METHODS IN MICROBIOLOGY,VOLUME 3B, Volume 3B (v. 3B)

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METHODS in MICROBIOLOGY

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METHODS in MICROBIOLOGY Edited by J. R. NORRIS Milstead Laboratory of Cliem’cal Enzyiiiology, Sittingbourne, k‘ent, England

D. W.RIBBONS Department of Bioclieinistry, University of Miami School of iIIedicine, and Howard Hughes Medical Institute, Miami, Florida, C7.S.A.

Volume 3B

@

ACADEMIC PRESS, INC.

( I izirr.ourt Brare J o v o i i o v i c ~ h Puldishrrs) .

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ACADEMIC PRESS INC. (LONDON) LTD 24-28 Oval Road London NW1

U.S.Edition published by ACADEMIC PRESS, INC.

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Copyright 01969 By ACADEMIC PRESS INC. (LONDON) L T D

All Rights Reserved No part of this book may be reproduced in any form by photostat, microfilm, or any other means, without written permission from the publishers

Library of Congress Catalog Card Number: 68-57745 SBN: 12-521543-6

PRINTED INTHE UNITEDSTATESOFAMERICA

85 86 87 88

9 8 7 6 5 4

LIST OF CONTRIBUTORS ELLAM. BARNES,Food Research Institute, Norwich, Norfolk, England EVEBILLING,East Mulling Research Station, Maidstone, Kent, England T. D. BROCK,Department of Microbiology, Indiana University, Btoomington, Indiana, U.S.A. N . G. CARR,Department of Biochemistry, University of Liverpool, England VERAG. COLLINS,Freshwater Biological Association, Ambleside, Westmorland, England M . R. DROOP, Scottish Marine Biological Association, Oban, Scotland R. J . FALLON,Ruchill Hospital, Glasgow, Scotland N . E. GIBBONS,Division of Bwsciences, National Research Council, Ottawa, Canada P. N. HOBSON,The Rowett Research Institute, Bucksburn, Aberdeen, Scotland R. E. HUNGATE,Department of Bacteriology, University of California, Davis, California, U.S.A. JOHN E. PETERSON, Department of Botany, University of Missouri, Columbia, Missouri, U S .A. A. H . ROSE,School of Biological Sciences, Bath University, Bath, England P. WHITTLESTONE, School of Veterinary Medicine, University of Cambridge, England A. T . WILLIS,Public Health Laboratory Service, Luton and Dunstable Hospital, Luton, Beds., England

V

ACKNOWLEDGMENTS For permission to reproduce, in whole or in part, certain figures and diagrams we are grateful to the following publishersMessrs Baird & Tatlock (London) Ltd; Council of the Marine Biological Association of the United Kingdom; Gustav Fischer, Stuttgart; H. K. Lewis & Company Ltd; Masson et Cie, Paris; Royal Society of Sciences of Uppsala, Sweden. Detailed acknowledgments are given in the legends to figures.

vi

PREFACE Volume 3 of “Methods in Microbiology” is concerned with the techniques used for isolating, growing and preserving micro-organisms, We considered that information on these themes was required in two distinct forms : a comprehensive list of growth media which would provide the reader with easy access to formulae and growth conditions for a wide range of microorganisms, and detailed descriptions of the special methods used for certain selected groups of micro-organisms. In addition general articles describing the principles involved in enrichment techniques for different types of micro-organisms and for the isolation of mutants and the design of mutation/selection programmes are also relevant to the main theme. As the contributions to Volume 3 took shape it became apparent that the amount of material involved was too much for inclusion in one volume and the material split relatively easily into two sub-volumes which are called Volumes 3A and 3B. Volume 3A contains Chapters concerned with the composition of growth media and media tables. Tabulated information about the preservation of micro-organisms and general articles concerned with enrichment, mutation and strain selection procedures are also provided. Volume 3B deals entirely with selected groups of micro-organisms, the emphasis being on methods of isolation,growth and handling in the laboratory, and preservation of cultures. In selecting the particular groups described we have been concerned to choose organisms which are not well described in other publications or which involve, because of their unusual physiology, special techniques. The actual treatment of the material we have left very largely to the choice of the individual authors. Our aim throughout has been to provide a useful treatment of important topics which are not well covered elsewhere while at the same time avoiding pointless repetition of readily available information.

J. R. NORRIS D. W. RIBBONS October, 1969

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CONTENTS LISTOF CONTRIBUTORS.

V

.ACKNOWLEDGMENTS.

vi

PREFACE

.

vii

Chapter I. Isolation, Cultivation and Maintenance of AutotrophsVERAG. COLLINS .

1

Chapter 11. Growth of Phototrophic Bacteria and Blue-Green Algae-N. G. CARR .

53

Chapter 111. Techniques for the Study of Anaerobic, Spore-forming Bacteria-A. T. WILLIS .

79

Chapter IV. A Roll Tube Method for Cultivation of Strict Anaerobes-R. E. HUNGATE .

117

Chapter V. Rumen Bacteria-P.

N. HOBSON

.

Chapter VI. Methods for the Gram-negative Anaerobes-ELLA M. BARNES . Chapter VII. Psychrophiles and Thermophiles--T. A. II. ROSE

133 Non-sporing

151

D. BROCKAND 161

Chapter VIII. Isolation, Growth and Requirements of Halophilic Bacteria-N. E. GIBBONS .

169

Chapter IX. Isolation, Cultivation and Maintenance of the Myxobacteria-Jol%N E. PETERSON .

185

Chapter X. Isolation, Cultivation and Maintenance of Mycoplasmas -R. J. FALLON AND P. WHITTLESTONE .

211

Chapter XI. Algae-hl.

K. DROOP .

Chaptcr M I . Isolation, Growth and Preservation of Bacteriophages -EVE BILLING . ,

269 315

AUTHORINDEX

.

331

SVBJLCT INDEX

.

345

ix

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Isolation, Cultivation and Maintenance of Autotrophs VERAG. COLLINS Freshwater Biological Association, Anibleside, Westmorland, England I. The Environmental Site in Nature

.

1

.

4

111. Isolation and Cultivation of Some Representatives of the Various Groups of Organisms . .

12

IV. Discussion on the Autotrophic Way of Life in the Natural Environment

25

11. Groups of Autotrophic and Facultatively Autotrophic Organisms

V. Media and Methods References

.

.

. .

26 49

I. T H E ENVIRONMENTAL SITE I N NATURE A. Stratified freshwater lake A stratified freshwater lake offers an ideal natural site for the isolation of facultatively autotrophic bacteria. T h e period of stratification of a lake starts in the late spring and early summer, when the temperature of the surface water rises owing to increased solar radiation. T h e resulting difference in density of the water gives rise to the formation of two distinct layers of water, the upper layer or “epilimnion” and the lower layer or “hypolimnion”. Under some conditions the temperature is approximately the same at all depths in the epilimnion; below this layer, in the transitional zone known as the “thermocline”, the temperature falls rapidly with increasing depth. The nature of the thermocline and the dcpth at which it occurs vary greatly, but in general the depth tends to increase as the summer advances and, therefore, the ratio of the volume of epilimnion to volume of hypolimnion increases. When the layering has begun, the amount of dissolved oxygen becomes gradually less in thc hypolimnion owing to the stagnation of the water and the increased activity of the aerobic bacterial flora of thc mud surface (Collins, 1963). l’his incrcascd activity of thc hcterotrophic facultatively 2

2

V. G . COLLINS

anaerobic bacteria and the mud's chemical oxygen demand results in an oxygen deficit in the bottom waters of a stratified lake. When this situation prevails, the first group of autotrophic bacteria to become active are the sulphate reducers. This results in the production of hydrogen sulphide and the formation of ferrous sulphide on the mud surface and in the overlying Bacteria, 1000/ml

!32527 29 31

3 (C)

i I

3-

1

A

I

I

..

4-

,

.

5-

I E

6-

'.

f a 0) 7 n

C

Temperoture, "C 17 18 19 2 0 21 ?I 22 2? 2 3

10

20

jb

I

30 40 I 60 6b Oxygen, % saturation

I

I

70

I

I

80

FIG.1. A typical environmental situation for a stratified lake. Bfelham Tarn, Boathouse Buoy, August 20th, 1968. Curves: (a) bacteria; (b) temperature; ( c ) oxygen. A. epilimnion; B thermocline ; (c) hypolimnion.

water, creating ti zone of anaerobiosis. At the interface between this zone and the upper zone containing oxygen, members of the coloured sulphur bacteria develop where the light penetration is sufficient to support their photosynthetic mechanisms. Above this zone in the lake and therefore in the thermocline system, where there is sufficient oxygen and organic matter in the form of decayed plank-

I . AUTOTROPHS

3

tonic organisms, members of the photosynthetic purple non-sulphur bacteria develop. Within this system, during stratification, the activities of the filamentous iron organisms must be superimposed. They are not recovered from the oxygenated water immediately above the mud surface, but they are present in very large numbers in the thermocline and above it, and offer, by means of their filamentous growth, a matrix of colloidal sheaths impregnated with iron compounds-an ideal site for the attachment of other micro-organisms. A typical environmental situation for a stratified lake is shown in Fig. 1, where the temperature and dissolved-oxygen determinations were measured by means of a Mackereth oxygen electrode (Mackereth, 1964). This instrument can be obtained from The Lakes Instrument Co., Ltd., Oakland, Windermere, Westmorland. T h e results for bacteria per ml were obtained from aerobic plate counts on a standard medium (Collins and Willoughby, 1962).

B. Sampling methods 1. Water samples For the purpose of obtaining water samples from a profile depth series in any lake system, a piece of apparatus known as a Friedinger water bottle is invaluable. These bottles can be purchased from Messrs Hans Buchi, Berne, Switzerland, and are made in two capacities, 1 litre and 2 litre. They can be sterilized, if necessary, between different depth samples by alcohol swabbing. If it is essential to obtain samples of water at a given depth in the sampling profile without any risk of contaminating water from any other depth entering the sampler, then the bacteriological sampler designed by Mortimer (1940)is recommended.

2. Mud samples Samples of mud can be obtained by means of a Jenkin surface-mud sampler, Fig. 2. ‘I’his apparatus can be obtained from The Lakes Instrument Co., Ltd., Oakland, Windermere, Westmorland. The sampler takes a mud core approximately 30 cm in depth, with the ovcrlying water in situ ovcr the mud. l’here is some slight disturbance of the surface mud during its enclosure in the Perspcx sampler tube, but not enough to make any significant difference to the contents of the sample. 1he samples, collcctcd by means of this apparatus, are excellent for the purpose of making Winogradsky cylinders (Winogradsky, 1887) for enrichment culture techniques, for measuring redox potentials of mud, and for respiration studies using pcrfuscr methods. r ,

4

V. G . COLLINS

11. GROUPS OF AU'L'O'rROI'HIC AND FACULTATIVELY AUTOTROPHIC ORGANISMS

A. Group a: Ammonia-oxidizing bacteria ORDER I I-'SElJDOMONADALES Rergey (1957) SUBORDER I I €'SEIJI)OMONAI)INEAl~ Family I NITl~OiIA(:TIiIlh~'fiAE Genus I Nitrosomonas

FIG. 2. Jenkin surface-mud sampler. (a) The sampler; (b) empty core tube mounted on the sampler; (c) mud-core sample. Photograph by A. E. Ramsbottom.

5

I. AUTOTHOPHS

B. Group b: nitrite-oxidizing bacteria ORDER I PSEUDOMONADALES Bergey (1957) SUBORDER I1 PSEUDOMONADINEAE Family I NITROBACTERACEAE Genus VI Nitrobacter

C. Group c : sulphur-oxidizing bacteria, that oxidize inorganic sulphur compounds and deposit sulphur globules internally ORDER Family Genus Genus Genus ORDER Family Genus

VII I I 111 IV VII IV I

BEGGIATOALES BEGCIATOACEAE Beggiatoa Thioploca Thiothrix REGGIATOALES ACHROMATIACEAE Achromatium

Bergey (1957)

Bergey (1957)

For illustration of representative species of Group c, see Fig. 3. D. Group d : sulphur-oxidizing bacteria that oxidize inorganic sulphur compounds and deposit sulphur both internally and externally ORDER SUBORDER Family Genus Genus Genus Genus

I I1 I11 I1 I11 IV V

PSEUDOMONADALES Bergey (1957) PSEUDOMONADINEAE

THIOBACTERIACEAE Macromonas Thiovulum Thiospira Thiobacillus

For illustrations of representative species of Group d, see Fig. 4. E. Group e: photosynthetic sulphur bacteria, producing red to

purple pigments, and able to use hydrogen sulphide as a hydrogen donor ORDER ' I SUBORDER I Family I Genus II Genus III Genus IV Genus V Genus VI Genus VII Genus VI I I Genus IX Genus X Genus XI Genus XI I Genus XI11

PSEUDOMONADALES Bergey (1957) RH~DOBACTERIINEAE

THIORHODACEAE Thiopedia Thiocapsa Thiodictyon Thiothece Thiocystis Lamprocystis Amoebobacter Thiopolycoccits Tiiiospirilluni Rhabdomonas Rhodothrce Chrowatittm

For illustrations of some rcprescntativc species of Group e, see Figs 5 and 6.

FIG.3. Group C : illustrations of some representative members of this microbial group. 1. Beggiatoa mirabilis (from Bavendamm, 1924), x 115 ; 2. Beggiatoa arachnoidea (from Skuja, 1956), x 670; 3. Beggiatoa minima (from Bavendamm, 1924), x 450; 4. Beggiatoa alba (from Bavendamm, 1924) x 450; 5. Thiothrix nivea (from ; Thiothrix niveu, growing attached to an algal cell Bavendamm, 1924), ~ 4 5 0 6. (from Bavendamm, 1924), x 67; 7. Thiothrix annulata (from Bavendamm, 1924), x 125; 8. Thiothrix tenuis (from Winogradsky, 1949), x 50; 9. Thioploca ingrica (from Bavendamm, 1924), x 225; 10. Achromatium oxuliferum, cells about to divide, original; 11. Achromatium oxalz’ferum, internal granules, original ; 12. Achromatium mobile (from Bavendamm, 1924), ~ 3 5 5 .T h e diagrams from Ravendamm are reproduced by courtesy of Gustav Fischer, Stuttgart; that from Skuja b y courtesy of the Royal Society of Sciences of Uppsala, Sweden; that from Winogradsky by courtesy of Masson et Cie, Paris.

I. AIJTOTHOPNS

7

Fig. 4. Group d : illustrations of some.representative members of this microbial group. 1 . Macronionas mobilis (from Skuja, 1956), x 840; 2. Macromonas hipiinctata (from Skuja, 1956), x 804; 3. Marvomonas frtstyormis (from Skuja, 19ih), x 804; 4. Mucromonas minutissiwta (from Skuja, 1956), x 804; 5 . Tltin7:zthtm ntinrrs (from Skuja, 1956), x 513; 6. Thiovitlum nrajiis (from Skuja, 1956), x 513; 7. Thinzw/rim Mttlleri (from Skuja, 1956) x 5 1 3 ; 8. Thiospira agilis (from Skuja, 19.56), x 804; 9. Thiospira Winogradskzi (from Bavendamm, 1924), x 420; 10. ‘I%io.ipira rfcstogyru (from Skuja, 1956), x 804; 11. 7hiospira tanrii.r (from Skuja, 1956), x 804; 12. Tlliospira Winogradshy (from Skuja, 1056), x 804. T h e diagrams frcim Skuja arc reproduced b y courtcsy of the Iioyal Society I J f Sciences of Upps;rl;i, Swedcn ; that from naveridamm by courtcsy of (iustav I:ischcr, Stuttgatt.

FIG.5 . Group e : illustrations of some representative members of this microbial group. 1. Thiopedia rosea (from Bavendamm, 1924), x 560; 2 . Thiocapsa Y O S ~ O pevsicina (from Bavendamm, 1924), x 486 ; 3. Thiodictyon elegans (from Ravendamm, 1924), x 560; 4. Thiothece gehtinosa (from Bavendamm, 1924), x 5 6 0 ; 5. Thiocystis violarea (from Bavendamm, 1924), x 560; 6. Lamprocystis roseo-persicina (from Bavendamm, 1924), ( a ) x 135 ; (b) x 486; 7. Lamprocystis roseo-persicina (from Winogradsky, 1949), schematic ; 8. Amoebobacter roseum (from Winogradsky, 1949), (a) colony of cells, (b) cells prior to colony formation; schematic; 9. Amoebobacter bacillosiis (from Winogradsky, 1949), schematic ; 10. Thiopolycoccus rubm (from Ravendamm, l924), x 56; 11. lihodotliccr ntrda (from Skuja, 1956), x 804; 12. Rhodothece conspicira (from Skuja, 19-50), x 804. The diagrams from Bavendamm are reproduced by courtesy of Gustav Fischer, Stuttgart; those from Winogradsky by courtesy of Masson e t Cie, Paris; those from Skuja by courtesy of the Royal Society of Sciences of Uppsala, Sweden.

Fla. 6. Group c : illustrations o f some representative mcrnbers of this microbial group. 1 . Thiospiuillrcm Hosenberfiii (from Skuja, 1950), x 804 ; 2. 7*l~io.spi~ill1c~~~ jeizcnsc (fn,m Skuja, 1056), x 804; 3. H / r u / ) ~ f ~ / c . / / / . c t i i(R/rnhcfoirronu.s) /u~;~/i/~ ro.sfw/i (from Skiija), x 804; 4. R/rabdoc.lrrowrutiriiir Sp. (from I%;iventlarnm,1924), involution forms, x 67.5 ; 5 . Rlrahdoiiroirus S'p., original ; 6. I~l/cihtfoc.lcri)iiintirc?ir (Rlrabdomonas) frrsifornre (from Ih\-cmtlamm, 1024) ; 7. ~ ' / ~ r o i i r o t ; r c.Sp., ~ u original ; 8. Clruoviatirem Sp., intcrml glol~tilc~s, original. 'I'hc diiigranis f r o m Skuja arc rcproduccd by courtesy of thc R ( J ~ : Socicty II o f Scicnccs i)t' Lrpps:il:i, S\vedcn ; those from I h \ endamm by courtcsy o f (; u s t : i \ , I;ischcr, Stuttgart.

10

v.

G . COLLISS

FIG.7. Group f : illustrations of some representative members of this microbial group. 1. Clathrochloris hypolimnica (from Skuja, 1956), x 804; 2. Chlorochromatium glebultcm (from Skuja, 1956), x 804; 3. Pelodictyon Sp., original; 4. Cylindrogloea bacterifera (from Skuja, 1956), x 804; 5. Chlorobiztm Sp., enrichment culture, original. T h e diagrams from Skuja are reproduced by courtesy of the Royal Society of Sciences of Uppsala, Sweden.

11

I . AUTOTHOPHS

F. Group f: photosynthetic sulphur bacteria, producing green pigments and able to use hydrogen sulphide as a hydrogen donor ORDER SUBORDER Family Genus Genus Genus Genus Genus

I I 111 I II III V VI

PSEUDOMONADALES Bergey (1957) RHODOBACTERIINEAE C'IfLORODACTERIACEAE

Chlorobirim Pelodictyon Clathrochloris Cl~lorochrornatirini Cylindrogloea

For illustrations of some representative species of Group f, see Fig. 7 .

G. Group g : photosynthetic, non-sulphur bacteria, producing red to brown pigments ORDER SUBORDER Family Genus Genus

I II I II

PSEUDOMONADALES Bergey (1957) RHODOBACTERIINEAE ATZIIORHODACEAE

Hhodopseudomonas Rhodospirillum

H. Group h: bacteria capable of reducing sulphates, and able to obtain their energy from atoms of hydrogen or the labile hydrogen atoms of low molecular organic substances ORDER I SUBORDER I I VII Family Genus II

PSEUDOMONADALES Bergey (1957) PSEIJDOMONADINEAE SPIRILLACEAE Desulfovibrio

I. Group i :methane-oxidizing bacteria ORDER I SUBORDER I1 Family I1 G~I~LIS I

PSIXJDOMONADALIIS Uergey (1957) PSEUDOMON~DINEAE METHANOMONADACEAE Alethanomonas

J. Groupj :hydrogen-oxidizing bacteria ORDER SUBORDI:R Family Genus

I1 I1 II II

PSEUDOMONADAIXS I3ergey (1957) PSEUDOMONADINEAE METHANOMONAIMCEAE Hydrogenomonas

K. Group k: carbon monoxide-oxidizing bacteria ORDER I SUBORDICR I I Family II Genus III

PSEUD0MONADALI:S PSEUDOM0NAT)INEAE R.~ET€1ANOMONAI)A('I:i\E

Carbosyc-iomonas

Uergey (I 9.57)

v.

12

G . COLI.INS

L. Group 1: bacteria capable of oxidizing iron and manganese compounds ORDER

I1 CIHLAMYDOBACTERIALES Bergey (1957)

Family Genus Genus

I Splraerotihrs I1 Leptotkrix

Family Genus Genus

I1 PELOPLOCACEAE I Pcloploca I I I’cloncma

Family (;enus (;cnus

ORDER SUBORDER Family Genus Famiiy Genus Genus Genus Genus Genus Genus Genus

1 CI-ILAMYDODACTERIACEAE

I I I CI3ENOT RICI3ACI:AE I C‘renotlirix I 1 I CIonotlirix I PSEUDOMONADALES I1 PSEUDOMONADINEAE

Bergey (1957)

V CAULOBACTEHACEAE

II Gallionella

VI SIDEROCAPSACEAE I IV V VI VI I IX

x

Siderocupsa Ferribacterium Sideromonas Naumanniella Ochrobium Siderobacter Ferrobacillus

For illustrations of some representative species of group 1, see Figs 8, 9 and 10. 111. ISOLATION AND CULTIVATION OF SOME REPRESENTATIVES OF THE VARIOUS GROUPS 01; ORGANISMS

A. Group a ; ammonia-oxidizing bacteria, Nitrosotnonus species 1. Site These organisms can be isolated from the surface muds of stratified lakes, where the mud is covered with shallow layers of water within the oxygenated zone. An ideal recovery site in a freshwater lake is the surface mud where a sewage bearing inflow first flows into the lake.

