D. Manuals of Bacteriologic Technic:
Abel, R.: ''Bakteriologisches Taschenbuch," 1927.
American Public Health Association: "Standard Methods for the Exam-
ination of Water and Sewage," 6th Ed., New York, 1925.
Conn, H. J.: *'An Elementary Laboratory Guide in General Bacteri-
ology," 98 pp., 27 figs., Baltimore, 1927.
Cunningham, A.: "Practical Bacteriology," 188 pp., London, 1924.
Giltner, Ward: "Laboratory Manual in General Microbiology," 3d Ed.,
472 pp.. New York, 1926.
Hastings, E. G. and W. H. Wright: "A Laboratory Manual of General
Agricultural Bacteriology," 83 pp., 28 figs., Madison, 1927.
Levine, M. : "Laboratory Technique in Bacteriology," 149 pp., New York,
1927.
Norton, J. F. and I. S. Falk: "Laboratory Outlines in Bacteriology and
Immunology," 114 pp., Chicago, 1926.
Koch, A.: " Mikrobiologisches Praktikum," 109 pp., 4 figs., Berlin, 1922.
KusTER, E.: "Kultur der Mikroorganismen," 3d Ed., Leipzig, 1921.
Lohnis, F.: "Landwirtschaftlich-bakteriologisches Praktikum," 3d Ed.,
BerHn, 1926.
MuiR, R. and J. Ritchie: "Manual of Bacteriology," 7th Ed.; 753 pp.,
London, 1921.
E. Classification of Bacteria:
Bergey, D. H.: "Manual of Determinative Bacteriology," 2nd Ed.,
Baltimore, 442 pp., 1925.
Buchanan, R. E.: "General Systematic Bacteriology," 597 pp., Balti-
more, 1925.
Lehmann, K. B. and R. O. Neumann: " Bakteriologie,. insbesondere
bakteriologische Diagnostik.," I Band, "Technik, Allgemeine Diagnos-
tik, Atlas," 172 pp., 65 plates. Mfmchen, 1926; II Band: "Allgemeine
und spezielle Bakteriologie," 876 pp., 43 figs., Miinchen, 1927.
LIST OF LABORATORIES
WHERE CULTURES MAY BE SECURED
1. American Type Culture Collection,
George H. Weaver, Curator,
637 South Wood Street,
Chicago, Illinois.
2. The Lister Institute of Preventive Medicine,
National Collection of Type Cultures,
Chelsea Gardens, London, S. W., England.
3. Professor Dr. E. Pribram's mikrobiologische Sammlung,
vorm. Krai's bakteriolog. Museum,
Wien, IX/2. Michelbeuerngasse la,
(Wien) Vienna, Austria.
140 LABORATORY MANUAL OF MICROBIOLOGY
4. Centraalbureau voor Schimmelcultures,
Professor J. Westerdijk,
Baarn, Holland.
5. Professor Alfred Jorgensen,
Laboratory of Fermentology,
Copenhagen, Denmark.
6. Professor Dr. E. de Herics-Toth,
Roy. Hungarian Institut of Fermentology,
Budapest II. Debroi u. 15. Hungary.
7. L' Institut Pasteur,
Director, Dr. E. Roux,
Institut de Microbiologie,
25 Rue Dutot,
Paris 15. France.
INDEX
Acid, resistant finish for tables, 91
sucrose agar, 13
Actinomyces, media for, 18
Aeration method for ammonia deter-
mination, 63
Agar, chemical composition, 5
general properties, 5
washed, 5
Albuminate agar, 9
Algse, culture media for, 20
in soil, 93
number in soil, 97
AUzarine, 52
Amino acids, decomposition of, 138
nitrogen, 69
Ammonia, determination in soil, 127
quantitative, 62
production, 127
test for, 56
Anaerobic bacteria, 11, 43
culture media, 11
number in soil, 100
cellulose fermentation, 133
culture methods, 46
nitrogen fixation, 118
Analysis of plant material, complete,
76
Apparatus for one student, 89
Artificial cultures, 110
Ashby's medium, 20
Asparagin, mannitol agar, 11
sodium lactate gelatin, 27
starch agar, 14
Autotrophic bacteria, 2
Azotobacter colonies, 115
growth on different carbohydrates,
117
Azotobacter colonies, in soil, 114
isolation of, 114
nitrogen fixation by, 116
B
Bacillus celluloses dissolvens, 135
cereus, 128
Bacteria in soil, 92
number in soil, plate method, 99
Bacterium fluorescens, 128
hartlebii, 122
pyocyaneum, 122
radiobacter, 116
Bacteroid formation, 34
Barium hydroxide solution standard,
56
Barlow stain, 51, 109
Bean extract, 34
Beef extract agar, 8
Beef extract gelatin, 8
Black finish for tables, 91
Bouillon, 8
Brain, sheep or beef, 43