2. Methods Samples of the surface mud are obtained by the use of the Jenkin surfacemud sampler. T h e surface layer of the mud core thus obtained is removed aseptically and transferred to a sterile, covered 500 ml beaker. Sterilized CaC03 is added in sufficient quantity to cover the layer of mud, the beaker

I . AUTOTROPHS

13

and contents then being incphatcd overnight at 20°C. Enrichment cultures are then made, kiftcr miling thc contents of the beaker, by inoculation of 5 g amounts of the mud with l’opc and Skcrman’s liquid mineral salts medium with additions (Slterinan, 1967). Alternatively inoculations can

FIG.8. Group c : illustrations ( I f some represcntati\,e membcrs of this microbial group, also shown in Figs 9 and 10.1. Spiiaerotilits natans, original; 2. Leptotlirix Sp., original; 3 . Leptothrix ochracea, a chain of rods gliding out of a sheath impregnated with iron, original; 4.Leptotlirix ochracea, a chain of rods completely emerged from an iron impregnated sheath, original ; 5 . I,eptot/irix sidevopozrs, Filamcnts growing out of an iron impregnated attachment disc, original ; 6. Crrnothrix polyspora (from Dorff, 1934), x 750; 7 . Clonothrixfitsca (from Uorff, 1934), x 500. ‘I’he diagrams from Dorff are reproduccd by courtesy o f Gustav I:ischcr, Stuttgart.

14

FIG.9. G r o u p 1 : 1. Pelonenia tenur (from Skuja, 1956), (a) x 600, ( t i ) x 402; 2 . Peloplocaferruginea (from Skuja, 1956), (a) x 810, (1)) x 600, ( c ) x 2 8 2 ; 3. Gallionella ferruginea (from Do&, 1934), (a) x 600, (b) x 1440; 4. (;ulliondla tenuicaulis (from Skuja, 1956), x 804; 5. Gallionella tcniiicaulzs, showing the “kidney cells” a t the ribbon terminals (from Skuja, 1956), x 804. T h e diagrams from Skuja arc reproduced by courtesy of the Royal Society of Sciences of Uppala, Sweden; that from Dorff by courtesy of Gustav Fischer, Stuttgart.

I. AUTOTROPIIS

15

FIG.10. Group 1 : 1. Sicferorapsa Treitbii (from Dorff, 1934). schematic; 2. Siderocupsa major (from Dorff, 1934), schematic; 3. Siderorapsa coronata (from Dorff, 1934), schematic; 4. Siderocupsa geniinata (from Skuja, 1956), x 670; 5 . Sideyoderma (Ferribacterium)dubiicm (from Skuja, 1956), x 670; 6. Naunianniella neustonica. (from Dorff, 19341, schematic; 7. Sideyocystis (Sideromonas) conjercurum (from Dorff, 1934), x 500; 8 Sidcrobacter (from Ihrff, 1934), (a) Sidmhacter lineare, (b) Siderobucter di+/ex, both schematic; 9. Ochrobiun/ tecfitnt (from I h r f f , 1934), x 1000; 10. Ochrobium techtni (from Skuja, 1956), x 670. T h e diagrams from I>orf€ are reproduced by courtesy of Gustav E’ischer, Stuttgart; those from Sltuja b y courtesy of the Royal Society of Sciences of Uppsala, Sweden.

16

v. c. COLLINS

bc niaJc into the medium of Meililcjohn (Skerman, 1967, p. 218). Subsequent transfcrs are inadc to silicd-gel plates, with nutrient additions, using the method of Sonimcrs and IIarris (1968) for preparation of the silica gel. When colonies develop on the siliea-gel plates, they are then transferred back to the above liquid media for A’itrosomonas, with the addition of a shallow layer of sterile fine sand on the bottom of the culture flask. Purity checks on the flask contents can bc performed by the inoculation of ordinary nutrient agar plates. As growth of the pure culture proceeds, estimations of the accumulation of nitrite (Skerman, 1967, pp. 218-220) and the absence of growth on repeated purity checks on nutrient agar give clear evidence of the presence of Nitrosonionas species. Re-plating on silica-gel plates, and subsequent transfer of pure colonies back to liquid media, with sand, should give pure cultures. T h e medium and methods of Soriano and Walker (1968) are greatly to be recommended.

B. Group b: nitrite-oxidizing bacteria, Nitrobacter species 1. Site The environmental site in a freshwater lakc is the same as for Group (a), as are the sampling operations.

2. Method T h e mud for enrichment cultures of Croup (b) is inoculated into Skerman’s medium for Nitrobacter (Skerman, 1967, pp. 215, 216) and into Meiklejohn’s medium (Skerman, 1967, p. 218). Again purity checks can be made on nutrient agar plates. Cultures of the nitrite-oxidizing bacteria are maintained in the liquid medium. As growth proceeds, determinations of the disappearance of nitrite and the formation of nitrate (Skerman, 1967, p. 218) in the culture flask, combined with the absence of growth on the nutrient agar purity check plates, give positive evidence of the presence of Nitrobacter species. For pure culture studies and literature reviews relating to the organisms of Groups (a) and (b), reference should be made to the follouing pubiications; hleiklejohn (1950; 1952; 1953a; 1953b; 1954), I,ws (19S4), and Smith and Iloare (1968).

C. Group c :sulphur-oxidizingbacteria that are capable of oxidizing inorganic sulphur compounds, depositing sulphur as globules within their cells-Beggiatoa, Thioploca, Thiothrix and Achromatium 1. Site T h e representatives of this group are found on the surface muds of stratified lakes, just prior to the complete de-oxygenation of the hypolimnion

I. AUTOTROPI-IS

17

water when the dissolved oxygen concentration is in the range of 0*15-0*30 mg &/litre in the deepest part of the lake. I n the English Lake District, this oxygen depletion of the hypolimnion occurs in five lakes where the depth ranges from 13 to 21 m.

2. Method The samples of mud are obtained from Jenkin surface-mud cores, taken at the maximum depth of the lake sampling profile. This enables the actual environmental conditions which prevail at the time of sampling to be maintained under laboratory conditions. That is, until the facultative heterotrophicbacteria increase their respiration rate by the increased temperature of the laboratory, and remove the remaining traces of oxygen. Therefore, it is essential for the successful recovery of this group of organisms to make enrichment cultures from the surface mud before the onset of complete de-oxygenation within the Jenkin core-sampling tube. As a procedure to check for the presence or absence of these organisms in the surface mud of the sample, small drops of the surface mud, removed by pipette, can be mounted on microscope slides under cover slips and viewed directly under the microscope, using darkground illumination and a lox objective (Leitz Heine system phase-contrast) or a 25 x phasecontrast objective with bright field phase lighting. T h e unique morphology and size of the organisms of this group makes rapid identification possible. An essential feature of successful enrichment cultures is a slow generation of hydrogen sulphide and a low concentration of dissolved oxygen. One enrichment method consists of removing the top layer of mud from a Jenkin core sample, and adding it directly to a Winogradsky Cylinder (Winogradsky, 1887; Larsen, 1952; Collins, 1963). I t is advantageous to siphon off the overlying water of the original mud core and use this as the liquid for the medium of the Winogradsky cylinders. Air is then slowly bubbled into the top layer of the water in the cylinder of the enrichment culture. Jenkin core tubes act as excellent cylinders. T h e entire unit is then stored in a cool incubator (Gallenkamp) at a temperature of ll-l2”C, this being the temperature of the natural environment of the surface mud at the time of sampling. The majority of the organisms in this group will develop on the surface of the mud and in the zone of water immediately abbe the mud. The slow generation of hydrogen sulphide from the bottom of the cylinder and the slow diffusion of oxygen from the top of the water column in the cylinder enables an artificial “poising” of the oxygen concentration. Insertion of an oxygen probe into the cylinder during incubation enables measurement of the dissolved-oxygen content to be carried out. Control of the rate of oxygenation can then be maintained between 0.15-0*50 mg Oz/litre in the over3

18

V. G. COLLINS

lying water of the cylinder. This type of enrichment culture will yield viable cells of all the organisms of this group, over a period of six to eight weeks. T h e next stage of obtaining pure cultures of these organisms is more difficult. T h e author, so far, has only succeeded in isolating species of Beggiatoa in pure culture using the methods of Faust and Wolfe (1961), and the methods of Cataldi (1940). However, for taxonomic purposes it is possible, from the enrichment culture technique, to harvest large concentrations of the cells of Thioploca, Thiothrix and Achromatium for microscopic examination. For reference purposes the descriptive work of the following authors should be consulted: Keil(l912); West and Griffiths (1913); Bavendamm (1924); Ellis (1932); Winogradsky (1949); Scotten (1953); Bissett and Grace (1954); Pochmann (1959); Lackey (1961); and Burton and Morita (1964).

D. Group d: organisms capable of oxidizing inorganic sulphur and usually depositing sulphur externally-Mucromonus, Thiovulum, Thiospira and Thiobaciilus 1. Site The main source of material for this group is from Jenkin surface-mud cores taken in the oxygenated zone of a stratified lake, that is mud from above the thermocline regime. I n this case the overlying water of the mud would have an oxygen concentration from 70 to 80% saturation, but the surface mud would have avery low oxygen value, of the order of 1-0-2.0 mg Oz/litre. An excellent time to take samples for this group is on the occasions when mass blooms of algae occur in the surface waters of a lake. When the algal cells die and sink onto the surface mud above the thermocline, large populations of Macromonas, Thiovulum, and Thiospira develop on the surface mud.

2. Method The simple expedient of storing the Jenkin mud cores with the top of the tube uncovered, at a temperature of 15"-20"C, usually results in large numbers of the afore-mentioned species developing on the surface of the mud. I n the case of Thiovulum, dense zones of growth develop in the overlying water about half-way between the surface of the tube and the mud. Pure cultures of these three species have as yet not been procured by the author, microscopic studies only have been possible for taxonomic purposes. For enrichment cultures of thiobacilli direct from the natural environment of the surface mud of a freshwater lake, 1 g amounts of mud are inoculated into the medium of Lieske (1912), Waksman and Starkey (1922); Starkey (1953) using tk serial-dilution technique. T h e liquid media in this instance are dispensed in shallow layers in 100 ml Pyrex conical flasks, and sterilized by steaming for one hour on three consecutive days. Two other

I. AUTOTROPHS

19

useful media are those described by Skerman (1967, p. 216), using the PopeSkerrnan basal mineral salts medium with additions. Enrichment cultures are best incubated at 30"C, when turbidity and pellicle formation are evident in the flask contents. A transfer to medium of the same composition with the addition of agar usually results in pure colonies of Thiobacillus species from the higher dilutions of the mud. Subsequent transfer of colony material back to liquid media, accompanied by determinations to assess the disappearance of thiosulphate gives positive evidence of the presence of thiobacilli. For the methods of estimation involved in determining the reactions of these organisms during active growth, see Skerman (1967, pp. 234-238). For studies on the species of Thiobacillus concerned in the oxidation of thiocyanate the work of Happold et al. (1954) should be consulted. The following publications should be used for reference to the ironoxidizing species of Thiobacillus: Lyalikova (1958) ; Colmer (1962); Lazaroff (1963); Razzell and Trussell (1963); and Duncan et al. (1964). The anaerobic thiobacilli have been extensively studied by the following workers: Woolley et al. (1962); London (1963); Hutchinson et al. (1965; 1966; 1967); and Jackson et al. (1968). For general reviews on the organisms of Group (d) the following work should be consulted; Bunker (1951); Baalsrud (1954); Lees (1955); Vishniac and Santer (1957); and Sokolova and Karavaiko (1964).

E. Group e : photosynthetic, sulphur bacteria, producing red to purple pigments-Thiopedia, Thiocapsa, Thiodictyon, Thiothece, Thiocystis, Lamprocystis, A moebobacter, Thiopolycoccus, Thio spirillum, Rhabdomonas, Rhodothece, Chromatium 1. Site The source of material is Jenkin surface-mud cores obtained from the de-oxygenated zone 'underneath the thermocline of a shallow, stratified lake, when the temperature' at the bottom of the lake is in the range of 11"-12°C and the dissolved-oxygen concentration 0.09 mg Ozflitre.

2. Method The original mud-core sample can be used as a Winogradsky cylinder, either stored, closed to laboratory atmosphere while exposed to diffuse daylight, or incubated untouched, at 3OoC, with a series of 25 W light bulbs positioned at a distance of 20-30 cm. The natural generation of hydrogen sulphide proceeds in the mud core, and any remaining oxygen is taken up by the heterotrophic bacteria present on the surface mud under the completely closed conditions of the core tube. When pink pigmented patches appear on the inside walls of the tube,

20

v.

G. COLLINS

enrichment cultures can be made by siphoning off the overlying water of the mud core, the entire core being removed from its tube by means of a piston. Slices of the mud are then placed in Petri dishes, mixed with a glass rod and 5 g amounts transferred to empty sterile 4 oz glass-stoppered bottles. T h e bottles are then completely filled with the medium devised by Pfennig (1961; 1962), and incubated at 30°C with 25 W light bulbs placed at a distance of 20-30 cm from the bottles. As growth of the enrichment cultures proceeds, the contents of the bottles turn a bright pink to deep red in colour. At this stage, pure cultures can be achieved by inoculating 5-10 ml of the enrichment culture into deep-culture tubes of van Niei’s medium (van Niel, 1931). These tubes are constructed from pieces of glass tubing 8 in. in length, and having an internal diameter of $ in. They are plugged at both ends with cotton wool and sterilized. After sterilization a sterile rubber bung is inserted at one end of the tube, the enrichment culture inoculum is added, the cooled molten agar medium is poured in, a sterile rubber bung is inserted in the other end of the tube, and the contents are gently mixed by repeated inversion of the tube. When mixing is complete, the tube is immersed in cold water to allow quick setting of the agar. When the agar has set, the tubes are removed from thc water, dried, and a transparent Viscap sealing cap (Baird and Tatlock, Ltd.) placed over the rubber bung at either end of the tube. For incubation at 30°C the tubes are mounted in a rack to allow maximum length of the tubes to be exposed to the 25 W light source. Brightly pigmented colonies soon develop in the agar, which can be removed by opening the tube and, using a sterile glass piston slightly smaller than the diameter of the tube, pushing the long plug of agar out and slicing it into sterile Petri dishes. Separate colonies are then cut out of the agar slices and transferred back to the medium of Pfennig, as liquid cultures in glass stoppered bottles, for maintenance and sub-culture purposes. For descriptive works with many illustrations see Winogradsky (1 888; 1949); Bavendamm (1924); Skuja (1956). For culture studies see the work of van Niel (1931), and Pfennig (1961; 1962).

F. Group f: photosynthetic, green pigment producing sulphur bacteria-Chlorobium, Pelodictyon, Clathrochloris, Chlorochromatium, Cylindrogloea 1. Site T h e source is the same as that for Group (e). 2. Method For this group enrichment cultures are obtained by the Winogradsky (1887) and Larsen (1952) cylinder method, and exposed to the same conditions as those described for Group (e). Pure cultures are obtained using the

I. AUTOTROPHS

21

methods and medium of Larsen (1952)) or the methods for green sulphur bacteria described by Pfennig (1961 ; 1962). For reference purposes the review by Larsen (1954) covers both groups of photosynthetic sulphur bacteria. T h e isolation by Pfennig (1968) of new species of green sulphur bacteria is proof that the green sulphur group of organisms is far from complete.

G. Group g : photosynthetic, non-sulphur bacteria, producing red to brown pigments-Rhodopseudomonas, Rhodospirillum 1. Site , The source is the same as that for Group (e). T h e bacteria can also be obtained from water samples taken in the thermocline zone of a stratified lake towards the end of the period of stratification in August and September of each season, when the temperature within this zone would be in the range of 17'-19"C, and the dissolved oxygen concentration within the range of 55-90% saturation.

2. Method Enrichment cultures can be obtained from Winogradsky cylinders using the surface mud, and from direct inoculation of the water samples using the methods and medium of van Niel(l94-4). For maintenance of pure cultures of these organisms the methods and media of Hutner (1944; 1946; 1950) give excellent results.

H. Group h: organisms capable of reducing sulphates and able to obtain their energy from atoms of hydrogen or the labile hydrogen atoms of low molecular organic substances. Also able to use low organic substances and C 0 2 as C-sourcesfor growth-Desulfovibrio 1. Site The source is Jenkin surface-mud cores taken from the deepest part of a stratified lake, when the bottom waters of the lake have remained stagnant for the maximum period under the thermocline. This period usually extends from the end of May to the end of September, just before the overturn and complete mixing of the lake.

2. Method The overlying water of the mud-core sample is siphoned off, and the entire core of mud is sliced into Petri dishes and placed directly into an anaerobic jar and incubated at 30°C. After 2 days incubation, sub-samples from the mud in the Petri dishes are inoculated into freshly autoclaved medium that has not been exposed to laboratory atmosphere. The methods and media recommended and used by Butlin et al. (1949) are excellent. These authors modified the original medium of Baars (1930) and Starkey

22

V. G. COLLINS

(1938); their media and methods are described by Skerman (1967, pp. 267268). These media are excellent for enrichment cultures. For serial dilution counts direct from the natural environment, and for pure culture maintenance and experimental work, the medium first described by Miller (1950) and later used by Crossman and Postgate (1953), is to be recommended; this medium is quoted by Skerman (1957, pp. 268-269). It is quite acceptable to omit the incubation of the mud in an anaerobic jar, and to inoculate the slices from the mud core directly into any of the above-mentioned liquid media in glass-stoppered 2 oz bottles. Serial dilutions can be used for enumeration purposes, blackening of the bottles after 7 days incubation at 30°C is a good indication that active sulphate reduction is taking place. Transfer of enrichment culture material to agar media of the same composition incubated in an anaerobic jar usually results in pure colonies of sulphate-reducing bacteria. For pure culture studies see Postgate (1965). T h e culture vessel system designed by Pankhurst (1967) serves as an excellent method of maintaining pure cultures of these organisms for a considerable time period, before sub-culturing is necessary. For studies on the spore-forming sulphate-reducing bacteria reference should be made to the publications of Campbell et al. (1957), Coleman (1960), and Postgate and Campbell (1963). A very good general study of sulphate reducing bacteria is that of Miller et al. (1968). J. Groupj :hydrogen-oxidizingbacteria-Hydrogen0 rnonas 1. Site T h e source is mud cores obtained from the littoral regions of freshwater lakes, especially lakes that receive their drainage from agricultural land which has been fairly heavily manured and also treated with inorganic fertilizers. T h e surface mud, obtained from such an environmental site as this, is always exposed to fully oxygenated overlying water.

2. Method T h e author has tried a number of different methods for the isolation of this group of organisms from the natural environment. For maintenance of autotrophic growth conditions the method of Atkinson and McFadden (1954) should be used. Cohen and Rurris (1955) also describe a very effective method of culturing these organisms in liquid media, which serves as an ideal basis for obtaining enrichment cultures from direct inoculation of the surface mud to the culturing system. Transfer of material to either silica-gel plates or agar, as described by Schatz and Bovell (1952), and Wilson et al. (1953) usually results in the growth of pure colonies of these organisms. For maintenance purposes, cultures survive for a longer period of time under the conditions described by Cohen and Burris (1954).

I. AUTOTROPNS

23

Enrichment cultures of related strains of these organisms have also been recovered from surface mud from the decpcr parts of a stratified lake, when the oxygen saturation has been in the range of 6-15%. Below 6% oxygen saturation of the overlying water of the mud, attempts at isolating this group of organisms have been negative. For information on the autotrophic and heterotrophic growth of this group of organisms, reference should be made to De Cicco and Stukus (1968). For the isolation of pure cultures of this group, under reduced oxygen tension conditions, the medium of Schatz and Bovell(l952) is ideal; this is described by Skerman (1967, p. 220).

K. Group k: carbon monoxide-oxidizing bacteria-Carboxydomonas The author has no personal experience with this group of organisms; repeated attempts to obtain enrichment cultures from the surface mud of a freshwater lake have been negative. On several occasions, using the medium of Kistner (1953), several isolates have been studied and were found to oxidize CO to CO2. However, without exception these isolates also oxidized hydrogen when grown under the conditions described by Cohen and Burris (1954). Kistner’s medium is described by Skerman (1967, p. 221), and on the following pages Skerman describes a good technique for the observation of the oxidation of carbon monoxide in actively growing cultures of this group. Since some organic matter in some form or another is nearly always present in the natural environment of a freshwater lake, this may not be an ideal site for the initial isolation of this group of organisms. It is a matter for speculation that some of the other facultative autotrophs may be capable of oxidizing CO to C02 under certain conditions of laboratory culture. Reference should be made to Bergey (1957, p. 77) for a discussion on the one species of this group so far described.

I. Group i :methane-oxidizing bacteria-Methanomonas 1 . Site The source is Jenkin surface-mud cores obtained from the de-oxygenated zone of a stratified lake, and stored, covered, in the laboratory, with some of the overlying water removed to leave an “airgap” in the closed-core tube. After a period of 2-3 weeks, when the mud has warmed to laboratory temperature (i.e., 14”-18”C), the top 6-8 cm of the mud core breaks away, and rises to the top of the Fube, This is due to the evolution of gases proceeding in the lower depths of the column of mud. !

$

2. Method Samples, taken from the bottom of the 6-8 cm “plug” of risen mud, and inoculated into the autotrophic medium for Methanomonas, described by Skerman (1967, p. 221), and used under an atmosphere of 50% methane

24

V. G . COLLINS

and 50% air, yields enrichment cuItures of this group. For pure cultures the use of washed agar is essential. T h e inorganic salts of the aforementioned medium are incorporated and the cultures maintained in an atmosphere of one part methane and two parts air. Skerman (1967, p. 221) gives a very useful description of methods for observing the oxidation of methane, and also methods for the preparation of gas mixtures. For alternative cultivation methods see Overbeck (1969, Anagnostidis and Overbeck (1966).

L. Group 1: organisms capable of oxidizing compounds of iron and manganese-Sphaerotilus, Leptothrix, Crenothrix, Clonothrix, Gallionella, Siderocapsa, Naumanniella, Ochrobium, Siderobacter, Ferribacterium, Sideromonas, Peloploca, Pelonema, Ferrobacillus 1. Site Sources are water samples and mud cores obtained from a stratified lake during the period of summer stagnation. 2. Method Enrichment cultures for most of the organisms in this group can be obtained directly from water samples and mud cores, stored at laboratory temperatures. I n both cases the samples should be stored with access to atmospheric oxygen, which allows the reduced iron compounds, in samples from the de-oxygenated zone of the lake, to slowly oxidize. As re-oxygenation of the sample proceeds, large flocs of iron “complex” material are formed, and with the flocs copious growths of the iron bacteria develop in the stored samples of water and mud. For organisms belonging to the Sphaerotilus-Leptothrix group, the methods and media of Mulder and van Veen (1963) are ideal for the purpose of obtaining both enrichment and pure cultures. For species of Gallionellu, the methods and media of Nunley and Krieg (1967) are the only ones that the author has found successful for the pure culture isolation of Gallionella. See also Hanert (1968). T h e iron flocs that form on the surface of stored mud cores provide a good source of material for the isolation of Ferrobacillus species, using the medium and culture growth methods of Silverman and Lundgren (1959a). For pure culture studies concerned with the chemical activities of these organisms, reference should be made to the work of Dugan and Lundgren (1964), and Temple and Colmer (1951). T h e author has not been able, so far, to isolate any of the species of Siderocapsaceae, in pure culture. Enrichment cultures for these organisms can be obtained from the surface of submerged plants in the littoral regions of a stratified lake. It is possible to sediment large accumulations of the growth of these organisms from water samples taken at the period of over-

I. AUTOTROPHS

25

turnof astratified lake. This is the period when the temperature and dissolved oxygen concentration are the same from the surface of the lake to its greatest depth, the temperature range is ll0-12"C, and oxygen usually 90-100% saturation, A useful technique for obtaining material representative of the organisms in the Siderocapsaceae for taxonomic purposes is to submerge squares or strips of P.V.C. sheeting in the lake, at depth intervals during the stratified phase and also on the overturn of the lake. These organisms very quickly colonize the P.V.C. sheeting and, on removal to the laboratory, small sections of the sheeting can be cut out and mounted on microscope slides for direct observation. Since most of the species in this family form some kind of attachment extrusion, surrounded by a "torus" of iron compounds, the entire assemblage can be viewed microscopically on the pieces of P.V.C. sheeting. Descriptive studies on these organisms have been published by Cholodny (1926); Dorff (1934); Hardman and Henrici (1939); and Beger (1941). Pringsheim (1949) gives a most useful review of the iron bacteria with 199references. A most useful study on the bacteria associated with manganese nodules from the aquatic environment is reported by Trimble and Ehrlich (1968). Concerning the filamentous bacterium Sphaerotilus natans a great deal of published work now exists. Particularly because of its association with paper mill effluents, stream pollution and sewage treatment plants. Some of the industrial implications of the biological activities of this organism are described in the following publications; Waitz and Lackey (1958); Mulder (1964); Phaup and Gannon (1967); Muellar and Litsky (1968); and Phaup (1968). IV. DISCUSSION O N T H E AUTOTROPHIC WAY OF L I F E I N T H E NATURAL ENVIRONMENT A stratified freshwater lake has been chosen by the author as a natural environmental site for the isolation of the major groups of organisms thought to be autotrophic. From the evidence presented by detailed ecological and microbiological studies, covering twenty-one years of research on the freshwater environment, it would appear that there are very few strictly autotrophic bacteria existing and functioning in their natural environments. Many of the so-called autotrophic bacteria can be recovered from the natural situation of a stratified lake when it is known that organic matter is present in both the waters of the lake and the surface mud at the bottom of the lake. The gap existing between a laboratory culture of an apparently autotrophic bacterium and the natural site from whence the culture was derived is a very large one indeed. In many cases, the ability of thc cultured

26

V. G . COLLINS

organism to grow without organic matter in vitro may simply be the result of our inability to measure the micro amounts of organic matter derived from “contaminating” substances in the ingredients used for making the culture media in which the organism is growing. T h e author therefore prefers the use of the term “facultative autotroph” when referring to organisms known to be actively participating in the various chemical cycles of the natural environment, such as the nitrogen cycle, the sulphur cycle, and the iron cycle, T h e reader, the author hopes, will appreciate that there are many more natural sites where these organisms can be recovered, and reference to the “habitat” descriptions in Bergey (1957) give adequate proof of this fact. It should also be noted that the author has omitted certain genera of the various groups under consideration, having had no personal experience with the particular genera omitted.

V. MEDIA AND METHODS A. Organisms of groups a, b, d, i, j, and k For these organisms Pope and Skerman mineral salts media are used (Skerman, 1967). For the preparation of various mineral salts media the solutions described in Tables I and I1 are required. They should all be prepared with glassdistilled water and acid-cleaned glassware. TABLE I Solutions required for the preparation of various mineral salts media, Group A (from Skerman, 1967) Solution __---

__

Amount

.

-

__

2. 0.074 M 3. Solution from 2 , above

1 litre 1 litre 200 ml

4. Solution from 3, above

1 litre

1.

N

NaOH

5. NaHC03 in 100ml water 6. CaClz in 100ml water 7. NaNOz in 100 ml water 8. Glucose in 100 ml water 9. Mannitol in 100 ml water 10. Sucrose in 100 ml water 11. Sodium citrate in 100 ml water 12. Phenol in 100 ml water 13. Na&03 in 100 ml water

Procedure

_.

8.333 g 5 .O g 5 .O g 10.0 g 10.0 g 10 .O g

Sterilize at 121°C for 20 min Sterilize at 121°C for 20 min Dilute to 2000 ml(O.0074 M); sterilize as for Solution 1 Neutralize with use of N NaOH ; sterilize as for Solution 1 Sterilize as for Solution 1 Sterilize as for Solution 1 Sterilize as for Solution 1 Sterilize at 110°Cfor 25 min Sterilize at 110°Cfor 25 min Sterilize at 110°Cfor 25 min Sterilize at 121“C for 20 min

2.0 g an hydrous (or 2.77g hydrated) Sterilize at 121“Cfor 20 min 10 g Sterilize at 121°C for 20 min 10 g

27

I. AUTOTROPHS

TABLE I-continued Solution

Amount . .

-

100 ml 1 litre

14. 0.5 M He1 15. 0.0167 M H3P04

16. Monoethylamine hydrochloride in 100 ml water

5 ml

___

__

Procedure ~

___

Sterilize at 121"Cfor 20 min Neutralize 500 ml with N NaOH; sterilize as for Solution 11 Sterilize by filtration

TABLE I1 Solutions required for the preparationof various mineral salts media, Group B (from Skerman, 1967) Amount per Solution

100 ml solvent

Solvent required

3.0 g 6.6 g 21 .O mg 80.0 mg 106.0 mg 600.0 mg 123.0 mg 110.0 mg 109.0 mg 60.0 mg 30.0 mg 30.0 mg 629.0 mg 1.4g 36 .O mg 300.0 mg

0.0074 M H3P04 0.074 M H3P04 0.0074 M 0.0074 M 0.0074 M H 8 0 4 0'0074 M H3P04 0.0074 M 0 -0074 M 0.0074 M 0.074 M Water Water 0.074 M &PO4 Water Water Water

Final concentration (pgllitre medium) 300,000 660,000 21 80 106 600 123 110 109 60 30 30 629 140,000 36 300

Preparation of the basal mineral salts medium is carried out as folIowsStep 1. Pipette into a 1 litre standard flask the following amounts of solutions from Group B : 10.0 ml of Solutions 1 and 2 and 0.1 ml of each of Solutions3-10. Step 2. Add approximately 600ml of 0.0074~H3PO4(Solution 3, Group A) and 210 ml of water. Step 3. Adjust the p H to 7.0 with N NaOH (Solution 1, Group A). Step. 4. Add 0.1 ml of Solutions 11 and 12 from Group B. Step 5 . Take 0.1 ml of the MnClz solution (Solution 13, Group B), add 9.9 ml of 0,074 M &Po4 (Solution 2, Group A), and adjust the p H to 7.0. Autoclave and filter. Add the filtrate to the medium.

28

V. G . COLLINS

Step 6. Add 10 nil of Solution 14, Group B, and 0.1 ml of 15 and 16, Group B. Step 7. Using the neutralized 0.0074 M &PO4 (Solution 4, Group A), make the final volume to 1 litre. Step 8. Sterilize at 121°C for 20 min. This solution is crystal clear and will remain so for long periods if kept in acid-washed glassware. It provides a complex mineral salts base with ammonium-N, which has been found suitable, after addition of specific components, for a wide range of autotrophic and exacting heterotrophic bacteria. The use of Pope and Skerman mineral salts media for some of the groups of organisms covered in the preceding Sections is described below.

1. Group a :ammonia-oxidizing bacteria, Nitrosomonas species The medium for Nitrosomonas (Skerman, 1967) is used. (a) Liquid medium. Adjust the pH of the basal mineral salts medium to 8-2 before sterilizing. Add aseptically 20 ml of the sterile NaHC03 solution (Solution 5, Group A) per litre. (b) Silica Gel Plates. In the preparation of the silica-gel plates the medium is diluted 1 : 2. T o allow for this, prepare a double strength basal mineral salts medium (D.S.B.M.S.M.) as followsFollow the instructions for the single strength medium to Step 2. Add 105 ml of distilled water and make the volume up to approximately 400 ml with the use of the 0.0167 M H3P04 (Solution 15, Group A). Neutralize with N NaOH (Step 3). Then follow Steps 4, 5, and 6 as indicated. Make the final volume to 500 ml with the use of the neutralized 0.0167 M H3P04 (Solution 15, Group A) and sterilize at 121°C for 15 min. Prepare the silicic acid by the method of Pramer (see preparation of silica gel below). Before gels can be prepared it is necessary to determine the quantity of N NaOH (Solution 1, Group A) required to adjust the pH to the desired level after the addition of NaHC03 and CaClz solutions (Solutions 5 and 6, Group A). As a trial, mix 10 ml of silicic acid and 10 ml of D.S.B.M.S.M. Neutralize with N NaOH and note the amount added (x). Add 0.4 ml of the NaHC03 solution (Solution 5, Group A), and then adjust the pH to 8.2 with N NaOH (Solution 1, Group A). Divide the sample (approximately 20 ml) into four aliquots and add varying amounts of the CaClz solution to each. Allow 2 to 3 h to gel, and then determine the most suitable amount of CaClz solution (y) required for the whole 20.0 ml sample (approximately 0.6-0.8 ml is required),

29

I . AUTOTROPIIS

Before procecding to pour the plates, prepare another test plate as follows. MixSilicic acid D.S.R.M.S.M. N NaOH (Solution 1, Group A) CaClz solution (Solution 6, Group A) Inoculum NaHC03 solution (Solution 5 , Group A)

10 ml 10 ml x ml 0.6-0.8 ml 1 ml 0.4 ml

Immediately determine the p H and add N NaOH (Solution 1, Group A) until the pH rises to 8.2. Note this amount (XI). To prepare the plates substitute the value of (x + XI) for x in the previous mix. Allow 2 h to gel and then bcubate in a humid chamber at 28°C. (c) Preparation of silica gel. Pramer’s method is used as described by Skerman (1967). (d) Preparation of silicic acid sols. To prepare an ion-exchange column take approximately a 70 cm length of 25 mm glass tubing and fit it at one end with a glass tap and mount it vertically in a stand. Place some glass beads in the base and cover with a layer of glass wool. Pack 120 g (wet weight) of IR-120 Amberlite resin in the tube and add sufficient 2~ HCI to cover the resin. Remove any air bubbles from the column with a glass rod. Open the tap and pass 1000 ml of 2 M HCI through the column. Drain and then flush with distilled water until the effluent no longer gives a test for chloride with silver nitrate. Leave the column full of water. Immediately after using the column for the preparation of the silicic acid, wash it again with water and regenerate with 2 M HCl. Prepare 500 ml of a solution of sodium silicate containing 1.5% of SiOz and allow it to flow through the column at 5 ml/min. Check the p H of the effluent and collect for use when the p H falls below 3.4. Adjust the pH of the solution to 2.0 with HCI. T h e solution should be stable at this pIi. Sterilize at 110°C for 25 min.

2. Group b: nitrite-oxidizing bacteria, Nitrobacter species The medium for Nitrobacter (Skerman, 1967) is used. (a) Liquid medium. Prepare the basal mineral salts medium with omission of the ammonium sulphate (Solution 2, Group B) and sterilize. Add aseptically 8 ml of the NaNOz solution (Solution 7, Group A) and 20ml of the sterile NaHC03 solution (Solution 5 , Group A) per litre of medium. Adjust the p H aseptically to 8.8 with sterile N NaOH (Solution 1, Group A).

30

V. G . COLLINS

(b) Silica gels. Proceed as for Nitrosotnonas with omission of the ammonium sulphate (Solution 2, Group B) in the preparation of the double strength basal mineral salts medium (D.S.B.M.S.M.). The mixSilicic acid D.S.B.M.S.M. NaNOz solution (7, Group A)

10 ml 10 ml 0.16 ml

Neutralize with N NaOH (Solution 1, Group A), noting the amount used (x). Add 0.4 ml of the NaHC03 solution (Solution 5 , Group A), and then adjust the p H to 8.8. Divide the sample into four aliquots; add varying amounts of CaCl2 solution (Solution 7, Group A) to each and allow to stand for 2-3 h to gel. Determine the amount of CaCl2 solution (y) that will give the optimal gel in the total (approximately 20 ml) sample. Prepare another mix as followsSilicic acid D.S.B.M.S.M. NaNOz solution (Solution 7, Group A) Inoculum N NaOH (Solution 1, Group A) CaClz solution (Solution 6, Group A) NaHC03 solution (Solution 5, Group A)

10 ml 10 ml 0.16 ml 1 ml x ml Y ml 0 - 4 ml

Determine the p H immediately; adjust to p H 8.8 with N NaOH (Solution 1, Group A) and note the amount (XI). To prepare plates substitute the value of (x+xl) for x in the previous mix. Allow 2 h to set and then incubate at 28°C in a humid chamber.

3. Group a and Group b (a) Autotrophic media for Nitrosomonas and Nitrobacter (J. Meiklejohn personal communication to Skerman 1967). NaCl MgS04. 7Hz0 FeS04.7HzO H2O 0.1 M KH2P04

+ (NH4)zS04 (for Nitrosomonas)

+ NaNOz (for Nitrobacter)

0.3 g 0.14 g 0.03 g 90 ml 10 ml (Previously boiled for 30 min, cooled, and made up to volume). 0.66 g 0.5 g

Dilute to 1000 ml and add 10 g of powdered CaC03 and 0.4 ml of a trace element solution supplying Mn, 22 pg; €3, 21 pg; Cu, 17 pg; Zn, 16 pg; and CO, 14pg. Dispense in layers not more than 1 cm deep in Erlenmeyer flasks. Sterilize at 121°C for 15 min.

I. AUTOTROPHS

31

(b) Quantitative determination of nitrite. T h e method of Lees and Quastel described by Skerman (1967) is used. Dilute a sample containing approximately 0.5-5.0 pg of nitrite-N to 11 ml with distilled water. Add 2 ml of Griess-Llosvay reagent. After 30 min read the colour density in a photoelectric colorimeter with the use of a suitable filter and determine the nitrite-N concentration from a curve prepared from a series of standard nitrite solutions. The Griess-Llosvay reagents are prepared as follows(i) Sulphanilic acid solution: Dissolve 0.5 g in 30 ml of glacial acetic acid. Add 100 ml of distilled water and filter. T h e reagent is stable for onemonth. (ii) a-Naphthylamine solution: This should not be more than one week old. Dissolve 0.1 g of a-naphthylamine in 100 ml of boiling distilled water. Cool and add 30 ml of glacial acetic acid. Filter. For use mix Solutions 1 and 2 in equal quantities. The standard nitrite solution is prepared as followsDissolve 0-493 g of pure sodium nitrite in water. Make up to 1 litre. Dilute 10 ml of this solution to 1 litre. This final solution contains 0.001 mg of N as nitrite. T h e standard curve should be prepared over a range of 0.0005-0.001 mg of nitrite nitrogen.

4. Group a : ammonia-oxidizing autotrophic bacteria, additional medium The medium of Skinner and Walker (1961), as described by Soriano and Walker (1968), is used. The medium contains: (NH4)$304, 0.5 g; KHzP04, 0.2 g; CaC12.2HzO 0.04 g; MgS04.71-120, 0.04 g; Fe (as ferric citrate or Fe-EDTA chelate), 0.5 mg; phenol red, 0.5 mg; distilled water, 1. litre. Plates are prepared by adding 1-1.5% of special Noble (Difco) or purified (Merck) agar to the above solution. After sterilizing at 120°C for 15 min, the p H of the medium is adjusted to 7.5-8.0 by adding a sterile aqueous 5% sodium carbonate solution. The author has used the isolation techniques described by Soriano and Walker (1968). These are excellent and, therefore, are included here for reference. Dilution. Serial dilutions in sterile medium were made from an enrichment medium culture so that the final dilution contained an average of 1or 2 organisms/ml. Tubes (usually forty to fifty) containing 8 ml of medium were inoculated with 1 ml of the final dilution and incubated at 25" or 30°C for some weeks. At intervals, tests were made for the presence of acid and nitrite, and positive tubes were subcultured on peptone agar to check for heterotrophicorganisms,

32

V. G. COLLINS

Plating. An inoculum consisting of a drop of undiluted or diluted enrichment culture was spread over the surface of a dried agar plate by means of a sterile bent platinum wire (the microspatula of Beijerinck). T h e plates were incubated at 25” or 30°C in a closed jar containing dilute aqueous ammonia until nitrite was detected in the agar and presumptive colonies of nitrifiers could be seen under the microscope by transmitted light at x 80 or x 100 magnification. Contaminant colonies were mostly visible without magnification. Isolated microcolonies were sucked into capillary pipettes and transferred to tubes of sterile liquid medium. B. Organisms of groups c, e, f, and g Winogradsky cylinder technique for Group (c) non-photosynthetic sulphur bacteria, Groups (e) and (f) photosynthetic sulphur bacteria and Group (g) photosynthetic non-sulphur bacteriaT h e mud is mixed with CaS04 and some insoluble organic matter, e.g. cellulose, poured into a tall glass cylinder, whereupon the cylinder is filled completely with water containing O.lyoNH4C1 and 0.1% phosphate buffer at pH 7.3 and incubated with continuous illumination. T h e enriched mud serves as a continuous H2S generator (bacterial sulphate reduction), and the photosynthetic sulphur bacteria soon appear as coloured spots at the mudglass interphase. T h e superiority of the Winogradsky method is not only due to the fact that it permits the use of large amounts of mud as inoculum; additional advantages are the continued formation of H2S and the localization in space of the photosynthetic bacteria, thus facilitating their detection in the form of “colonies”.

1. Group c (a) Enrichment culture techniquefor Beggiatoa species. This is based on a modification of Cataldi’s (1940) technique, and described by Faust and Wolfe (1961). Dried roadside grass, which has been exposed to the weather for the autumn months is cut into small pieces and added to boiling tap water in a ratio of about 100 g of grass to each litre of water. After boiling for about 10 min, the water, which then contains soluble components from the grass, is decanted. Additional water is added and boiling is repeated. After five cycles of boiling and decanting, the extracted hay is allowed to stand overnight in water, and on the following day the extraction procedure is repeated three additional times before the hay is spread out and dried. About 0.5 g of extracted hay and 60 ml of stream water are added to each 125 ml Erlenmeykr flask. Each flask is inoculated with a small piece of decaying leaf or other debris from a polluted stream and is plugged with

33

I. AUTOTROPHS

cotton to retard evaporation. Enrichment flasks are incubated at room temperature (about 28°C). (b) Methods for obtainingpure cultures of Beggiatoa species (Faust and Wolfe, 1961). By means of a capillary pipette, a tuft of trichomes from a crude enrichment flask is transferred through several drops of steril tap water to remove most of the contaminating bacteria. T h e tuft is then placed on the dry surface of a solid medium (medium A of Faust and Wolfe) which consists of yeast extract, 0.2 g; agar, 1.5 g ; tap water, 10 ml; glass distilled water, 90 ml; and is adjusted to p H 7.0. Standard sterilization techniques are used in all media preparation. A dry surface is obtained by allowing excess water to evaporate from the surface of the medium in a 30°C incubator for about 2 h with the lid of each dish propped open slightly. Between 4 and 6 h after inoculation each plate is examined under a dissecting microscope at 30 x magnification. A small agar block containing a well isolated trichome is cut and transferred aseptically to 2 ml of sterile liquid medium A. Pure cultures are then obtained, after repeated plating and washing procedures where necessary, and the pure trichomes of the organism are finally transferred to a semi-solid medium (medium D of Faust and Wolfe). This is adjusted to p H 7.0 and contains yeast extract, 0.2 g; agar, 0.2 g ; CaC12,O.Ol g; sodium acetate, 0.1 g; and distilled water, 100 ml. T h e medium is filled out into test tubes (15 by 150 mm) with screw caps, and growth occurs in a narrow disc about 0.5 cm beiow the surface of the medium. This characteristic is typical of gradient organisms which require a specific oxidation-reduction potential for growth. Incubation should be at 17"C, on this medium; for maintenance of stock cultures repeat transfers must be made every seven to ten days.

2. Group d: isolation media (a) Themedium of Lieske (1912). This medium, described by Collins (1963), is particularly useful for the isolation of Thiobacillus dentrificans, and consists of the following ingredientsNazSz03.5H20 KNOB NaHC03 K2HP04 MgCh

0.2g 0.1 g

CaCIz

traces

FeC13 distilled water

traces 1 litre

5.og

5.Og 1.og

(b) The medium of Waksman and Starkey (1922). This medium, described by Collins (1963), is useful for the isolation of Thiobacillus thio-oxidans, and

34

V. G . COLLINS

is composed of the following ingredients per litre of distilled water0.2 g 0.1-0.5 g 0.01 g

0.25 g 3-5 g 10 g

(c) The medium of Starkey (1935). This medium, described by Collins (1963), is useful for the isolation of Thiobacillus thioparus, and is composed of the following ingredients per litre of distilled water5.0 g 0.4 g 4.0 g 0.25 g 0.5 g 0.01 g

(d) Neutral medium for nonaciduric species of Thiobacillus (Skerman, 1967). Use the Pope and Skerman Basic Mineral Salts Medium. Follow the preparation of the basal medium, except at Step 2 add 110 ml of water instead of 210, and at Step 7 make the final volume to 900ml instead of 1000. Sterilize at 121°C for 20 min. Cool. Then add aseptically 100 ml of sterile NazSz03 solution (Solution 5, Group A). The medium should be dispensed in a depth of not more than 1 cm in Erlenmeyer flasks.

3. Groups e and f : isolation media (a) Detailed description for the preparation of culture medium for red andgreen sulphur bacteria. This is described by Pfennig (1961 ; 1962), and in personal communication to the author. Solution 1 Distilled water CaClz anhydrous

2500 ml 2s

500 ml of this solution are autoclaved separately in an Erlenmeyer flask. 2000 ml are distributed in amounts of 75-80 ml into 127 ml (4 0 2 ) screwcapped bottles and autoclaved. Before adding sterile filtered sohtions 2 and 3 (below) the bottles are kept cold (4"-1OoC).