Bristol's medium for algae, 21
Brom cresol purple milk, 39
Brucine reagent, 59
Bryan's medium, 45
Burri's PeUkan Tusche, 49
Caffein bean extract, 34
Calcium hypochlorite, seed sterihza-
tion, 83
Capsule stain, 49
Carbohydrates, determination of,
69-76
Carbohydrates, effect on nitrogen
fixation, 120
141
142
LABORATORY MANUAL OF MICROBIOLOGY
Carbol fuchsin, 47
thionin (Nicolle's), 48
Carbon dioxide, determination of,
79, 80
from soil, 136
total, 80
Carrot extract agar, 33
agar for yeast, 18
Casein hydrolysis, 42
Caseinate agar, 9
Cellulose agar, 37
bacteria, media for, 35
determination of, 69
decomposing bacteria,
isolation of, 135
thermophilic, 134
decomposition
aerobic, 134
Characteristics of root nodule bac-
teria, 108
of soil population, 92
Chemosynthetic bacteria, 2
Cleaning solution, 90
Clostridium pasteurianu7n, 101, 114,
118
Clostridium thermocellum, 134
Clover sucrose agar, 15
Cochineal, 52
Color change of indicators, 53
Congo red, method for examining
bacteria, 96
negative stain, 51
Corn meal medium, 43
Crone's medium, 45
Crystal violet, ammonium oxalate,
48
Culture media, 1
Czapek's solution, 13
D
Davisson-Parsons method for nitro-
gen determination, 68
Denitrification by pure cultures, 121
in soil, 122
Denitrifying bacteria, isolation of,
121
media for, 25
Depth, effect on number, 102
Detmer's solution for algae, 20
Diphenylamine reagent, 59
Direct method of measuring number
of bacteria, 104
Directions for media, 3
Dorner capsule stain, 49
spore stain, 50
of nitrogen fixing, bacteria, 119
Embedding nodules. 111
Enrichment media, 1
Erythrosin, 49
method of staining soil bacteria,
51
Ethyl alcohol solution, 25
Exercises in soil microbiology, 87
Fat decomposition, 41
Ferric ammonium citrate, 43
Ferrous carbonate medium, 32
Filtration of culture media, 4
Flemming's stain, 112
Frazier's gelatin, 42
Fungi, culture media, 12
in soil, 93, 98
Gelatin, chemical composition, 5
general properties, 5
liquefaction, 42
Gelidium corneum, 5
Giltay's medium, 25
Glucose phosphate, nitrogen-free, 1 1
Glycerol agar, 129
Gram stain, 48
Green plants, culture media, 44
Gum formation, 35
H
Hanging-drop preparations, 95
Hansen's solution, 16
Hay infusion for protozoa, 19
INDEX
143
Hayduck's medium, 18
Heidenhains hsematoxylin, 112
Heterotrophic bacteria, 2
Heyden agar, 9
Humus, determination of, 78
Hydrogen bacteria, medium for, 30
sulphide bacteria, 139
Bavendamm, 30
Hydroxylamine, test for, 58
I
Indicators, 53
Indol, test for, 61
Ink method, capsule stain, 49
Insects, number in soil, 105
Invertebrate population of soil, 94
Iron bacteria, 133
oxidizing bacteria, media for, 31
precipitating bacteria, 132
Isolation of anaerobic bacteria, 118
Mannitol, nitrogen free, 32
phosphate solution, 20
soil extract, 20
Manure, effect on number of micro-
orgamisms, 103
Media, 1
Mercuric chloride, disinfectant solu-
tion, 90
seed steriUzation, 82
Methane bacteria, medium for,
31
Methods of staining, 47
Methyl orange, 52
red, 67
Methylene blue as indicator of
anaerobiosis, 46
Microscopic examination of soil, 104,
105
Milk, culture media, 39
brom cresol purple, 39
Moisture determination, 61
K
N
Koch, 1
Koser's medium for colon-aerogenes
group, 43
Krainsky's medium, 13
Laboratories, supplying cultures, 139
Laboratory rules, 89
Lagerberg's spore stain, 50
Leguminous plants, bacteria of, 106
Lignins, 75
Literature, 137
Litmus, 52
milk, 39
Loeffler's alkahne methylene blue, 47
Lugol's iodine, 48
M
Malt extract agar, 16
Manganese, oxidizing
media for, 31
bacteria,
Nahrstoff-Heyden agar, 9
Negative stain, 109
Nematodes, number in soil, 103
Nessler's reagent, 56
Nesslerization of ammonia, 62
Nigrosin negative stain, 50
Nitrate formation, 123
reducing bacteria, media for, 25