35

I. AUTOTROPHS

Solution 2

Distilled water Heavy metal solution Vitamin-Biz solution KH2P04 KCI NH4CI MgCI2.6H2O Na ascorbate

32 nil 50 ml 15 ml 1g 1g 0.8 g 0.8 g

2.4 g

(To be added only as a separate sterile solution to the ready prepared medium. Ascorbate can be omitted, if larger inocula are used.) Solution 3 900 ml 4.5 g

Distilled water NaHC03

Gaseous CO? is bubbled through for at least 30 min; p H about 6.1. Solution 4

Distilled water NazS .9Hz0

200 ml 3g

Prepared using a magnetic stirrer rod. This solution is autoclaved. After CO:! saturation of solution 3, solution 2 is added and the mixture immediately filtered through a Seitz filter using C 0 2 pressure to push the liquid through (no suction). T h e sterile filtered solution 2+ 3 is added to the 127 ml bottles containing cold 75-80 ml CaClz-solution 1. T h e sterilized cold solution 4 is partially neutralized by adding (on a magnetic stirrer) drop by drop 2 ml sterile 2 M HzS04. This partially neutralized solution is added to the bottles in 6 ml amounts for Chromatiurn and Chlorobium and 3 ml per bottle for Thiospirillum jenense. T h e bottles are nearly filled up with liquid by adding sterile solution 1 (from the 500 ml flask). A small air bubble is left. T h e bottles are immediately closed. ?'he final pH has to be 6.6-6.8, and this will be obtained, if these directions arc followed exactly. The freshly prepared medium becomes slightly turbid due to the oxidation of some HzS to sulphur by dissolved oxygen. After storage of the medium for 1-2 days the turbidity disappears and a slight sediment is formed which turns black after some days. If the tightly closed bottles are stored in the dark the culture medium keeps for several months. The freshly prepared cdture medium should be aged in the bottlcs for at least 24 h before inoculation., The cultures are incubated at room temperature (20°-25"C) at a distance of 30-40 cm from a 40 Wtbulb. Light periods of 16 h and dark periods of 8 hare favourable.

36

V.

G. COLLINS

If the growing organisms have used up both the HzS and the stored sulphur (disappearance’ of the chalky appearance) fresh neutralized NaZS solution has to be added. A suitable volume of solution 4 is added to a sterile Erlenmeyer flask with a magnetic stirrer rod and neutralized on a magnetic stirrer by adding drop by drop sterile 2 M HzS04 only until a slight sulphurturbidity appears, This turbidity will disappear if an excess of HzS04 is avoided. T h e solution will be slightly yellow but clear. 5-6 ml of the neutralized solution arc added to each 127 ml bottle of the Chromatium or Chlorobium strains; 3 ml are added to the Thiospirillum jenense cultures. After the addition of the sulphide solution the cultures are kept in the dark for some hours; thereafter they are put back into the light. Heavy metal solution Distilled water Ethylene diaminetetra acetate

Modified “Hoagland trace element solution” FeS04.7Hz0 ZnS04.7HzO MnClz .4Hz0

1000 ml 1* 5 g (EDTA has to be dissolved first) 6 mi 200 mg 100 mg 20 mg

Vitamin Bl2 solution Vitamin BIZ(Cyanocobalamin, Merck) 2 mg/100 ml distilled water Vitamin solution 100 ml Distilled water 0.2 mg Biotin Nicotinic acid 2 mg 1mg Thiamine p-Aminobenzoic acid 1 mg Pantothenic acid 0.5 mg 5 mg Pyridoxamin-HC1 Modified “Hoagland trace element solution” AIC13 IF! KI 0.5 g KBr 0.5g LiCl 0.5 g MnCk .4Hp0 7g H3B03 11 g ZnCla f g CUCI? Ig NiCls 1g COClZ Ig SnClz. 2H20 0.5 g BaCl2 0.5 g NazMo04 0.5 g NaV03. H20 0.1 g Se-salt 0.5 g

I. AUTOTROPHS

37

Each salt is dissolved separately in distilled water. Before mixing together, the pH of each solution is adjusted below pH 7.0. The total final volume is 3.6 litres (1/5 of the total final volume of Hoagland, 18 litres). The pH of the final solution is adjusted to pH 3-4. The flaky yellow precipitate which is formed after mixing transforms after a few days into a very fine white precipitate. Before use the solution is mixed thoroughly. (b) Group e: the medium of van Niel (1931). This medium is described by Collins (1963). Inoculate water or mud samples into medium of composition 1.0g NHdCI, 0.5 g K2HP04, 0.2 g MgC12, 1.0 g NaHC03, 1.0 g Na2S.9HzO per litre of distilled water (pH 8-8*5), contained in 4 oz glass-stoppered bottles. The NaHC03 is prepared as a 5’7’ solution, and sterilized by filtration through a Seitz filter; the Na2S.9H20 is prepared as a 10% solution and autoclaved separately, the remainder of the ingredients being autoclaved in bulk. After sterilizing, 10 ml/litre of the NazS. 9Hz0 are added to the bulk, along with 20 ml/litre of the NaHC03 solution to complete the medium. (c) Group f: the medium of Larsen (1952). This medium is described by Collins (1963). The medium consists of tap water with 0.1% each of NH4C1, KH2P04, and Na2S.9H20; 0.05% MgCl2; 0.2% NaHC03, and NaCl as required; the initial pH is adjusted to 7.3 (van Niel, 1931). See also the description of the Winogradsky (1887) cylinder enrichment method, as described by Laisen (1952). (d) Group g (photosynthetic non-sulphur bacteria): the medium of van Niel (1944). This medium is described by Collins (1963). The composition of the medium is 1.0 g (NH&S04 or NHdCI, 0.5 g KzHPO4, 0.2 g MgS04.7H20 or MgCl2, 2.0 g NaCI, 5.0 g NaHC03, 0.15-2% proteose peptone, or an alternative organic substance per litre of distilled water (pH 7.1-7-2 adjusted by means of sterile solutions of lisp04 and Na3C03 as required). Note, when chloride is substituted for MgS04 use 0.25 ml of a saturated solution of MgClz per Iitre, an approximate equivalent. (e) Group g: Hutner’s (1946) method and medium for agar slant cultures. Many different media proved suitable for the maintenance of cultures. The substratewas adequately supplied as lactate, 0*2-0-4%, or malic acid (natural or synthetic), 0.1-0.4~0;occasionally sodium acetate (hydrated), 0.05-0.1%, or sodium butyrate, 0*0470,was added to malate and lactate media. Media not containing malate contained sodium citrate (hydrated), 0.025-0.1 %, to ensure full availability of heavy metals and calcium. The unidentified requirements were adequately supplied as trypticase (Baltimore Biological

38

V.

G. COLLINS

Laboratories), 0.1--0*2y0,or thiopeptone (Wilson), 0.10/,. Yeast extract (Difco) was inhibitory to inany strains, while trypticase was non-inhibitory and permitted extremely good growth. T h e remainder of the medium consisted of agar, 1.5yo; small amounts of K2HP04, MgS04.7H20, (NH&HP04, and Fe, 0.1-0.4 mg%; and Mn, 0.05-0.2 mg%. T h e p H was adjusted to 6.5-6.8. These slants were made in screw-capped tubes (bottles would do equally well) with the rubber liners of the metal caps removed. Media were autoclaved for 10 min at 118°-121”C. Hutner states that a few isolates designated “Rhodovibrio”, which appeared to be microaerophilic, and a few isolates of Rhodospirillum rubrum, which although uninhibited by air, seemed unusually sensitive to inhibitory substances or were exacting for other reasons, were grown on the same media rendered semi-solid by decreasing the agar to 0.2-0*4%. T h e growth of slant cultures was usually heavy in 24-48 h. They were then stored in the dark at 6°C. They remained satisfactorily viable for at least a month. There was no obvious impairment of photosynthetic ability as a result of this treatment. (f) Group g: Hutner’s (1950) medium for growth-factor requirements under anaerobic conditions. KaHP04 MgS04.7HzO DL-Mak acid Sodium succinate (hydrated) L-Glutamic acid Glycerol Potassium acetate Sodium indigodisulphonate or benzylviologen(B.D.11.) Zn Ca

Mn Fe

cu Mo

co

0.05 g 0.025 g 0.3 g 0.4 g

0.2 g 0.2 g 0.1 g 1.0 g 0 . 5 rng 1 .O m g 0.4 g 0 . 2 rng 0.1 mg 0.1 rng

0.05 mg 0.05 mg

Distilled water was added to 100 ml, pH adjusted to 6.8-7.1 with KOH. Sodium formaIdehyde sulphoxylate 0.05 g/100 ml was added separately as a freshly autoclaved 2% solution (w/v) reducing agent. Growth factors supplied when necessary as follows: aneurin, 0.1 mg; nicotinic acid, 0.1 mg; p-aminobenzoic acid, 0.01 mg; and biotin, 0.4 pg. T h e concentrations of metals listed refer to the metal content of the salt used. These were usually sulphates.

39

I . AUTOTROPIIS

Hutner also lists a very useful “preservative” mixture for stock solutions as follows: a mixture (v/v) of part o-fluorotoluene, 2 parts n-butyl chloride, and 1 part 1,2-dichloroethane (Hutner and Bjerknes, 1948). This preservative volatilizes on autoclaving. C. Organisms of Group h-Desulfovibrio

1. Gvoup h: isolation media (takenfrom Skerman, 1967). The following modifications of media described by Baars (1930) and Starkey (1938) are recommended by Butlin, Adams, and Thomas (1949). Baars’ Medium K2HP04 NH4Cl CaS04 MgSO4.7HzO Sodium lactate, 70% solution Tap water

0.5 g 1.og 1.og 2-0 g

5.0 g 1000 ml

Dissolve the salts; adjust the p H to within the range of 7-0-7.5 and sterilize at 121°C for 20 min. Prepare separately a 1% solution of FeSO4. (NH4)2S04.6€1~0and sterilize by steaming for 1 h on three successive days. Add 5 ml of the supernatant per 100 ml of the above medium immediately before use. Note: This medium has a heavy precipitate but is quite suited for crude cultures. Starkey’s Medium K2HP04 NH4Cl Na&04 CaC19.2H20 h~gS04.71320 Sodium lactate, 70:/0 solution Distilled water

0.5 g 1.og 1.og 0.1 g 2.0 g 5.0 g

1000 ml

Dissolve the ingredients and adjust the p H to between 7.0 and 7.5. Sterilize at 121°C for 20 min. Note: This medium has a slight precipitate, which may be removed by filtration after sterilization, following which the medium may be resterilized. Prepare a 1% solution of FeS04. (NH&S04. 6 H Z 0as for Baars’ medium above, and add 5 m1/100 ml of medium just before use. For halophilic strains, add 1-3% NaCl to each of the above media before sterilization,or, alternatively, replace the tap or distilled water with seawater.

2. Group h: medium for the isolation and maintenance of pure cultures (from Skerman, 1967) The media of Baars and Starkey, described above, are not ideal for pure culture studies. Skerman recommends the following medium, described by

40

V. G . COLLINS

Butlin and his associates and modified by Postgate. T h e medium is similar in most respects to that published independently by Miller (1950). K2HP04 NH4Cl Na2S04 CaClz .6H2O MgS04.7Hz0 Na lactate, 70% solution (sterilize separately) Difco yeast extract FeS04.7HzO Distilled water

0.5 g 1.og 1.og

0.1 g 2-0g 3.5 g 1-Gg 0.002 g 1000 ml

Dissolve the ingredients, adjust the p1-I to 7.5, and autoclave at 121°C for 20 min. Filter off the sediment. dispense as required, and resterilize. Prepare separately a 0.6% solution of cysteine hydrochloride in distilled water, and sterilize by autoclaving at 121°C for 20 min. This acid solution has a p H of 1.8 and is relatively stable to oxidation, provided it is not neutralized. Add 1 ml to each 9 ml of medium immediately before use. T h e final concentration of cysteine is 5 pmoles/ml. Pick black colonies showing the correct morphological types on microscopic examination into the liquid medium. Incubate aerobically. If it is desired, the cultures may be plated on the same medium containing 2% agar and incubated anaerobically in an atmosphere of hydrogen and 5% C 0 2 in a McIntosh and Fildes Jar, with a dried pad of absorbent cotton wool, impregnated with lead acetate, between the cultures and the catalyst. See text for references to the authors mentioned in connection with this medium.

D. Organisms of Groupj-Hydrogenomonas A synopsis of a method for growing organisms of this group as described by Cohen and Burris (1955). The medium for growth of 11.facilis contains the following macro-nutrients per litre: NaHC03, 1-0g ; NHdCl, 1 4 g; KH2P04, 0.5 g; MgS04.7I180, 0.1 g; NaCl, 0-1 g ; CaC12, 0.1 g ; and Fe(NH&(S04)2. 6H20, 8 mg. T o this is added a mixture of micro-nutrients containing: H3B04, 228 pg; CoCl2.6H20, 80 pg; CuSO4. 5H20, 8 p g ; MnC12.4Ha0, 8 pg; ZnS04.7Hz0, 176 pg; and NazMoO4.2H20, 50 pg. The gas mixture made from commercial cylinder gases consists of 6 parts (by volume) of hydrogen, 2 parts oxygen, and 1 part carbon dioxide. The temperature for growth is maintained at 30°C by means of a thermostated bath. T h e pH of the medium as determined with a glass electrode after equilibration with the gas phase is initially 6.8-7-0. A small amount of Dow Corning “antifoam A” or silicon stopcock grease is added as an antifoam agent. The 8 litre of sterile medium are inoculated with 250-300 ml of a 36-48 h old culture grown in liquid medium in shaken flasks; each 500 ml

I. AUTOTROPHS

41

shaken flask contains approximately 125 ml of medium and the gas mixture described. In growing the hydrogen bacteria a gas mixture is circulated with a pump through the gas reservoir and thcn into the culture vessel; concomitantly the pressure of the system is maintained at essentially atmospheric pressure by displacing water into the gas reservoir as the gas mixture is used. T h e culture vessel is a narrow 10 litre Pyrex bottle (Corning No. GBYDI) fitted with a rubber stopper through which the gas inlet, gas outlet, and medium sampling outlet are attached. Both the gas inlet and outlet are connected externally to filter plugs (40 by 200 mm tubes packed with plugging cotton). A porous gas dispersion ball (“Marc0 Ball O’Mist Releaser”, obtained from the J. B. Maris Co., Bloomfield, N. J.) is connected to the end of the gas inlet, which extends nearly to the bottom of the vessel. In practice, 8 litre of medium are placed within the vessel and autoclaved. After cooling it is inoculated, and the sterile rubber-stopper assembly is put into position. T o avoid the hazard of evacuating this large vessel, C02 is blown through it for 5 min to displace air; the leads into and out of the vessel then are clamped off, and the vessel is set into the thermostated bath and connected to the gas reservoir and pump. After evacuating and filling the reservoir system with the gas mixture, the leads are freed, and the pump is turned on. Then the displacing water reservoir is connected. Gas reservoir bottles are 9 litre Pyrex serum bottles with concave bottoms. These bottles are suitable for evacuation. Connections in the system are made with rubber stoppers, butyl rubber tubing, and glass tubing; and these are made gas tight by sealing with either a mixture of 1 part beeswax to 1 part rosin or with ordinary finger-nail polish. T h e system is checked for leaks by partial evacuation. A gas inlet and evacuation port are connected through a three-way T stopcock to the apparatus. A manometer for the system serves in making up gas mixtures as well as in checking the system for leaks. The authors state that they employ four serum bottles to give a gas reservoir volume of 36 litre because it is convenient to fill the system with gas in the late afternoon and to harvest the next morning without having to refill the system. Any type of bottle which holds slightly less than the total gas volume of the system may be used as a water reservoir. Sulphuric acid sufficient to lower the pH of thc displacing fluid to p€I 4 is added to prevent binding of COa as bicarbonate in the displacing fluid. A “Ccnco Pressovac-4 Pump” is used to circulate the gas mixture; cotton gauze filters and a simple trap formed by using a 500 ml filter flask serve to filter the oil spray from the,gas train. A bypass around the culture vcssel permits control of the volume of gas passing through the culture vessel. In summary, the authors state that rapid growth of H . facilis was achieved by growing the organism in liquid culture, with vigorous aeration, main-

42

V. G . COLLINS

tenance of a reasonably constant gas pressure, and addition of micronutrient elements to thc medium were important. The medium of Schatz and Bovcll(1952), as described by Skerman (1967), 1s-

KHzP04 NH4N03 MgS04.7Ha0 FeS04.7H.O CaClz .2Hz0 Distilled water

0.1 g 0.1 g 0.02 g 0.001 g 0.001 g

to 100 ml

Adjust the pH to between 6.8 and 7.2. Where desired, incorporate 1.5% washed agar. For autotrophic growth, supplement the base with 0.05% NaHC03. Autoclave stock solutions of the NaHC03 separately, flush with COz, and add to the sterile medium before inoculation. Incubate under an atmosphere of 10% C 0 ~ , 3 air, 0 ~and ~ 60% hydrogen. 1. Group j : autotrophic medium using the Pope and Skerman mineral salts solutions (Skerman, 1967) Prepare the same medium as that employed for Nitrosomonas but adjust the p H to 7.0. Incubate under an atmosphere of 10% COz, 30% air, and 60% hydrogen,

E. Organisms of Group k-Carboxydornonas 1. Group k : autotrophic medium using the Pope and Skerman mineral salts solutions (Skerman, 1967) Use the same medium as for Hydrogenomonas but incubate under an atmosphere of 20% oxygen and SOYo carbon monoxide. 2. Group K : the medium of Kistner (1953) (Skerman, 1967) Kh'03 KzHP04 MgS04.7HzO Peptone HzO

2.0 g 1.og 0.1 g 0.2 g 1OOOmI

Dissolve the ingredients and adjust the p H to 7.2. Sterilize at 121°C for 20 min. For an agar medium use only sufficient agar to make a moderately firm gel. An agar that is too hard inhibits growth. Incubate under an atmosphere of 80% CO and 20% 0 2 . 3. Group k : Parkes and Mellor's method for producing carbon monoxide fm thegrowth of organisms (Skerman, 1967) Close a round-bottomed 500 ml flask with a rubber stopper fitted with a gas outlet tube just penetrating the stopper and a dropping funnel with the

I. AUTOTROPHS

43

lower end reaching almost to the bottom of the flask. Add concentrated sulphuric acid to the flask so that the tip of the dropping funnel is immersed. Place concentrated formic acid in approximately half this volume in the dropping funnel. Place the flask over a steam bath and connect the gas outlet tube via a concentrated NaOH wash bottle of soda-lime tube to the gascollecting apparatus. Heat to 100°Cand then admit the concentrated formic acid drop by drop until the required amount of gas has been collected. Allow for the exclusion of air before collecting the gas.

HCOOH~I-I~SO~-~I~~SO~E-~~O+CO 4. Groups i, j and k : Skerman’s (1967) recommended method and description ofan apparatusfor the collection and mixing of gases In culturing autotrophic gas-utilizing organisms, it is necessary to prepare gas mixtures of various types. This can be done simply with the apparatus shown in Fig. 11. This consists of a series of 500 ml gas-collecting

H

Frc. 1 1 . Apparatus for the collection and mixing of gases.

burettes, A , h’,C, and I>, conncctcd at thc base via a manifold to a 3 litre rescrvoir, E,containing dilute sulphuric acid; and via taps at thc top to a second manifold closed at both cnds by taps F and G. ‘The tube from tap G

44

V. G . COLLINS

is connected to the gas generator and both G and F are opened. Gas is allowed to stream through G and F to expel air or any previous gas. Then with F still open, the tap to A is opened and then F is closed. T h e gas is diverted into A and dilute acid is expelled to E. If desired, E may be lowered to reduce the back pressure on the gas generator. With A nearly filled, open F and then close A immediately. Disconnect the gas generator and then close G and F. Repeat with other gas to B and C. Any number of gas burettes may be employed, but allowance must be made for mixing. To prepare a mixture of 200 ml of A, 100 ml of B, and 200 ml of C and D, open F and then A to flush out the manifold. Close F. Lower E so that its meniscus is level with the 200 ml mark in D. Open D and let the gas flow in, adjusting E so that its meniscus is finally level with the water in D at the 200 ml mark. Close D and A. Raise E and open B and F to flush the manifold. Close F. Lower E to the 300 ml level of D and open D. Allow gas from B to flow into D to the required level. Close D and B. Raise E and open C and and F to flush the manifold. Close F. Lower E to the 500 ml mark in D and open D.Allow gas from C to flow in to the required level. Close D and C and replace F. T o discharge D into the required containers, open G and D and flush the manifold. Close G. Connect F to the apparatus and open F.Allow some of the gas mixture to flow through the connection to the apparatus to expel any air before admitting the gas.

F. Organisms of Group i-Methanomonas 1. Group i: autotrophic medium using the Pope and Skerman mineral salts solutions (Skerman, 1967) Use the same medium as for Hydrogenomonas but incubate under an atmosphere of 50% methane and 50% air. For the production of methane use Weygand’s method as described by Skerman (1967), or purchase the highest grade of commercial methane. Pure methane may be prepared by reducing methyl iodide in alcohol with a copper-zinc couple. Add 100 g of zinc dust to 250 ml of a 4% aqueous solution of copper sulphate. Shake thoroughly and then allow the powder to settle. Wash several times with water by decantation and then dry. Place the powder in a 100 ml Erlenmeyer flask fitted with a stoppcrcd dropping funnel and a gas outlet tubc. Allow a mixture of cqual volumes of methyl iodide and absolute ethyl alcohol to drop slowly onto the zinccopper couple. Collect the methane over water aftcr allowing the air from the flask to escape.

45

I. AUTOTROPHS

2. Group :Foster's autotrophic medium (Skerman, 1967) NaN03 MgSO4.7Hz0 FeS04.7HzO NasHP04 NaHzP04 CuSo4.5Hz0 fIsnO3 MnS04. HzO ZnS04.7HsO MOO3 KCl CaClz HzO

2.0 g 0.2 g 0.001 g 0.21 g 0.09 g 200.0 pg 60.0 pg 30-0 pg 300.0 p g 15.0 pg 0.04 g 0.015 g 1000 ml

Dissolve the salts and sterilize. Incubate under an atmosphere of 50% methane and 50% air.

3. Groups i, j and k : Skerman's (1967), recommended method for obsercing the oxidation of carbon monoxide, hydrogen, or methane by growing cultures The apparatus is illustrated in Fig. 12. A consists of a 10 ml graduated

pipette sealed at the tip and joined above the 0.0 ml mark to a 12 ml tube fitted with a 12 mm side arm and pear-shaped bulb with a 16 mm outlet. The tube is plugged at the outlet with cotton wool and sterilized. B is a glass tube of 6 mm outside diameter; bent so that the external arm liesparallel to the main axis of the tube when the tube is inserted as illustrated. The external end is unconstricted and is plugged with cotton wool. T h e internal end is only slightly constricted and extends into the tube until it almost touches the wall when the rubber stopper D fits snugly into the base of the side arm. A small 1 mm aperture, E, is made in the tube exterior to the rubber stopper. A cotton wool plug is rolled around the stem of B between the rubber

46

V. G . COLLINS

stopper and the bend. The lower end (bearing the stopper) is inserted into a 150 by 16 mm tube; and the assembly is sterilized. A third tube, C, bent in the same manner as B but without the small hole E and with the external end extended to a length that brings its tip level with the top of the graduated tube, completes the apparatus. This tube is fitted with a rubber stopper similar to D. Method for Use: Pipette 40 ml of the sterile synthetic medium into the tube A and inoculate the medium. Replace the plug in A with tube B, leaving the stopper D only loosely seated in its base. Connect B to the gas-mixing burette and, holding the tube on its side, loosen the stopper D and allow a quantity of gas to escape into the pearshaped bulb to exclude the air in tube B. Then turn the tube upright and collect approximately 9 ml of gas. If too much is collected it can be released by inclining the tube. Disconnect the gas supply and seat D firmly into the side arm. Supporting the tubes by the outlet tube, immerse them for 20 min in a water bath at a temperature as near that of the room as possible. Holding the tubes by the outlet tubes, slightly tilt the tube until the menisci in the closed arm and pear-shaped bulb are level; read the gas volume (Vl) and note at the same time the temperature of the water bath. All subsequent gas volume readings must be taken at the same temperature. Incubate the tubes on their sides attached to a rocking arm in a water bath. This provides a maximal gas-liquid contact area during incubation. The small aperture E allows movement of liquid between the two bulbs, but the position of the internal end of B does not allow escape of gas. After incubation adjust to room temperature to eliminate errors due to gas expansion. Replace tube B with tube C. Seat the stopper well into the side arm and introduce water through C, while holding the tube in the vertical position by the outlet tube, until the levels in the closed arm and in C are equal. Read the gas volume (V2). Release the stopper in the side arm and insert 4 pellets of solid sodium hydroxide (approximately 0.4 g) into the closed tube. Seal the tube and slowly rock it on its side in a water bath at room temperature for 20 min to absorb CO2 and re-equilibrate the temperature. Place in a vertical position, readjust the levels through C, and again read the volume (V,). The differknce (V2 - V,) is C02. Release C and introduce 0-5 g of solid pyrogallic acid by allowing it to drop through into the closed arm. Reseal; rock the tubes to absorb the oxygen; equilibrate the temperature; adjust the levels; and read the volume (V4). The difference (V3-V4) is oxygen. The residual gas will be hydrogen, carbon monoxide, or methanc, depending upon the mixture uscd for culture,

I. AUTOTROPHS

47

If the final gas analysis is compared with the initial onc, the ratio of gas oxidized to oxygen utilized can bc dctermined.

G. Organisms of Group 1 1. Group I: the culture media of Mulder aiid van Veen (1963) Basal culture solution. 27 mg I
48

V. G. COLLINS

S- or Mn-agar. I n contrast to S . natans, typical Leptothrix discophora strains, when present in the inoculum, produced many colonies on Mn-agar, even when the plates were contaminated with many other bacteria. The tiny, black-brown, hairy colonies wcre easily visible in a microscope with low magnification. Pure cultures of both S. natans and L. discophora were obtained by subculturing from parts of the colonies which were microscopically free from contamination.

6. Group I: the medium and methods of Nunley and Krieg (1967), for the isolation of Gallionella ferruginea T h e authors used the medium of Wolfe (1958). This consisted of the following method being used in conjunction with the following ingredients. Screw-cap tubes are used to confine the carbon dioxide added to the medium, and by combining ferrous sulphide with agar and slanting before the liquid medium is added. The technique is as follows. Sterile ferrous sulphide precipitate-prepared as described above-is mixed with an equal volume of sterile melted 3% water agar at 45°C. This mixture is slanted in screw-cap tubes. A liquid medium consisting of ammonium chloride, 1.0 g/litre, dipotassium phosphate, 0.5 g/litre, magnesium sulphate, 0.2 g/litre; and calcium chloride, 0.1 g/litre, is then added to the tubes. Carbon dioxide is bubbled through this medium for 10-15 sec before it is added to the test tubes, p H 6.6. T h e tubes may then be inoculated with a drop of a suspension of a Gallionella deposit from a natural source. This medium was modified by Nunley and Krieg (1967) by the addition of a pH indicator (bromothymol blue 0.001%, plus bromocresol purple, 0-0004%). Ferrous sulphide used for the submerged layer of ferrous sulphide agar was prepared by the reaction of NazS and Fe(NH&(SO& in boiling water as described by Kucera and Wolfe (1957). Before combination with the solidified ferrous sulphide agar, the fluid salts medium was cooled to 5°C to enhance the capacity of the medium for dissolved carbon dioxide; the latter was then bubbled into the fluid medium for 15 sec or until the pH had decreased to 6.0. Culture vessels were screw-capped Pyrex milk-dilution bottles (150 ml capacity) containing 10 ml of ferrous sulphide agar and either 50 ml of fluid medium (in the case of propagation of cultures) or 100 ml (in the case of the formalin isolation method). For propagation, the inoculum used per bottle was 1.0 ml of a two to three-week culture. Screw caps were tightened and then loosened a quarter turn. ' The formalin isolation procedure, developed by the above authors, is as follows: Five-tenths millilitre of formalin (40% formaldehyde solution) was added to a dilution bottle of Wolfe's medium containing 10 ml of ferrous sulphide agar and 100 ml of fluid medium. Samples of Gaffionella

49

I. AUTOTROPIIS

from its natural source wcre centrifuged at 3000 x g for 3 min. One to five millilitres of sediincnt was transferred to the Wolfe’s medium with formalin. The medium was incubatcd at 2SnC for 1-2 days. One millilitre aliquots of the culture wcrc transferred to fresh Wolfe’s medium without formalin. Cultures were incubated at 25°C for 2-3 wceks.

7. Group 1: the medium of Silverman and Lundgren (19594, for Ferrobacillus ferro-oxidans The medium is designated (910 by the authors, and its composition is as follows(NH4)aSOd KCl &HP04 MgS04.7Hz0 Ca(N03)z Distilled HzO 10 M Has04 Energy source, FeS04.7HzO

3.0g 0.10 g

0.50 0-50g 0.01 g to 700 ml 1-0 ml 300 mlofa 14.74% w/v solution

The basal salts and iron soIution are autoclaved separately and combined when cool. Some oxidation of iron occurs during autoclaving but the loss of ferrous iron is not appreciable. The medium can be stored for at least two weeks at refrigerator temperature without noticeable oxidation. The medium exhibits a precipitate (probably ferrous and ferric phosphates), is opalescent and green, has a pH of 3.0 to 3.6, and contains 9000 ppm ferrous iron. The authors have been able to maintain their standard culture of the organism on this 9K medium for more than a year by transferring twice a week into 100 ml of medium 9K dispensed in 250 ml Erlenmeyer flasks which are incubated at 28°C on a reciprocal shaker. ACKNOWLEDGMENTS

Grateful acltnowledgment is given to the following: Mrs. J. E. hl. Horne for so carefully typing the general text and references, Mrs. M. Thompson for typing of the “Media and Methods” section, Mrs. M. R. Reynolds and Mrs. S. Pywell for most able technical assistance both in the laboratory and on lake sampling; Mr. G . J. Thompson, Laboratory Steward, for assistance over many years on “field-trip” organization. Mr. J. E. M. Horne, librarian, for assistance with the references. Also to Dr. Th. E. Cappenburg of the Limnological Institute at Nieuwersluis, for constructive comments on the text, especially the description of the capabilities of the sulphate-reducing bacteria-Deszdfovibrio.

REFERENCES Anagnostidis, K., and Overbeck, J. (1966). Ber. dt. bot. Ges.,79, 163-174. Atkinson, D. E., and McFaden, B. A. (1954).J. bid. Chem., 210, 885-893. Baalsrud, K. (1954). In “Autotrophic Micro-Organisms”. Synip. SOC. gen. Microbiol., 4,5467. 4

50

V. G. COLLINS

Baalsrud, K., and Ilaalsrud, I<. S. (1954). Arch. Mzkrobiol. 20 (l), 34-62. Baars, J. K. (1930). Thesis, Meirema, Delft, The Netherlands. Bavendamm, W. (1924). Pflanzenforschung, 2, 1-1 57. Beger, H. (1949). Zentbl. Bakt. ParasitKde Abt I Originale, 154, 63,65,66. Bergey (1957). “Bergey’s Manual of Determinative Bacteriology” (Ed. R. S. Breed, E. G. D. Murray, and N. R. Smith) 7th ed. Ballikre, Tindall and Cox, London. Bissett, K. A., and Grace, J. B. (1954) I n “Autotrophic Micro-Organisms”. Symp. Sac. gen. Microbial., 4, 28-53. Bunker, H. J. (1951). Chemistry Research Report No. 3. H.M.S.O., London. Burton, S. D., and Morita, Y. (1964).J. Bact., 88 (6), 1755-1756. Butlin, K. R., Adams, M. E., and Thomas, M. (1949). J. gen. Microbial., 3,46. Campbell, L. L., Jr., Frank, H. A., and Hall, E. R. (1957).J. Bact., 73, 516-521. Campbell, L. L., and Postgate, J. R. (1965). Bact. Rev., 29 (3), 359-363. Cataldi, M. S. (1939). Revta Inst. bact., B. A r e s , 9 (I), 5-96. Cataldi, M. S. (1940). Revta Inst. bact., B. Aires, 9 (4), 393-423. Cholodny, N. (1926). P’anzenforschung, 4, 1-163. De Cicco, B. T., and Stukus, P. E. (1968).J. Bact., 95 (4), 1469-1475. Cohen, J. S., and Burris, R. H. (1955).J. Bact., 69, 316-319. Coleman, G . S. (1960). J . gen. Microbiol. 22, 423-436. Collins, V. G., and Willoughby, L. G. (1962). Arch. Mikrobiol., 43, 294-307. Collins, V. G. (1963). Proc. Sac. W a t . Treat. Exam,, 12,40-73. Colmer, A. A. (1962). J . Bact., 83, 761-765. Dorff, P. (1934). PJIonzenforschzmg, 16, 1-62. Dugan, P. R., and Lundgren, D. 6. (1965).J. Bact., 89 (3), 825-834. Duncan, D. W., Trussell, P. C., and Walden, C. C. (1964). Appl. Microbial., 12 (2), 122-1 26. Ellis, D. (1932). “Sulfur bacteria”. Longmans, Green and Co., London. Faust, L., and Wolfe, R. S. (1961).J. Bact., 81 (l), 99-106. appl. Bact., 16 (l), 1-9. Grossman, J. P., and Postgate, J. R. (1953). Proc. SOC. Hanert, H. (1968). Arch. Mikrobiol., 60, 348-376. Happold, F. C., Johnstone, K. I., Rogers, H. J., and Youatt, J. B. (1954). J.gen. Microbial., 10,261-266. Hardman, Y., and Henrici, A. T. (1939).J. Bact., 37,97. Hutchinson, M., Johnstone, K. I., and White, D. (1965). J. gen. Microbial., 41, 357. Hutchinson, M., Johnstone, K. I., and White, D. (1966). J . gen. Microbial., 44, 373. Hutchinson, M., Johnstone, K. I., and White, D. (1967).J.gen. Microbial., 47,17-23. Hutner, S. H. (1944). Archs Biochem., 3, 439. Hutner, S. H. (1946). J . Bact., 52, 213. Hutner, S. H., and Bjerknes, Clara A. (1948). Proc. SOC. exp. Biol. Med., 67, 281. Hutner, S. H. (1950). J. gem Microbial., 4, 286. Jackson, J. F., Moriarty, D. J. W., and Nicholas, D. J. D. (1 968). J . gen. Microbiol., 53,53-60. Keil, F. (1912). Beitr. Biol.Pfl., 11, 335-372. Kistner, A. (1953). Proc. K. ned. Akad. W e t . Ser. C., 56, 443. Kucera, S., and Wolfe, R. S. (1957). J . Bact., 74, 344, 349. Lackey, J. B. (1961). W a t . Sewage Wks, 15 (1). Larsen, H. (1952). J . Bact., 64, 187. Larsen, H. (1954). In “Autotrophic Micro-Organisms”. Symp. Soc. gen. Microbiol., 4,186-201. Lazoroff, N. (1963). J. Bact., 85, 78-83.

I. AUTOTROPHS

51

Lees, H. (1954). In “Autotrophic Micro-Organisms”. Symp. SOC.gen. Microbiol., 484-98. Lees, H. (1955). “Biochemistry of Autotrophic Bacteria”. Butterworths, London. Lieske, R. (1912). Bey. dt. bot. Ges., 36, 12. London, J. (1963). Arch. Mikrobiol., 46, 329-337. Lyalikova, N. N. (1958). Mikrobiologiya, 27, 556-559. Mackereth, F. J. H. (1964). J. scient. Instrum., 41, 38-41. Meiklejohn, J. (1950). J. gen. Microbiol., 4,185. Meiklejohn, J. (1952). Nature, Lond., 170, 1131. Meiklejohn,J. (1953a).J. gen. Microbiol., 8, 58. Meiklejohn, J. (1953b). J . Soil Sci., 4 , 59. Meiklejohn,J. (1954). “Autotrophic Micro-Organisms”. Symp. SOC. g m . Microbiol., 4,6843. Miller, L. P. (1950). Contr. Boyce Thompson Inst. Pl. Res. 6 (3), 85. Miller, J. D. A,, Hughes, J. E., Saunders, G. F., and Campbell, L. L. (1968). J. gen. Microbiol., 52, 173-179. Mortirner, C. H. (1940). J. Hyg., Camb., 40 (6), 641-646. Muellar, W. S., and Litsky, W. (1968). Wat. Res., 2, 289-296. Mulder, E. G., and van Veen, W. L. (1963). Antonie van Leeuwenhoek, 29,121-153. Mulder, E. G. (1964). J . appl. Bact., 27 (I), 151-173. Nunley, J. W., and Krieg, N. R. (1968). Can.J. Microbiol., 14 (4), 385-389. Niel, C. B. van (1931). Arch. Mikrobiol., 3, 1-112. Niel, C. B. van (1944). Bact. Rev., 8, 1-1 18. Overbeck, J. (1965). Zentbl. Bakt. ParasitcKde Abt. I , Suppl. 1, 13-17. Pankhurst, E. S. (1967). Lab. Pract., 16 (l), 58-59. Pfennig, N. (1961). Naturzuissenschaften, 48, 136. Pfennig, N. (1962). Arch. Mikrobiol., 42, 90-95. Pfennig, N. (1968). Arch. Mikrobiol., 63, 224-226. Phaup, J. D., and Cannon, J. (1967). Waf.Res., 1, 523-541. Phaup, J. D. (1968). Wnt. Res., 2, 597-614. Pochmann, A. (1 959). “Die Rolle der Milrroorganismen im Stoffkreislauf der Seen von S. I. Kusnezow”. Veb Deutscherverlag der wissenschaften, Berlin. Postgate, J. R., and Campbell, L. I,. (1963). J. Bact., 86, 247-279. Postgate, J. R. (1965). Bact. Rev., 29 (4), 425-441. Pringsheim, E. G. (1 949). Biol.Rev., 24, 200-245. Razell, W. E., and Trussell, P. C. (1963). J. Bact., 85, 595-603. Rhodes, M . E. (1958). J. gen. Microbiol., 18 (3), 639-648. Scotten, H. L. (1953). A. M. Thesis, Indiana University, Bloomington, U.S.A. Schatz, A., and Bovell, C., Jr. (1952). J. Bact., 63, 87-98. Silverman, M. P., and Lundgren, D. G. (1959a). J. Bact., 77, 642-647. Skerman, V. 13. D. (1967). “A Guide to the Identification of The Genera of Bacteria”, 2nd ed. Williams and Wilkins, Baltimore. Skinner, F. A., and Walker, N. (1961). Arch. Mikrobiol., 38, 339. Skuja, H. (1956). Nova Acta R. SOC.Scient. upsal., Ser. IV,16 (3). Smith, H. J., and Hoare, D. S. (1968). J. Bact., 95 (3), 844-855. Sokolova, G. A., and Karavaiko (1964). “Nauka” Publishing House, MOSCOW. Translation (1968) IPST cat. no. 1851. Sommers, L. E., and Harris, R. F. (1968). J . Bact., 95 (3), 1174. Soriano, S., and Walker, N. (1968). J. appl. Bact., 31, 493-497. Stnrltey, R. L. (1 934). J. Bact., 28. 365.

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Starkey, R. L. (1935a). Soil Sci.,39, 197. Starkey, R. I,. (1935b).J. gen. Physiol., 18, 325. Starkey, R. L. (1938). Arch. Mikrobiol., 9,368-404. Temple, K. L., and Colmer, A. R. (1951).J. Bact., 62, 605. Trimble, R. B., and Ehrlich, H. L. (1968). Appl. Microbiol., 16 ( 5 ) , 695-702. Vishniac, W., and Santer, M. (1957). Bnct. Rev., 21, 195-213. Waitz, S., and Lackey, J. B. (1958). Q. J I Fla. Acad. Sci., 21 (4), 1959. Waksman, S. A., and Starkey, R. L. (1922).J. gen. Physiol., 5 , 285. West, G. S., and Griffiths, B. M. (1913). Ann. Bot. 27 (105),83-91. Wilson, E., Stout, H. A., Powelson, D., and Koffler, H. (1953).J. Bact., 65,283-287. Winogradsky, S. (1887). Bot. Ztg, 45, 489. Winogradsky, S. (1888). “Beitrage zur Morphologie und Physiologie der Bakterien.” Heft I. Arthur Felix, Leipzig. Winogradsky, S. (1949). “Microbiologie du Sol”, h4asson et Cie, Editeurs Libraires de 1’AcadCmie de MCdicine, Paris. Wolfe, R. S. (1958). J. Am. Wat. Wks. Ass., 50, 1241-1246. Woolley, D., Jones, G. L., and Happold, F. C. (1962). J. gen. MicrobioZ., 29, 311.

CHAPTER I1

Growth of Phototrophic Bacteria and Blue-Green Algae N. G. CARR Department of Biochemistry, University of Liverpool, England I. Isolation and Elective Culture . A. Development and purificationof Myxophyccae B. Photosyntheticbacterja . 11. Maintenance of Stock Cultures . A. Myxophyceae B. Photosynthetic bacteria . 111. Chemical Environment . A. Growth media for blue-green algae . B. Growthmedia for photosyntheticbacteria IV. Physical Conditionsfor Growth . A. Blue-green algae . B. Photosynthetic bacteria . V. Physiological Effects of Varying Light VI. Apparatus . References .

. . . .

. .

.

.

. . .

. . . *

.

53 53

57 59

59 60 60 60 65 69 69 73 74 75 75

I. ISOLATION AND ELECTIVE CULTURE A. Development and purification of Myxophyceae Of the many hundred species of Myxophyceae described only a few have been obtained in pure laboratory culture. It is noticeablc that the considerable amount of physiological and biochemical work employing blue-green algae usually involves only a few species; Anabamu vuriabilis, Anabaena q’zndrica, Anacystis nidulans, Nostoc muscorum and Tolypothrix tenuis are some whose relatively rapid growth rate and ease of cultivation permit convenient biochemical examination. Macroscopic growth of blue-green colonies may sometimes be seen in streams and ponds as well as on soil surface, old brickwork, concrete guttering, etc. The extensive distribution of Myxophyceae ensures that non-specific elective incubations will usually be successful but the development of a particular spccies, and its isolation in pure culture, is a more demanding and time-consuming task. I’ringsheim (1949) has described

54

N. G. CARE

isolation procedures for blue-green, as well as other microalgae and Allen (1952) has reviewed several methods of obtaining cultures of Myxophyceae. Initial liquid cultures often involve much simpler media than are used for the growth of pure species. Beijerinck (1902) grew blue-green algae from soil in tap water +0.02% KzIIP04 and Pringsheim (1949) quotes several simple salt solutions for the growth of micro-algae generally (Table I), and has emphasized the value of soil-water cultures for isolating Myxophyceae (Pringsheim, 1950). T h e developmcnt of nitrogen-fixing species does allow greater specificity in the elective culture, since certain blue-green algae are the only organisms that will combine nitrogen in the absence of organic carbon compounds or sulphide. Tret’yakova (1965) has used several media for isolating nitrogen-fixing blue-green algae, and Bunt (1961a) has used a medium with a large content of CaC03 for this purpose (Table I). A feature of these media is the alkalinity brought about by KzHP04 or CaCO3, a p H range 7.0-8.5 being generally recommended for Myxophyceae. Oscillatoria sp. have been isolated from mud in the presence of low (0.03%) concentrations of sulphide (Allen, 1952) and sometimes occur as the terminal flora of Chlorobacteriaceae enrichments. Initial cultures may be incubated (20”-30°C) in continuous light from warm white fluorescent strips or tungsten lamps. Conical flasks, half-filled with medium and containing a small natural inoculum should be placed at varying distances from the light source. Some species of Myxophyceae are distinctly thermophilic, an example being Hupalosiphon luminosus, a strain of which, isolated by Holton (1962) from hot springs, has an optimum growth temperature of 45°C. T h e production of a uni-algal culture is usually relatively easy compared with the development of pure, bacteria-free species of Myxophyceae. Most of the examples of this Class available in culture collections are contaminated with bacteria, and some which are described as “bacteria-free” should be examined with caution as they may yield bacteria when grown on nutrient media. Pringsheim (1914) isolated three uncontaminated species of bluegreen algae by repeated transfer on solid media of either gypsum, silica or agar; modifications of this procedure have since been used by many workers. Allen (1952) isolated several filamentous blue-green algae by repeated transfer on agar plates, making use of the phototactic movements of some species. After inoculation at one side of the plate it was illuminated from the other side and motile filaments moved toward the light. After microscopic examination of the peripheral zone of growth, filaments that appeared clean were inoculated on to a new plate. Non-filamentous species were isolated after growth on pour-plates of agar, surface streaking being ineffective in separating bacteria from bliie-green algae (Allen, 1952). T h e use of micromanipulative procedures is probably the most successful method that

TABLE I Culture media for isolating blue-green algae

A

KzHPO4 KHzP04 (NH4)zHPO.l (NH4)2N03 KNO3 Ca(N03)z MgS04.7Hz0 NaC1 CaSOl CaClz. 6H2O CaC03 NaC03 FeC12 FeS04.7Hz0 NaHC03 Sodium citrate h,licro-elements PH

0.2

.. ..