Nitrates and nitrogen fixation, 120
determination of (reduction), 65
quantitative determination of, 63
reagent, 57, 59
Nitrification in Hquid cultures, 123,
124
of various substances, 125
Nitrifying bacteria, enrichment cul-
tures, 123
media for, 22
Nitrite formation, 123
Nitrites, test for, 57
Nitrobacter, 125
solution for, 23
144
LABORATORY MANUAL OF MICROBIOLOGY
Nitrogen, determination of, 65
fixation in soil, 119
fixing capacity of soil
Winogradsky's method, 119
fixation by cultures of
Azotobacter, 116
bacteria, media for, 32
Nodule bacteria, isolation of, 106
and nitrogen content of plants,
113
in agar, 109
Normal sodium hydroxide, 55
sulphuric acid, 54
Nutrient agar, 8
broth, 8
gelatin, 8
Nutrition of bacteria, 1
Nutrose agar, 9
Oxalic acid, standard, 55
Oxidation of ammonia, 23
sulphur, 131
Paraffin agar, 41
Pasteur, 1
Pea extract, 34
Pentosans, 70
Peptone malt agar, 15
mannitol solution, 12
sucrose, 35
Phenoldisulphonic acid, 60
Phenolphthalein, 52
Physiological groups, 104
Plant material, analysis of, 76
Potassium nitrate, thiosulphate solu-
tion, 26
Potato culture media or potato
slants, 39
glucose agar, 14
Preparation of H ion indicators, 53
Preservation of stock cultures, 45
Preserving plates, 45
plate cultures, 46
Pressure and temperature, 7
Protein decomposition, 128
Protozoa, 94
media for, 19
number in soil, 98
Pyrogallic acid for absorbing oxygen,
46
R
Raisin extract, 17
Raulin's solution, 12
Reaction, 3
Reagents, 52
Reducing sugars, 72
Reduction of sulphates, 26
Rock phosphate, 131
Root, effect on number of bacteria,
102
nodule bacteria, isolation, 107
structure of. 111
Rose bengal, 96
S
Safranin, 48
Schweitzer's reagent, 69
Season, effect on number, 102
Sectioning nodules. 111
Seed sterihzation, 82
Selective media, 1
Shaffer and Hartman modified, 72
Silica gel, general properties, 5
preparation of, 6
Souleyre, 35
use of, 116
SiUcic acid gel for nitrite forming
bacteria, 24
Size of organisms in soil, 94
Sodium albuminate agar, 9
asparaginate, glycerol agar, 19
caseinate agar, 9
citrate (Koser), 43
Soil extract agar, 10
gelatin, 10
stock solution, 10
population, 92
samples, how taken, 97
Sorenson's phosphate (Na2HP04.-
2H2O), 40
Souleyre, silica gel, 35
INDEX
145
Spirochaeta cytophaga, 134
Spore-forming bacteria, determina-
tion of, 101
stain, 50
yeast medium, 18
Staining of bacteria, 47
Starches, 71
Sterilization of media, 6
of soil, 7
Straw, effect of on numbers of
microorganisms, 103
Structure of the root nodule, 111
Suction filter, 4
Sulphates, reduction of, 128
Sulphur bacteria, 130
high forms, 130
oxidizing bacteria, media for, 28
Sulphuric acid, normal solution, 54
Transferring cultures, 90
Trommsdorf reagent, 57
Tubeuf's medium, 15
Truffaut and Bezssonoff medium, 12
U
Urea bacteria, isolation of, 127
media for, 21
citrate solution, 21
decomposition, 136
soil extract solution, 22
Vegetable tissue for absorbing oxy
gen, 46
W
Tap-water gelatin, 11
Temperature and pressure, 7
Test for ammonia, 56
for ferric iron, 133
for hydroxylamine, 58
for indol, 61
for nitrates, 58
for nitrites, 57
ThermophiUc bacteria, 38, 101, 134
Thiohacillus thioparus, 131
thiooxidans, 132
Thionin, 48
Thiosulphate solution, 28
Thomas' test reagents, 57
Thornton's medium, 11
Total carbon, determination of, 80
nitrogen (Davisson-Parsons), 68
(Kjeldahl), 65
to include nitrates, 67
Worms, number in soil, 105
Washed agar for nitrifying bacteria,
23
Water-holding capacity of soil, 61
Winogradsky's direct count, 105
glucose-peptone agar, 12
medium, 11
solution for Nitrosomonas, 22
Yeasts and yeast spores, 96
infusion (dried), 17
infusion (fresh), 16
media for, 16
water sulphite agar, 27
Ziehl's carbol fuchsin, 47
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