.. 1.0 0.1 0.1

.. .. .. ..

.. 0.001 .. ..

..

.. ..

B

0.4

..

0.8

..

.. .. 0.4 .. 0.4

C

D

..

0.2

..

0.04 ..

0.02 .. 0.2

..

..

..

1.0

..

..

.. 0.25

.. 0.2

.. ..

0.01

..

.. .. 0.1 ..

.. 0.0005

..

..

.. .. ..

.. 0.001

.. 0*0005 ..

.. 0.001

.. ..

..

..

.. .. .. .. ..

.. ..

.. .. .. ..

E

.. 0.024

.. ..

F 0.2 ..

.. 0.116 .. ..

0.02 .. 0.20 0.2 0.165 .. 1 ml (a) . . 7.8 7.8

H

G

..

0.35

.. ..

0.25

..

1

..

..

..

0.25 ..

..

... .

0.125 0.125 I

.

..

0.15 0.20

5.5

0.20

..

.. ..

..

.. 0.0025

.. ..

1 ml (b) 1 ml (c) 8.0 7.3

S Quantities expressed as g/litre of distilled water, except A, B, C, D for which tap water W ~ used. Micro-element solutions contained in g/litre : (a) H3B03,2.86 ;ZnS04.7Hz0,0 -222;MnClz .4Hz0,1.81; NazMo04. 2H20,0.0252; CuS04.5I-I20, 0.079. (b) H3B03, 5.0;ZnS04.7Hz0,0.2; (NH4)zh!IoO4, 5.0;KI, 0.5;NaBr,0.5; x1 (so&,0 . 3 . (c) hInS04.4H20, 2.5;NazMo04.2Hz0,0.25. References: A, B, C, D, quoted b y Pringsheim (1949); E, F,G, quoted by Tret’yakova (1965); H, from Bunt, (1961a)-

U1

VI

56

N. G. CARR

does not involve physical or chemical agents that selectively kill contaminating bacteria or fungi. Taha (1963) isolated nitrogen-fixing species by repeated sub-culture on silica-gel plates. Purification of these species was assisted by treatment of the uni-algal cultures with ultraviolet light, which killed the contaminating bacteria. T h e use of such toxic agents has frequently been employed and ultraviolet treatment, introduced by Allison and Morris (1930), found to be the most successful. Cultures of Myxophyceae should be suspended with continual agitation in a quartz test tube and irradiated for 15-30 min with 275 nm ultraviolet light from a mercury-vapour lamp. Aliquots are removed at time intervals and several dilution cultures prepared from each. Prolonged irradiation kills both blue-green algae and bacteria; after very short exposure cultures may still be contaminated, but certain irradiation levels produce viable blue-green algae free from bacteria (Gerloff et al., 1950). The exact details of culture density and time of irradiation have to be established for each purification attempted. This technique is particularly valuable for isolation of species which form a heavy, external sheath, into which bacteria penetrate and are difficult to dislodge by washing. Krauss (1966) has purified ten species of blue-green algae free from contaminating bacteria by irradiation with y-rays from a 6OCo source. After treatment with 260,000 rad from a “Gammacell 220’’(Atomic Energy of Canada Ltd), bacteria were totally killed. The relative difficulty in manipulating powerful irradiation of this nature somewhat limits the general application of this method. Chemical methods for removing contaminant bacteria include the use of chlorine water (Fogg, 1942) and phenol (McDaniel et al., 1962). The latter workers purified Anacystis nidulans by exposure to 0.2% phenol in the presence of A R K 0 detergent (2 drops per 50 ml, Deko Chemical Co., Culver City, California). T h e surface-active agent was essential for success and presumably facilitated separation of blue-green algae and bacterial cells. Recently several species of blue-green algae have been purified by incubation at 47°C for 15-45 min, this technique being useful only against non-spore forming contaminants (Wieringa, 1968). Unsuccessful attempts have been made to free blue-green algae from bacteria by treatment with a range of antibiotics, although actidione and fungostatin have been effective in eliminating fungal contaminants (Tchan and Gould, 1961). An aspect of growth that has been only slightly explored is the rdle of “hormone-like” substances. I t has been reported that N . muscorum requires a “morphogenetic substance” for normal growth, and that this is available only under conditions of adequate illumination (Lazaroff and Vishniac, 1961). The interaction of bacterial Cuulobacter species and an unidentified species of Nostoc has been described by Bunt (1961b), who found that the growth of the blue-green algae was stimulated by presence

11. GROWTH OF PHOTOTROPHES

57

of the Caulobacter cells. Indole-3-acetic acid was capable of bringing about the recovery of bacteria-free homogenia that had degenerated to a colourless condition. It has been known for some time that blue-green algae often grow better in the presence of bacteria or other blue-green algae (see Fogg, 1953), and further information on this subject may well permit isolation in pure culture of hitherto impure species of Myxophyceae. If after repeated passage on growth media plus yeast extract there is no sign of bacterial development, the blue-green algal culture may be presumed pure. This is confirmed by inoculation into (1) 10% yeast autolysate, (2) 10% yeast autolysate plus 2% glucose and (3) 10% yeast autolysate, 2% glucose, 2% CaC03; if no contamination is observed after several weeks’ incubation, the culture may be judged to be pure (Allen, 1952).

B. Photosyntheticbacteria The green and purple bacteria have for many years attracted the attention of microbiologists, and considerable biochemical studies employing these bacteria, their ecological r6le and photosynthetic properties have been extensively reviewed (see Gest et al., 1963; Kondrat’eva, 1965; Pfennig, 1967). These organisms are generally divided into three families, which may be distinguished by pigment composition, nature of electron donor in photosynthesis,vitamin requirements, etc. Chlorobacteriaceae are green or yellowish-green and contain types of Chlorobium chlorophyll with in vivo absorption maxima at 725 and 747 nm. They are obligate phototrophic anaerobes and require in their medium sulphide, which is oxidized to sulphate during growth; carbon dioxide is the usual source of cell carbon but one species, Chloropseudomonas ethylicum, has a requirement for ethanol or acetate as a major carbon source (Shaposhnikov et al., 1960). The green photosynthetic bacteria are widely distributed in shallow polluted waters and mud, where visible development of species may occur when the environment is rich in sulphide; marine and estuarine species have been reported (see van Niel, 1931; Larsen, 1953). The purple sulphur bacteria (Thiorhodaceae) and the non-sulphur purple bacteria (Athiorhodaceae) both contain bacteriochlorophyll, the in vivo absorption maximum of which is 773 nm. Their reddish-brown colour is due to carotenoids. The Thiorhodaceae may utilize sulphide as an electron donor on photosynthesis and carbon dioxide, or various organic acids, serve as carbon sources. The non-sulphur purple bacteria however require reduced carbon compounds, such as maIate or succinate, as both electron donors and carbon sources, and, with one exception, are capable of aerobic growth in the dark as well as anaerobic growth in light. Most species of Athiorhodaceae, and some of the Thiorhodaceae, require certain B group vitamins. Species of purple bacteria are generally found in stagnant water,

58

N. G . CARR

especially if it contains decomposing organic matter. T h e absorption of longer wavelength light permits development of purple photosynthetic bacteria beneath a layer of green microalgae. Since water strongly absorbs in the near infrared region, inocula for these bacteria should be taken from shallow waters. Thiorhodaceae are found especially in environments rich in hydrogen sulphide and sometimes occur in salt lakes. When green algae overgrew developments of purple bacteria the use of filters that selected far-red light (700-900 nm) were employed (Scher et al., 1963; Carr, this Series, Vol. 2). Van Niel(l944) has described an elective culture medium for the Athiorhodaceae (Table 11) and has discussed procedures for eliminating ThiorTABLE I1 Elective media for photosynthetic bacteria Athiorhodaceae

Thiorhodaceae

Chlorobacteriaceae

-

(NH4)&04 (NH4)Cl K2HP04 KH2PO4 MgCh MgS04.7HzO NaCl NaHC03 NazS. 9Hz0 Yeast extract Organic substrate Fe

1.0

..

.. 1.0

0.5

1.0 1.0

..

..

1.0

..

1.0

0.5

..

0.2

2.0 5.0

..

0.1 1.5 pH 7.0

..

0-30 1 .o 1.0

2.0 1.0

..

pH 8.0-8’5

..

Quantities in g/litre of tap water. Organic substrate may be malate, succinate or fumarate.

hodaceae and green algae. T h e organic substrate (malate, acetate, fumarate, etc.) must in later cultures be supplemented with a source of B vitamins, such as yeast extract. Incubation in small completely filled glass-stoppered bottles 1 or 2 ft from tungsten bulbs will provide adequate light and heat. T h e use of particular carbon sources for isolating different species of Athiorhodaceae has been summarized by Pfennig (1967). Thus capryllate and pelargonate permit the selective enrichment of Rhodospirillum fulvum and Rhodospirillum molischianum, whereas benzoate or hydroxybenzoate have been used for isolating Rhodopseudomonas palustris. Ethanol provides the best carbon source for enrichments of Rhodomicrobium vannielii. The

11. GROWTH OF PIIOTOTROPHES

59

Thiorhodaceae may be developed in light in a medium lacking organic substrates and at a more alkaline pH (Table II), the concentration of sulphide employed prevents the development of blue-green algae (van Niel, 1931). Enrichment cultures of these organisms have been described in detail by van Niel(1944), Schlegel and Pfennig (1961) and Pfennig (1967). Wassink and Manten (1942) varied the NaHC03 and NazS concentrations, but did not succeed in isolating purple sulphur bacteria that grew faster in photoautotrophic than in photoheterotrophic conditions. T h e development and isolation of green photosynthetic bacteria has been discussed by Larsen (1953) who used a modified medium of van Niel (Table 11). I n a detailed account of his procedure, Larsen emphasized the value of using for an inoculum a pre-enrichment culture that had developed as discrete colonies on a Winogradsky column, as the numbers of purple bacteria may vastly exceed those of green bacteria in mud samples. T h e Winogradsky column consists of a mixture of equal parts cellulose, CaS04 and black mud covered with pond water. After an initial period in the dark the column is illuminated and, if successful, colonies of green and purple photosynthetic bacteria develop in the strongly reducing environment. T h e difference in p H is an important factor in the elective culture of purple sulphur and green sulphur bacteria (see Table 11). All photosynthetic bacteria may be purified by repeated anaerobic shake cultures, sealed with paraffin wax and mineral oil, and incubated in the light. After sufficient inoculum dilution, spatially separate colonies from single cells are formed and these may be dissected from the agar and transferred to fresh medium. Straightforward surface streaking onto agar plates and subsequent anaerobic incubation in the light has been used with purple bacteria (Skerman, 1959).

11. MAINTENANCE OF STOCK CULTURES

A. Myxophyceae Cultures of blue-green algae may be kept for several weeks on slopes of growth medium solidified with agar (1-2%). Some species demand liquid medium and all, generally, grow better in moist conditions, hence the frequent use of rather soft (1%) agar. It is important to use freshly prepared slopes, and if they are to be stored for more than 48 h before use they should be kept in a sealed plastic bag, or prepared in screw-capped bottles. A rather large inoculum, just visible to the naked eye, usually ensures good growth. Some species that grow poorly on solid media (e.g., Chlorogloea fritSchii) may be inoculated on to agar by adding 1 ml of a well grown liquid culture. It has been found convenient in certain cases to include in the inorganic growth agar, sodium acetate (0.01 M) and yeast extract (0.1%), thus encouraging any contaminant present in the culture to grow and be

60

N. G . CARR

detected. This procedure (used successfully with A. variabilis and An. nidulans) may partially inhibit other species of Myxophyceae, some of which are known to be inhibited by organic material in their media. Inoculated agar slopes, in boiling tubes, must be incubated at a temperature and light intensity appropriate to the species. Illumination at 30°C by 60 W tungsten bulbs, or warm white fluorescent strip lights (40 W), at 25-30 cm distance is satisfactory for many species. After growth, the boiling tubes are sealed with Parafilm (Gallenkamp Ltd, London) and stored on open shelves in the laboratory, away from direct sunlight. Several species of blue-green algae may be preserved for longer periods by rapid freezing and storage at low temperatures (Holm-Hansen, 1963), or by lyophilization (Watanabe, 1959).

B. Photosynthetic bacteria T h e Thiorhodaceae and Chlorobacteriaceae are strict anaerobes and must be maintained as stab cultures on growth media solidified with 1.5% agar, and sealed with sterile wax. All the Athiorhodaceae may be maintained on the malate-glutamate medium, supplemented with yeast extract as described by LasceIles (1956). Stab cultures in test tubes are sufficiently anaerobic for photosynthetic growth except for Rh. wannielii, which requires complete removal of oxygen by a few crystals of pyrogallol and drops of saturated potassium carbonate on top of the plug and a rubber bung inserted in the top of the tubes. After growth in front of tungsten bulbs, cultures of photosynthetic bacteria may be stored at 0°C for 4 weeks before transfer. T h e Athiorhodaceae may be stored for indefinite periods after lyophilization. 111. CHEMICAL ENVIRONMENT A. Growth media for blue-green algae T h e literature contains many media suitable for cultivating pure species of blue-green algae, and a successful recipe is often modified by different workers. This Section will describe some of the general media which various workers have found suitable for several species of Myxophyceae. A brief account will be given of some of the special media suitable for a particular species, or group of species. One of the earlier successful general culture media was that described by Chu (1942) as No. 10, this being an ionically dilute solution that included Feels. It was found that an organic source of iron increased the availability of that metal and improved the growth of cultures. Gerloff et al. (1950) described a modified Chu No. 10 medium (Table 111) in which they grew eight species of pure Myxophyceae. A feature of this medium is the maintenance of a relatively high p H ; the growth rate of N . museorurn fell

TABLE I11 Growth media for blue-green algae

Gerloff et al. Provasoli et al. Kratz and Myers (1950) (1957) (1955)

KN03 Ca(NO& KzHPOi MgS04.7Hz0 NaCl Na2C03 NaHC03 NazSiOa .9Hz0 NaN03 Fe2(S04)3 .6H20 KCl CaClz.6HzO TRIS EDTA VitaminBiz Micro-elements (see Table IV) Sodium citrate Ferric citrate Citric acid

..

ASP 2 base

0.04 0.01

..

.. 0.025

..

..

0.005 5.0 18.0

0.02

.. ..

0.025

0.150 0.05

.. .. .. ..

.. ..

..

C 1.0 0.025 1.00 0.25

.. ..

..

0.003 0.003

1.Autoclavedseparately. Salts are dissolved in 1 litre of glass-distilledwater.

..

D

M

0.01 1.00 0.15

..

.. ..

.. ..

0.04 0-25

..

.. 0.2t

.. ..

1-00 0.004

0.6 0.3 1.0 0-03 2.0 LCg “PMD”

Taha Allen and Amon (1963) (1955)

.. .. .. ..

A;(1 ml) 0-165

..

..

0-004

0.02

.. ..

0 * 024

..

.. ..

0.358 0.25 0.232

..

.. .. ..

.. ..

0.05 Hs (1 ml)

..

..

..

A,(1 ml)

0.11

.. ...

..

A4+B7 0.165

..

.. ..

..

I

0 P

0

5X

62

N. G. CARR

markedly when the p H of the medium was lowered to 7.1. It was apparent that the function of silicate and carbonate in the medium was one of pH control, rather than as acting as specific nutrients. Taha (1963) found that good growth of three nitrogen-fixing species was obtained with medium M, and Allen and Arnon (1955) report high growth rate for A. cylindrica and other nitrogen-fixing blue-green algae (Table 111). T h e latter medium was prepared with especially pure materials, and although not all the trace elements included were known to be essential, it was shown that A. cylindrica had a requirement for Ca and Mo that could not be replaced by other elements. T h e work of Allen and Arnon has stressed the desirability of using purified salts, and conditions of vigorous growth, for examination of mineral nutrients. T h e major variable open to studies on blue-green algae nutrition is the supply of nitrogen. T h e media of Taha and of Allen and Arnon described in Table I11 may be supplemented with various nitrogen sources. T h e latter workers suggest that about 0.2 g/litre of nitrate ion is necessary to adequately supply a rapidly growing culture, Nitrogen may be supplied as nitrate, nitrite or ammonia and the presence of such ions in the medium will prevent nitrogen fixation. Taha (1963) has reported that Calothrix elenkinii will not use nitrate as a source of nitrogen and continues to fix atmospheric nitrogen in its presence. Allen (1952) reports the widespread use of casein hydrolysate as a nitrogen source, although there is less information on the use of organic nitrogen by blue-green algae in chemically defined media. Urea has been used as sole nitrogen source by several species, e.g., A. variabilis and N . muscorum (Kratz and Myers, 1955) and A. cylindrica (Cobb and Myers, 1961). T h e latter workers showed that the growth rate of A. cylindrica increased with nitrogen, nitrate and urea, in that order. Van Baalen (1962) has found that the ASP-2 medium of Provasoii et al. (1957), containing vitamin BIZ(2 pgllitre) and no other vitamins, adequately supported growth in 15 species of marine blue-green algae. On examining the salinity requirement, Van Baalen (1962) found optimum growth of three species between 1-0-2-0% NaCI. T h e media that have probably been found to be most generally useful in Myxophyceae growth studies are those of Kratz and Myers (1955), two of which are described in Table 111. Medium C proved especially favourable to a range of blue-green algae and has the advantage of being relatively simple and, except immediately after autoclaving, is precipitate free. T h e sodium citrate functions as a chelating agent, and may be replaced by EDTA. T h e micro-element constituents were not accurately measured and Kratz and Myers (1955) observed that they could be varied without observable effects on growth. More than one group of workers has supplemented medium C with 0.05% NaHC03, which may be autoclaved with the medium

11. GROWTH OF PHOTOTROPHES

63

and is reformed after gassing with air-CO2 mixtures (Carr and Hallaway, 1965; Hoogenhout and Amesz, 1965). T h e supply of C 0 2 to the growing culture is best maintained by a flow of air-COz (95-5 v/v), although other proportions have been used with success; pure nitrogen may also be used in place of air. Since blue-green algae produce oxygen during photosynthesis, strict anaerobic conditions cannot be maintained. It is necessary to measure the pH of a medium after gassing with the mixture employed, as the proportion of C02 contributes to the final pH. Provided medium C is aerated sufficientlyvigorously air alone can supply sufficient COz for A . variabilis to grow to 1.0 mg dry wtlml. However, such rapid aeration and agitation considerably increases the evaporation and foaming of cultures in medium C, and gassing with an air-COz mixture is to be preferred. Considerable attention has been directed to laboratory growth of those species of blue-green algae that are toxic to man and animals, Microcystis aeruginosa has been grown on defined medium in which tris(hydroxymethy1)aminomethane was used as a buffer at pH 7.0 (McLachan and Gorham, 1962). The large-scale production of this organism has been described by Gorham (1964) whose "algae factory" produced up to 450 litres per day. The thermophilic blue-green algae are perhaps easier than most to isolate, and Holton (1962) describes the growth characteristics of H. laminosus, with an optimum temperature of 45"-5O"C, in a medium in which the initial pH of 7.0 was produced by K2HP04, Na2C03 and Na2Si03.9HzO. T h e medium described being better than those of Allen and Arnon and Kratz and Myers (Table 111) for this organism. An extremely rapid growing Synechococcus lividus has an optimum temperature of 52"C, at which it has a doubling time of 2.6 h (Dyer and Gafford, 1961) when growing on a defined medium containing tris. The r81e of organic materials in the nutrition of Myxophyceae is not clear. Most species demand C02 as the major growth material, but many grow better in the presence of small quantities of a wide range of organic materials. Allen (1952) considers that these may be facilitating the availability of various inorganic ions, rather than acting as nutrients in the normal sense. Recently the incorporation and accumulation of 14C-labelled glucose and acetate by A. vuriubilis and An. niduluns has been demonstrated (Hoare and Moore, 1965; Carr and Pearce, 1966; Pearce and Carr, 1967) and these organisms may no longer be considered strict photoautotrophs. A few species however are heterotrophs and will grow in the dark on reduced carbon compounds. N . muscorum will grow and fix nitrogen in the dark on a mineral medium plus glucose (Allison el al., 1937) as will Chlorog.fritschii in the presence of sucrose (Fay, 1965). Both these organisms grew extremely slowly and experiments were conducted over periods of many weeks. T. tmuis on a medium containing-

64

N. G . CARR

5.0 g 10.0 g 0.5 g 0.25 g 0.02 g 0.02 g 1 ml 1 litre

Casaminoacids Glucose MgS04.7HzO KzHPO4 CaClz.2Hz0 FeS04.7H40 Agmicro-elementsolution (see Table IV) Distilled water

grew in the course of 10 days when incubated in the dark at 32°C. This being the most rapid, heterotrophic growth of blue-green algae known (Kiyohara etal., 1960). TABLE IV

Trace-element solutions Allen&Amon(1955) Kratz&Myers(1955) Provasolietal. (1957 A5 H5 “PMD” A4 f H7 ppm metal g/litre Metal ion (as C1-) per 100 ml MnClz .4&0 MnS04.4Hz0 Moo3 ZnS04.4H20 CuS04.5HzO HsBOs NHaVOa CO(NO3)2.6HzO NiS04.6HzO Crz(SO)sK2SO4.24H20 NazWO4.2HzO TiO(Ca04)r. yHzO Fe (as EDTAcomplex) FeCls

.. 0.50 0.10 0.05 0.02 0.50 0.01 0.01 0.01 0.01

0.01 0.01 4.00

..

1 *81

1a 4 4

0.0177 0.222 0 * 079 2.86

0.71 8.82 1.57

..

..

..

..

.. ..

0.49

..

..

.. ..

.. ..

..

0.12 m g

.. ..

15.0 p g 0.12 p g 0.6 mg

.. ., .. ..

0.3

*. 0.08 mg

There are several organisms described in the literature as “colourless Cyanophyta”, which are pigment-less species of blue-green algae. The possible phylogenetic relationship between these and the flexibacteria has been discussed (Soriano and Lewin, 1965). Media that have proved successful for the growth of heterotrophic colourless Cyanophyceae, and the details of incubation and cultivation have been described: Saprospira sp. and Flex& bacter sp. (Holm-Hansen et al., 1965), Leucothrix mum and Vitreoscilla sp. (Webster and Hackett, 1966) and Beggiatoa (Scotten and Stokes, 1962).

11. GROWTH OF PHOTOTROPHES

65

B. Growth media for photosyntheticbacteria 1. Athiorhodaceae

The ready growth of these bacteria in laboratory culture and their capacity for photoheterotrophic and aerobic heterotrophic growth have contributed to their popularity with microbial biochemists. The media described will support either type of culture of species of Athiorhodaceae other than Rh. vannielii, which grows only under anaerobic photosynthetic conditions. Good growth with several species (Rhodopseudomonascapsulata, Rhodopseudomonasgelatinosa, Rhodopsnidomonaspalustris, Rhodospirillum rubrum and Rh. vannielii) has been obtained using the malate-glutamate medium developed by Lascelles (1959) for Rhodospeudotnonasspheroides (Table V). Not all the vitamins are necessary for each micro-organism (see Table VI) and Rhodops. palustris requires the addition of p-aminobenzoic acid (137 pg/litre). Glass-distilled water should be used to which the organic substrates are added first and the vitamins last; pH 6.8 is achieved by adding NaOH. The medium may conveniently be made double-strength and stored at 0°C with a few drops of toluene before sterilization. In order to produce cultures growing at maximal rate, modifications are necessary for different species of Athiorhodaceae; Rhodops. capsulata grows better, for example, when malate is replaced by succinate. Media specially designed for the growth of Rhodosp. rubrum (Ormerod, et. al., 1961) and Rhodops. palustris (Keane et al., 1963) have been described (see Table V). The later workers report growth yields some four-fold better than those obtained with a malate-glutamate media, but do not comment on the rate of growth. Rh. vannielii differs from the other non-sulphur purple bacteria in several respects, one of which is its nutrition. It does not require any vitamins in the growth medium and thrives on alcohols. The chemically defined medium described (Table V) comprises the basal salts of Lascelles malate-glutamate medium plus propanol and NaHC03, these being added after membrane sterilization. Other substrates (e.g., ethanol, butanol, lactate, pyruvate) may replace propanol, but substituted alcohols do not serve as substrates. Rh. vunnieZii grew well on malate-glutamate medium (Table V) and does not, on this medium, require NaHC03. The nature of the carbon source affectsthe level of bacteriochlorophyll (Kornberg and Lascelles, 1960) and of ubiquinone (Carr and Exell, 1965) in Athiorhodaceae. Growth on acetate produced a several fold decrease of both compounds when compared with malate-grown organisms. The vitamin requirements of the Athiorhodaceae have been examined by Hutner (1946) and Table IV summarizes those needed by seven species. A new species, Rhodopseudomonas isachenkoi, isolated from water layers 1.367 m deep, bclow petroleum deposits, also

TABLE V Growth media for some Athiorhodaceae

Rhodopseud-s spheroides (Lascelles, 1959)

Rhodospirillum Rhodopseudomonas Rhodomicrobium rubrum palustris vamielii (Ormerod et aE., (Bose, 1963, after (N. G. Carr, unpublished data) 1961) Keane et al., 1963) _

KH2PO4 K2H Po4 (NH4)2HP04 (NH4)zS04 NaHC03 MgS04.7H20 MnS04.4HzO CaC12.2Hz0 FeS04.7H20 Iron citrate,t 4 m~ Glycerophosphate(Na) Citrate (K) Glutamate (Na) L-Glutamic acid DL-Mahc acid Acetate (Na) Propanol L-Histidine

.

0.500 0-500 0.800 ..

..

0.600 0.900

.. ..

0.50-1 '25

0.040

.. ..

.. ..

0.200 2-23 mg

0.200

0-600

0.040

0.075

0.150

0.012

..

.. Id

..

..

..

_

_

_

~

0.500 0.500 0.800

..

1.00 0.200 2-23 mg 0.040

..

1 ml

..

..

.. ..

..

..

3 -00 4.008 0.200

..

..

2.00

..

..

2.70

.. 6.00

.. .. ..

..

_

2.00 0-500

..

.. 1-90

..

..

_

..

..

..

0.50

..

TABLE V-continued Rhodopseudomonas Rhodospirilhm Rhodopseudomonas Rhodomicrobium spheroides rubrum palustris vannielii (Lascelles, 1959) (Ormerod et at., (Bose, 1963, after (N. G. Carr, unpublished data) 1961) Keane et al., 1963)

Tyrosine Monobutyrin Homocysteine thiolactone-HCl EDTA Micro-elements Nicotinic acid ThiamineHCI Biotin p-Aminobenzoic acid PH

.. .. ..

.. ..

1 mg 1 mg 10 f %

..

6-8

.. ..

.. 0.020 1 mll

.. ..

15 Pg

..

6.8

0.100

0.400 0.200

..

see below7

.. .. ..

200 Pg 6.2-6.5

..

., .. .. ..

..

.. .. ..

..

Except where indicated, all quantities are in g/litre of distilled water. t Iron citrate: (NH4)zFe (SO&. 6Hz0, 157 mg and sodium &rate (&hydrate), 236 mg dissolved in 100ml distilled water and stored at 0°C. $ Per 100 ml de-ionized water: HsB03, 280 mg; MnS04.4HzOy21Omg; NazMoOr. 2Hz0,75 mg; ZnS04.7H20, 24 mg; Cu(NOs)~.3H~0,4rng. 8 Neutralized separately with KOH. 7 Am& containing (mg/litre) Fe, 18: Mn, 14; Zn, 9.0; Mo, 1.8; Cn, 0 - 9 ; Co, 0.18; B, 0.18; V, 0.18; 1,0.18; Se, 0.036.

8

68

N. G . CARR

TABLE VI

Vitamin requirements of Athiorhodaceae Organism

Rhodospirillum rubrum Rhodopseudomonas spheroides Rhodopseudomonas capsulata Rhodopseudomonas gelatinosa Rhodopseudomonas palustris Rhodomicrobiunt vannielii R hodopseudomonas isachenkoi

Biotin Thiamine

-I-

+ +

Nicotinic acid p-Aminobenzoicacid

-

-

i-

4-

i-

+ +

-

-

-

-

-

-

-

-

-

-

4-

does not require vitamins when growing on malic acid and mineral salts (Doman et al., 1962).

2. Thiorhodaceae These bacteria will grow photoautotrophically (with NaHCO3 as sole carbon source) or photoheterotrophically on various organic acids, malate being a favoured substrate. Growth is more rapid in the presence of an organic substrate, and either type of culture contains sodium sulphide, which acts as both an electron donor and as a deoxygenating agent. The medium of Eymers and Wassink (1938) provides a simple, but not entirely defined, solution for growth, as various trace elements are supplied by the tap water (Table VII). Autotrophic conditions are provided by Hendley’s (1955) medium, cultures in which are continually gassed with C02 and oxygen-free nitrogen (4-96, vlv) ; the indigo carmine serves as an indicator for the absence of oxygen, and may be omitted if desired. A fully defined heterotrophic medium is described by Bose (1963) who comments that reliable and reasonably rapid growth required large inocula (10-20%). Several workers have noted that calcium ions appear to be important in maintaining growing cultures in suspension, and that greater concentrations to those described are sometimes necessary. The media so far mentioned are suitable for small species of Chromatium, such as Chromatium minus and including the frequently studied Chromatium D.Schlegel and Pfennig (1961) and Pfennig (1961) describe a carefully developed chemically defined

11. GROWTH OF PHOTOTROPHES

69

medium that pcrmits the cultivation of pure cultures of “large” species, such as Cliromutium olzcnii and Chromatiurn wurmingii (Table VII). The final pH was varied according to species, and was more acidic (pH 6.6-7.2) than usual for Thiorhodaceae. A feature of these workers’ media was the inclusion of vitamin BIZ; before this, no vitamin requirements had been observed for the Thiorhodaceae. Recent modifications introduced by Pfennig (personal communication) are included in the medium described in TableVII.

3. Chlorobacteriaceae The media used for culturing green photosynthetic bacteria derive mainly from the work of Larsen (1953) who showed that the species available at that time (Chlorobium thiosulfatophilum and Chlorobium limicola) grow exclusively on inorganic material (Table VIII). The former bacteria will grow with Na2S or Na~S203as electron donors, while the latter will use only Na2S. Larsen (1953) observed that Chlor. thiosulfatophilum is much more sensitive to NazS203 concentrations than the non-photosynthetic sulphur bacteria, 0.4% causing distinct decrease in growth. Larsen’s media should be made in distilled water and the pH adjusted to 7.0-7-5 with H3P03,and the NaHC03, NazS and FeCl3 should be sterilized separately. More rapid growth was obtained if the medium (less FeC13) was stored in glass bottles for 24 h before inoculation, just before which the FeCl3 was added. It was suggested that this “aging” of media allowed the NazS completely to remove small amounts of dissolved oxygen. If thiosulphate is used as an electron donor, 0.1 gjlitre of Na2S is added to ensure anaerobiosis. The isolation of Chlorops. ethylicum, which has an obligate requirement for C02 and ethanol as carbon sources, provides an example of an anaerobic, heterotrophic green photosynthetic bacterium (Shaposhnikov et al., 1960). This may be grown on a medium described by Bose (1963), which was based on Larsen’s medium for green photosynthetic bacteria (TableVIII), the ethanol may be replaced by sodium acetate (0.034 M) and additional buffering provided by tris (0.1 M, pH 7.3) (Callely and Fuller, 1967).

IV. PHYSICAL CONDITIONS FOR GROWTH For definitions of the quantitative terms of light measurement see Carr (this Series, Vol. 2).

A. Blue-green algae The emissions from both tungsten bulbs and fluorescent striplights are suitable for growing blue-green algae. Cultures may be placed 20-30 ern

TABLE VII

Growth Media for Chromatium species Hendley (1955) Eymers and Wassink (1938) Distilled water, autotrophic T a p water, heterotrophic g/li tre g/litre --______

Sodium malate NazS203 (NH&S04 NaCl K2HPO4 MgSO4

t 10% NazS t 10% NazC03 t 10% H3P03

-.-

2.4 1.6 1.0

20.0 0.5 0.2 pH 7 - 4

18 ml 9 ml 6 ml

NaCl NH4Cl KH2PO4 MgC12.6H20 CaCl Trace elementsa Indigocarmine Conc. HCl

Na2C03 NazS203. SHa0 NazS. 9H20 EDTA (di Na)

Bose, 1963 (after Fuller) Distilled water, heterotrophic g/litre

Pfennig (modified after Pfennig, 1961)

.~

Solution 1 Distilled water CaClz. 2H20 Solution 2 Distilledwater 20.0 . Heavy metal solutionC VitaminBiz KH2P04 (120 ml) diluted to KC1 800 ml NH4Cl MgClz .6H20 Solution I1 M Tris, pH 7.5, Solution 3 Distilled water 50 ml NaHC03 Solution 111 KzHPOa(1 g) in50ml Solution4Distilledwater NazS.9Hz0 B Solution 1V NazS .9Hz0 (0.29) in 50 ml Solution V 5 * 5 g organic carbon source (neutralized) in 50 ml CaC12 NH4CI MgS04.7H20

9.0

'z: 1 0.1 0.2

Stock 0-66 solution

5000 ml 2 -0 g 40 ml 60 ml 0.12mg 2.0 g 2.0 g 2.0 g 2 .O g 900 ml 9g 400 ml 6g

t Added after sterilization

Solution A and B are sterilized separately and mixed in

equal volumes after cooling giving a final pH of 8.0.

Solutions I-V were autoclaved separately and mixed when cool;pH adjusted to 7 . 5 with conc. HCI.

The sterilization and addition of these

solutions are described below (4.

(a) Larsen's trace elements (see Table VIII).

(b) Dissolve in 750 ml distilled water, EDTA, 5Og; ZnS04.7H20,22g; H3B03,11.4 g; MnC12.4Hz0, 5.1 g ; FeS04.7Hz0, 5.0 g; CoCh.6Hz0,1.6 g; CuS04.5Hz0,1-6 g; (NJ&)&f0@~4.4H20,1.1 g. Boil and after cooling bring to pH 6 - 5 - 6 . 8 with KOH make to 1 litre final volume. (c) Heavy metal solution. T o 1 litre of distilled water containing EDTA disodium salt (500 mg) were added FeS04. 6HzO, 200 mg ; ZnS04.7Hz0, 10 mg; MnC12.4H20, 3 mg; H3B03, 30 mg; CoC12.6H20, 20 mg; CuC12.2Hz0, 1 mg; NiC12.6Hz0, 2 mg'; NaMOO4.2H20; 3 mg. ( d ) Solution 1 (105 ml) was placed in a 135 ml screw-capped flask and autoclaved with loose caps. Solution 3 was flushed with co2 for at least 30 min while being vigorously stirring, until the pH is below 7.0 and then mixed with solution 2. The mixture was sterilized by passage through a washed Seitz filter under a pressure of 15psi ( 2 0 2 , and aseptically added (22 ml aliquots) to the sterile cold solution A. Solution 4, after autoclaving, was added to the flasks in 4 ml (giving pH7 .O)to 6 .O ml (giving pH 7 2) amounts. The flasks were filled with sterile 0 -04% CaCl2 solution. 3

72

N. C . CARR

TABLE VIII Growth media for green photosynthetic bacteria _

_

~

~

Chlorobium thiosulfatophilum and Chlorobium limicola (Larsen, 1953) g/litre I _

NH4Cl KHzPO4 MgClz NaCl TNaHCOs NazSzO3 t o r NazS .9H@ CaClz tFe(FeCls.6HzO) B (H3B03) Zn (ZnSo4.7Hzo) Co(Co(NOs)s.6H20) Cu (cuso4.5Hz0) Mn (MnC12.4HzO)

Chloropseudomonus ethylicum (Bose, 1963)

..-__

1.0 1.0 0= 5 10.0 2.0

KHzPOa NH4C1 MgClz .6Hz0 NaCl CaClz Trace elements

i:::

10.0 g

40.0g 80mg 2.0 ml

1

Basal salts, per500 ml

1.0 0.1 5OOpg 100 p g 100 pg

50 p g 5 pg 5 pg

FeC13 NazB.107. lOHzO ZnS04.7H20 cosO4.7Hzo CuClz. 2Hz0 MnS04.HzO Versenol iron solution$ 110 ml Mix 5 I. Basal salts 500 ml distilled water 450 ml 11. 10% Naz S.9Hz0 2 ml 111. 10% NaHC03 40 ml IV. 0.05% FeS0.7HzOin 5 ml 0.3 N HCI V. 70% ethanol 3 ml

+Sterilized separately 1Versenol iron solution: 59 g EDTA are dissolved in 500 ml distilled water and FeS04.7H20 (24.9 g) added. Dilution to 1 litre and aeration overnight yielded a final pH of 9 * 7. $ Solutions I-IV were sterilized separately and mixed after cooling. Ethanol was sterilized by membrane filtration and the complete medium adjusted with cone. HCl topH7.3.

distant from several parallel 20 W strip lamps (giving 200 Im/sq. ft at culture) or approximately the same distance from a bank (3 or 4) of 60W bulbs (yielding 240 lm/sq. ft at culture). It has been found convenient to stand vessels (Roux bottles or medicinal flats) in a glass tank (30 x 50 x 25 cm) and illuminate from lights fixed in a reflector hung on to the side. Large quantities (10 litres) may be grown in a hot-room in spherical flasks, around the equator of which is hung a 60 W circular strip lamp. T h e degree of illumination supplied varies considerably between workers, Gerloff et al,

11. GROWTH OF PIIOTOTROPHES

73

(1950) used 40 ft cd for several species and Dyer and Gafford (1961) used 1500 ft cd for Syn. lividus. Data from Kratz and Myers (1955) indicates that, under their conditions, the approximate optimum light intensity for growth of A . variabilis was 250 ft cd; An. nidulans, 330 ft cd and N . muscorum, 260 ft cd. Considerably higher light intensities (7000-15,000 m cd) were employed by Hoogenhout and Amesz (1965). The growth of a strain of An. nidulans (originally from Dr. M. 13. Allen) was inhibited by light intensities greater than 700 Im/sq. f t at 31'C (N. G. Carr, unpublished results). Many species of blue-green algae grow at temperatures between 30" and 35"C, and there are thermophiles, already mentioned, whose optima are considerably higher. Microcyst. aeruginosa grew well at 23"C (McLachlan and Gorham, 1962) and T. tenuis grew both heterotrophically and photosyntheticallyat 32°C (Kiyohara et al., 1960). The proportions of saturated to unsaturated fatty acids in A n . nidulans increased when the growth temperature was raised from 35" to 41°C (Holton et al., 1964). An important aspect of culturing blue-green algae is the problem of keeping the organisms in a uniform suspension during growth. Most species (An. nidulans is an exception) tend to clump together and settle, or stick to the surface of the flask. Agitation by gently rocking the cultures while illuminated from overhead minimizes these problems, as does stirring with a Teflon-covered magnetic stirrer. The flow of gas through a culture, especially if in a tall, thin vessel, may materially assist in keeping the organism suspended. Blue-green algae may be gassed with any mixture of air and CO2 up to lo%, and the changes in pH caused by high concentrations of C02 may be buffered with NaHCOs, 0.5 g per litre. Gusev (1962) suggests that an artificial diminution of oxygen concentration, especially during early stages of growth, was beneficial.

B. Photosyntheticbacteria All species of Thiorhodaceae and Chlorobacteriaceae require strict anaerobiosis for growth. In test-tube cultures, this may be achieved by a pyrogallol seal or a plug of sterile wax (see Section I, and Willis, this Volume, p. 79). Larger volumes are grown in completely filled glass bottles sealed with a ground-glass stopper or continuously flushed with an appropriate gas phase; screw-capped jars may also be used. A Roux bottle with a B29 ground-glass socket joined and annealed to its neck provides a reasonably shallow, flat-surfaced vessel. The Athiorhodaceae typically require only semi-anaerobic conditions for photosynthetic growth and Roux bottles, or medical flat bottles, three quarters full of medium are adequate. Rh. vannielii requires strict anaerobic conditions and should be treated like the photosynthetic,purple bacteria. Photosynthetic bacteria must be illuminated with tungsten bulbs, fluorescent lamps producing insufficient emission in

74

N. C. CAHR

the near infrared region. Athiorhodaceae and Thiorhodaceae will grow at intensities of 200-300 ft cd, i s . , some 10 in. from a row of 60 W bulbs. Green photosynthetic bacteria usually demand less illumination and optimum growth is obtained at intensities lower than 100 ft cd; several strains were grown by Cohen-Bazire et al. (1964) at not more than 40 ft cd. It is difficult to control entirely the illumination actually received by a culture, factors such as reflection may cause significant alteration in light intensity within a growth vessel. The growth of the bacterial culture itself, from a faintly turbid solution to a deeply pigmented thick suspension clearly causes continuous decrease in the light received by each microbial cell. If reliable growth is not obtained the possibility of over-illumination should be considered, especially of cultures in the early stages of growth. Photosynthetic bacteria grow at temperatures between 20"-30°C ; some species, (e.g., Rhodops. spheroides) will grow at 34"C, whereas others (e.g., Rh, vannielii) are particuIarIy sensitive to temperatures in excess of 30°C. It is usually necessary to dissipate heat produced by the lamps with a fan that is linked either to an air thermostat or a thermostat in a "control" vessel filled with water. T h e typical Athiorhodaceae will grow in the dark under aerobic conditions, and provided the aeration is sufficient, pigment synthesis is suppressed. Growth on malate-glutamate medium (500 ml) in 2 litre Erlenmeyer flasks agitated in a rotary shaker (New Brunswick Scientific Co., or Gallenkamp Ltd) at 100 rev/min at 30°C yielded unpigmented organisms with growth rates comparable to those obtained under photosynthetic conditions. Smaller cultures have been grown in inverted T tubes of about 50 ml capacity clamped to a rocking device that moves through a 90" arc in a water bath (Lascelles, 1956). Excess aeration will inhibit the aerobic growth of Athiorhodaceae, and 3-5 mm Hg was found to be the optimum partial pressure of 0 2 for Rhodosp. rubrum (Biedermann, et al., 1967).

V. PHYSIOLOGICAL EFFECTS OF VARYING LIGHT T h e most marked effect of variation of light intensity other than considerations of growth, is on the concentrations of photosynthetic pigments within the microbial cell. T h e levels of bacteriochlorophyll and carotenoids vary inversely with light intensity. This has been examined in detail by Cohen-Bazire et al. (1957), who showed that when organisms were transferred from high to low light intensity the rate of pigment synthesis relative to cell mass synthesis increased. The bacteriochlurophyll content ofR4we.r. qheroz2es is some sevenfold higher after growth in light intensity of 50 ft cd than after growth at 5000 ft cd. A similar result has been noted for Chlorops ethylicum (Holt et al., 1966) and for Rh. vannielii (Trentini and Starr. 1967)

11. GROWTH OF PHOTOTROPHES

75

In the latter organism, ultrastructural changes occur as light intensity varies, growth in weak light increases the total amount of intracellular lamellae and localizes this around the periphery of the cell. Sistrom (1962) has shown that the bacteriochlorophyll content of lamellae is determined by the illumination operative at its biosynthesis; thus adjustment of bacteriochlorophyll content must take place in conjunction with increased or decreased rate of lamellae formation. The changes in ratios of chlorophyll, carotenoids and phycocyanin with varying light intensity have been examined in the blue-green algae, An. nidulans (Myers and Kratz, 1955). A threefold decrease in chlorophyll and a fourfold decrease in phycocyanin was observed after growth under higher illumination ;a much smaller change was noted in carotenoid pigments. The ratio of phycoerythrin to phycocyanin formed by T.tenuis alters in response to the spectrum of illumination, fluorescent lamps increased twenty-fold the level of phycoerythrin over that produced in tungsten light (Hattori and Fujita, 1959). The proportion of COz to ethanol assimilated by Chlorops. ethylinrm during growth increased when light intensity was altered from 50-7000 lux (Balitskaya and Kondrat’eva, 1963) and the use of acetate and glucose is differentially affected by alteration in luminous flux (Shaposhnikov and Balitskaya, 1964). VI. APPARATUS There are many modifications to laboratory glassware that have been useful in culturing photosynthetic microbes. The two most general problems to be solved are providing a uniform degree of illumination to all the culture and the elimination of “settling” of cultures by some form of mechanical agitation. Myers (1960) has described a conical flask altered to increase the effectiveness of gassing, and a simple test-tube procedure for growing a series of cultures (see also Hoogenhout and Amesz, 1965). A cone-shaped base to a boiling tube, and a gassing tube that almost touches the bottom of the tube assist in keeping blue-green algae in suspension. Incubated gyrorotary shakers that are illuminated by fluorescent lamps are suitable for some blue-green algae (New Brunswick Scientific Co. ; Gallenkamp Ltd). A continuous-culture apparatus designed for micro-algae generally (Myers, 1960) and a similar apparatus for anaerobic cultures of Rhodosp. rubrum (Hendley, 1955) have been described. REFERENCES Allen, M. B. (1952). Arch. Mikrobiol., 17, 34-53. Allen, M. B., and Arnon, D. I. (1955). Plant Physiol., 30, 366-372. 71,221-223. Allison, F.E., and Morris, H. J. (1930). Science, N . Y.., Allison, F. E., Hoover, S.R . , and Morris, I f . J . (1937). Hot. Guz., 98, 433-463.

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Balitskaya, R. M., and Kondrat’eva, E. N. (1963). Mikrobiol., 32, 167-171. Beijerinck, M. W. (1902). PYOC. I(. ned. Akad. Wet., 4, 5-9. Biedermann, M., Drews, G., Marx, R., and Schroder, J. (1967) Arch. Mikrobiol., 56,133-147. Bose, S. K. (1963). In “Bacterial Photosynthesis” (Ed. H. Gest, A. San Pietro and L. P. Vernon), pp. 501-510. Antioch Press, Yellow Springs, Ohio. Bunt, J. S. (1961a). Nature, Lond., 192, 479-480. Bunt, J. S. (1961b).Nature, Lond., 192,1274. Callely, A. G., and Fuller, R. C. (1967). Biochem. J., 103, 7 4 ~ . Carr, N. G., and Exell, G. (1965). Biochem.J., 96,688-692. Carr, N. G., and Hallaway, M. (1965).J. gen. Microbiol., 39, 335-344. Carr, N. G., and Pearce, J. (1966). Biochem. J.,9 9 , 2 8 ~ . Chu, S. P. (1942). J. Ecol., 30, 284-325. Cobb, H. D., andMyers, J. (1961). Am.J. Bot., 51,753-762. Cohen-Bazire, G., Sistrom, W. R., and Stanier, R. Y. (1957). J. cell. comp. Physiol., 49,25-68. Cohen-Bazire, G., Pfennig, N., and Kunisawa, R. (1964).J. Cell. Biol., 22,207-225. Doman, N. G., Romanova, A. K., and Terent’eva, 2. A. (1962). Mikrobiologiya, 31,157-161. Dyer, D. L., and Gafford, R. D. (1961). Science, N. Y.,134,616-617. Eymers, J. G. and Wassink, E. C. (1938). Enzymologia, 2, 258-304. Fay, P. (1965).J.gen. Microbiol., 39,ll-20. Fogg, G. E. (1942). J. ex$. Biol., 19, 78-87. Fogg, G. E. (1953). “The Metabolism of Algae”. Methuen, London. Gerloff, G. C., Fitzgerald, G. P., and Skoog, F. (1950). Am.J. Bot., 37, 216-218. Gest, H., San Pietro, A., and Vernon, L. P. (Eds). (1963). “Bacterial Photosynthesis’’. Antioch Press, Yellow Springs, Ohio. Gorham, P. R. (1964). In “Algae and Man” (Ed. D. F. Jackman), pp. 307-336. Plenum Press, New York. Gusev, M. V. (1962). Dokl. Akad. Nauk SSR,147,947-950. Hattori, A., and Fujita, Y. (1959).J. Biochem., Tokyo, 46, 633-644. Hendley, D. D. (1955).J. Bact., 70, 625-634. Hoare, D. S. and Moore, R. B. (1965). Biochem. biophys. Acta, 109,622-625. Holm-Hansen, 0. (1963). Physiologiu PI., 16, 530-540. Holm-Hansen, O., Prasad, R., and Lewin, R. A. (1965). Phycologia, 5, 1-14. Holt, S. C., Conti, S. F., and Fuller, R. C. (1966).J. Bact., 91, 349-355. Holton, R. W. (1962). Am.J. Bot., 49, 1-6. Holton, R. W., Blecker, H. H., and Onore, M. (1964). Phytochemistry, 3,595-602. Hoogenhout, H. and Amesz, J. (1965). Arch. Mikrobiol., 50’10-24. Hutner, S. H. (1946).J. Bact., 52,213-220. Keane, M., Zahalsky, A. C., Hutner, S. H., and Lubart, K. J. (1963). In “Studies on Microalgae and Photosynthetic Bacteria” (Japanese Society of Plant Physiology Ed.), pp. 163-169. University of Tokyo Press, Tokyo. KiyoharaiT., Fujita, Y., Hatton, A., andwatanabe, A.~l96O).J.gen.appl.Mic~obiol. 6.176-182. Kondrat’eva, E. N. (I 965). “Photosynthetic Bacteria”. Israel Program for Scientific Translation, Jerusalem. Kornberg, H. L., and Lascelles, J. (1960). J. gm. Microbiol., 23, 511-517. Kratz, W. A., and Myers, J. (1955). A m . J . Bot., 42, 282-287. Krauss, M. P. (1966). Nature, Lond., 211, 301.

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Larsen, H. (1953). K. norske Vidensk. Selsk. Skr., 1-205. Lascelles, J. (1956). Biochem. J.3 62, 78-93. Lascelles, J. (1959). Biochem.J.,72,508-518. Lazaroff, N., and Vishniac, W. (1961).J.gen. Microbiol.,25, 365-374. McDaniel, H. R., Middlebrook, J. B., and Bowman, R. 0. (1962). Appl. Microbiol., 10,223. McLachan, J., and Gorham, P. R. (1962). Can.J. Mikrobiol., 8,l-11. Myers, J. (1960). In “Handbuck der Pflanzenphysiologie” (Ed. W. Ruhland), Vol. V, pp. 21 1-233. Pt. I. Springer-Verlag, Berlin. Myers, J., and Kratz, W. A. (1955).J. gen. Physiol., 39, 11-22. Ormerod, J. G., Ormerod, K. S., and Gest, H. (1961). Archs Biochem. Biophys., 94,449463. Pearce, J., andCarr, N. G. (1967).J.gen. Microbiol., 49, 301-313. Pfennig, N. (1961). Naturwiss enschajten, 48, 136. Pfennig, N. (1967). A. Rev.Microbiol., 21, 285-324. Pringsheim, E. G. (1914). Beitr. Biol. PfE.,12, 49-107. Pringsheim, E. G. (1949). “Pure Cultures of Algae”, University Press, Cambridge. Pringsheim, E. G. (1950). In “The Culturing of Algae” (Ed. J. Brunel, G. W. Prescott and L. H. Tiffany), pp. 19-26. Charles Kettering Foundation, Yellow Springs, Ohio. Provasoli, L., McLaughlin, J. J. A., and Droop, M. R. (1957). Arch. Microbiol., 25, 392-428. Scher, S., Scher, B., and Hutner, S. H. (1963). In “Marine Microbiology” (Ed. C. H. Oppenheimer), pp. 580-587. Charles C. Thomas, Springfield, Illinois. Schlegel, H. G., and Pfennig, N. (1961). Arch. Mikrobiol., 38,l-39. Scotten, H. L., and Stokes, J. L. (1962). Arch. Mikrobiol., 42, 353-368. Shaposhnikos, V. N., and Balitskaya, R. M. (1964). Mikrobiol., 33, 343-346. Shaposhnikov, V. N., Kondratiesa, E. N., and Fedorov, V. D. (1960). Nature, Lond., 187,167-168. Sistrom, W. R. (1962).J.gen. Microbiol., 28,599-605. Skerman, V. B. D. (1959). “Genera of Bacteria”, p. 160. Williams and Wilkins, Baltimore. Soriano, S., and Lewin, R. A. (1965). Antonie z’an Leeuwenhoek, 31, 66-80. Taha, M. S. (1963). Mikrobiologiya, 32, 498-503. Tchan, Y. T., and Gould, J. (1961). Nature, Lond., 192,1276. Trentini, W. C., and Starr, M. P. (1967).J. Bact., 93, 1699-1704. Tret’yakova, A. N. (1965). Mikrobiologiya, 34, 420-424. vanBaalen, C. (1962). Botanica Mar., 4,129-139. van Niel,C. B. (I 931). Arch. Mihobiol., 3, 1-1 12. van Niel, C. B. (1944). Bact. Rev.,8, 1-118. Wassink, E. C., and Manten, A. (1952). Antonie can Leeuwenhoek, 8, 155-163. Watanabe, A. (1959).J. gen. appl. Microbiol., 5 , 153-157. Webater, D. A., and Hackett, D. P. (1966). PZ. Physiol., Wash., 41, 599-605. Wieringa, K. T. (1968). Antonie wan Leeuzuenhoeh, 31, 54-56.

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CHAPTER I 1 1

Techniques for the Study of Anaerobic, Spore - forming Bacteria A. T.WILLIS Public Health Laboratory Service, Luton and Dunstable Hospital, Luton, Beds., England I. Introduction

.

11. Methods of Growing Anaerobes . A. Removal of oxygen by palladium catalysis . B. Removal of oxygen by alkaline solutions of pyrogallol C. Removal of oxygen by chromium and sulphuric acid D. Use of aerobic organisms to absorb oxygen . E. Other methods for removal of oxygen from air . F. Shake cultures and fluid cultures . G. Reducingagents .

III. Cultural and Other Techniques

A. B. C.

Tetanus Clostridial myonecrosis (gas gangrene) Clostridium welchii food poisoning

.

C.

. .

Lactose-egg-yolk-milk agar D. Nutrient gelatin . E. Glucose gelatin . F. Milkagar . G. Fermentation media . H. Cooked-meat medium . Acknowledgments References

.

.

81 81 86 87 87 88 88 88

. . .

.

. .

.

V. Tables of Classification of Common Clostridia of Clinical Interest

VI. Appendixon Media . A. Fresh-blood agar B. Egg-yolkagar

.

. . . . . . .

A. B. C.

IV. Bacteriological Diagnosis of Clostridial Infections in Man

80

. . .

.

Media . Selective agents . Inoculation of media . D. Stock cultures of anaerobes . E. Study of pure cultures . F. Isolation and purification of anaerobes

.

. . . . . . *

.

. . .

. . . .

. .

89 89 90 91 92 92 102 104 104 105 106 108 111 111 111 111 112 112 112 112 112 112 112

80

A.

T. WILLIS

I. INTRODUCTION During the last forty years a great variety of methods has been proposed for the study of anaerobic bacteria. These have been concerned, not only with methods for producing an anaerobic environment in which these organisms can be grown in artificial culture, but also with developing cultural techniques that facilitate rapid isolation and specific identification, especially of the human and animal pathogens. Since non-pathogenic anaerobes have not been studied so intensively, we are less well informed about these organisms; indeed, the authenticity of many of the described species is suspect. Thus, amongst the anaerobic cocci, Breed et al. (1957) admit that some species, possibly many of them, may be identical with one another, whilst amongst the 93 species of clostridia listed in the seventh edition of Bergey’s Manual (Breed et al., 1957) only a small proportion of the non-pathogens appear to fulfil the requirements of valid species. Isolating anaerobes whose characters do not accord with those of any species described in the literature is a common experience which, all too often, has led workers to publish descriptions of new species, many of which have been given a cloak of respectability by their inclusion in contemporary determinative classifications. “One of the first duties of any author who refers to a living organism by its generic and specific names is to make sure by all means at his command of the identity of the form in question. . . There is no practice so conductive to confusion or so detrimental to progress in the science of bacteriology as the willingness of workers to accept uncritically the names upon so called ‘authentic. stock cultures’ ” (Hall, 1926). It is regrettable that this observation is still pertinent today. In comparing the characters of an unknown anaerobe with those of described species, reference to Bergey’s Manual is most helpful. The method of classification used is basicaily a dichotomous key, which follows a logical development. Obviously related species are not separated from one another, and all species are fairly presented in so far as their descriptions allow. Though one may disagree with some points of detail, such as the division of the fusiform bacilli into four genera on minor morphological differences (Weiss and Rettger, 1937), and the separation of the proteolytic species of Clostridium botulinum from the non-proteolytic Bergey’s Manual presents a good account of anaerobic organisms, and It 1s recommended for reference. It is wise for bacteriologists who have had little experience with anaerobic organisms to study some earlier publications that deal with them, and with the principles of anaerobic culture. From the extensive literature on these subjects (see McCoy and McClung, 1939; McClung and McCoy, 1941),

111. TECHNIQUES FOR STUDYING ANAEROBIC BACTERIA

81

the following is a uscful selection: Robertson (1915-16); Henry (1916-17); McIntosh (1917) ; Committee upon Anacrobic Bacteria and Infections (1919); Hall (1922, 1929); Slanetz and Kettger (1933); Spray (1936); Dack(1940); Reed and Orr (1941, 1943); Hayward (1947); van Heyningen (1950); McVay and Sprunt (1952); Hare et al. (1952); McClung (1956); Parish and Cannon (1961) ; MacLennan (1962) ; and Hare (1967). The present text offers a short account of the best methods used for the culture and isolation of anaerobes of medical interest, especially the clostridia, and of the techniques and tests of the greatest value in their identification. More detailed accounts are to be found in the publications of Smith (1955), McClung and Lindberg (1957) and Willis (1962, 1964, 1965).

11. METHODS OF GROWING ANAEROBES When an oxygen-free or anaerobic atmosphere is required for obtaining surface growths of anaerobes, anaerobic jars provide the method of choice. Some of the methods for securing anaerobiosis in a jar are also applicable to single tube or plate cultures, but for laboratories that intend to undertake anaerobic bacteriology seriously, anaerobic jars are essential. A. Removal of oxygen by palladium catalysis Of the various anaerobic jars that have been devised, those that utilize the principle of Laidlaw (1915) are recommended. I n these platinum or palladium is used to catalyse the combination of oxygen with hydrogen to form water. Mclntosh and Fildes (1916) adapted Laidlaw’s method of producing an anaerobic atmosphere to a number of devices for anaerobic growth, and modified forms of their anaerobic jar today provide quite the best means of securing anaerobic conditions. There are three important types of McIntosh and Fildes anaerobic jars, all basically the same in construction, but differing in the form of the catalyst capsule.

1. Types of Mclntosh and Fildes anaerobicjars (a). In the original jar, and in some modified types of it, the capsulefor example, Wright’s capsule (Wright, 1943)-is composed of asbestos wool impregnated with finely divided palladium and enclosed in a finemesh brass or copper gauze envelope. The palladium is activated by heating in a Bunsen burner flame immediately before sealing the jar, after which hydrogen is run in. This method of operation is still widely used, since the capsule is easily made in the laboratory and can be used with any air-tight container of suitable size. 5

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Preparation of Wr2ght’s capsule. i. Preparation of palladinized asbestos. Dissolve 1 g of palladium chloride in 15 ml of distilled water acidified with a few drops of concentrated HCI. Thoroughly soak about 1.5 g of asbestos wool in this solution and dry it in the incubator. Tease out the impregnated wool and coat it with carbon in a smoky flame, and finally burn off the carbon deposit in a blow lamp. This reduces the palladium chloride and leaves a finely divided deposit of palladium on the asbestos wool fibres. Alternatively, the palladium chloride may be chemically reduced by immersion of the impregnated asbestos in boiling 5% sodium formate solution, T h e palladinized asbestos may be tested by directing a fine jet of hydrogen on to a sample piece; if the preparation is successful the catalyst will glow and ignite the hydrogen. T h e quantity of catalyst thus prepared is sufficient for 5-6 capsules.

ii. Preparation of the capsule. Fold a layer of the catalyst into a piece of fine mesh copper or brass gauze, to form an envelope, the open side of which is then secured with a strip of soft metal to ensure a good seal and to facilitate handling. After completion, capsules should be carefully examined for integrity of the envelope and for pieces of catalyst that may project outside them. Any projecting pieces must be removed, otherwise there is a great risk of the glowing catalyst igniting the hydrogen in the jar and causing an explosion. (b). I n electrically operated anaerobic jars the palladinized asbestos, similarly enclosed in a wire gauze capsule, is heated by a 12 V element, which is connected to two terminals in the lid of the jar (Fildes and McIntosh, 1921). Here, the catalyst is not heated until the jar has been sealed and filled with hydrogen. (c). Messrs Baird and Tatlock (London) have developed an anaerobic jar which utilizes a room-temperature catalyst (Heller, 1954). T h e catalyst, which is manufactured under patent as “D” catalyst by Engelhard, Surrey, consists of pellets of alumina coated with finely divided palladium. The Baird and Tatlock anaerobic jar (Fig. 1) using this type of capsule is very efficient and may be set up in a few minutes.

2. The hydrogen source T h e most convenient source of hydrogen is a cylinder of the compressed gas. Since it is important that the hydrogen should be supplied to the jar at low pressure, the hydrogen cylinder should be fitted with a reducing valve and the gas delivered at a pressure of not more than 0.5 Ib/inz. If a reducing valve is not available, a convenient low-pressure source is a foot-

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111. TECHNIQUES FOR STUDYING ANAEROBIC BACTERIA

ball bladder filled from the cylinder. T h e pressure-reducing mechanism isattached to a gas-washing bottle, which acts as a flow meter, and this in turn is connected to the anaerobic jar. SCREW

I

/

CLAMP

,H Y D R O G E N

VACUUM VALVE.

LID

--_

SACHET CONTAINING CATALYST’

,’

-

/

VALVE

0 -RING

- _ _ SIDE --RUBBER

ARM

TUBING

--INDICATOR CAPSULE

FIG. 1. B.T.L. anaerobic jar. (Reproduced by courtesy of Messrs Baird and Tatlock (London) Ltd.)

Brewer et al. (1955) criticized the use of hydrogen compressed in cylinders, They described a technique for producing hydrogen inside the closed jar, using moist sodium borohydride which slowly liberates hydrogen in the presence of a cobalt chloride catalyst. The advantages claimed for this method of hydrogen production are that it eliminates the need for manometers and cylinders and the danger of explosion from hydrogen under high pressure, After these and subsequent observations by Brewer and Allgeier (1966), a commercial anaerobic jar operating on this principle of hydrogen production within the jar was developed by Baltimore Biological Laboratories in America, under the trade name of the Gaspak System. This apparatus is not yet available in the United Kingdom, so that it has not been possible to conduct comparative trials. T h e Gaspak jar apparently utilizes a cold catalyst, the hydrogen being generated from a disposable hydrogen-carbon dioxide generator envelope which is activated by the ,additionto it of water. T h e cost of operating this system is likely to be high.

3. Setting up the anaerobicjar Cultures are placed in the jar and the lid secured. If only a few cultures are involved it is a good plan to fill up unused space with wooden blocks,

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A. T. WILLIS

thus minimizing the volume of gas. If a Wright’s capsule is used, this is heated by Bunsen burner and clipped into position on the underside of the lid before sealing the jar. T h e jar is then attached to a vacuum pump, which is fitted with a vacuum gauge, and air is evacuated until there is a “negative pressure’’ in the jar of about 30 cm of mercury. T h e tap on the jar is closed; the vacuum pump is disconnected ; and the low-pressure hydrogen source is connected to the jar. T h e tap is opened to admit the hydrogen, which is drawn in rapidly as the jar loses its partial vacuum. If a Wright’s capsule or the cold catalyst is used, there is then a short pause in the flow of gas until catalysis decreases the volume of gases in the jar and so draws in more hydrogen. After a steady flow of hydrogen into the jar has been established for 10-15 minutes, the tap is closed and the jar placed in the incubator, where catalysis will continue until all the oxygen in the jar has been used up. With electrically heated capsules, catalysis does not begin until the heating element is turned on. This is done after the jar has been filled with hydrogen. After the heater has been turned on, the gases in the jar expand slightly and force some of the water in the gas-washing bottle a short distance up the supply tube. Then, as catalysis commences, hydrogen is drawn steadily into the jar. The jar should remain connected to the hydrogen source for at least 30 minutes, the heater being turned on for 3 minutes on three occasions during that period. Since “D” catalyst is active at room temperature, heating is unnecessary. T h e jar is evacuated to a “negative pressure” of 30 cm of mercury, filled with hydrogen, and left attached to the hydrogen source for about 10-15 minutes. After the initial inrush of gas, a short pause followed by a constant flow of hydrogen shows that the catalyst is active. The jar is then placed in the incubator, where catalysis continues and rapidly produces anaerobic conditions. Since palladium catalysts are inactivated by moisture, the capsule must be quite dry before use. For Wright’s capsule, this is ensured by heating it in a Bunsen flame. Electrically heated capsules are conveniently dried in an incubator; the room-temperature catalyst is best dried in a hot-air oven and stored in a warm, dry place. Anaerobic jars of these three types may be bought, and spare capsules, gaskets, and so on, are also available from the manufacturers (Messrs Baird and Tatlock (London) Ltd., Chadwell Heath, Essex). Although jars with a Wright’s capsule have the advantage that they can be made in the laboratory, explosion risks with this type of jar are higher than with others. This risk is greatly reduced in electrically heated jars and appears to be minimal with the Baird and Tatlock anaerobic jar. For routine use I favour the Baird and Tatlock jar with the Engelhard “D” catalyst. It is relatively inexpensive, and the capsule remains active

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85

for a considerable time and is easily replaced. The jar is fitted with a replaceable rubber gasket which ensures an air-tight seal, and with an external indicator attachment which shows whether all free oxygen in the jar has been removed. These features, together with the ease and speed with which the jar can be set up, make it ideal for all anaerobic work.

4. Indicatorsfor anaerobiosis For all types of anaerobic jars it is important to include some system which serves as a check on the development of anaerobic conditions. A faulty capsule or a leaking jar may well prevent the development or maintenance of complete anaerobiosis. The most commonly used methods for indicating anaerobiosis are those Rammended by Fildes and McIntosh (1921) and Lucas (see Stokes, 1960). They depend on the reduction of methylene blue. I n an anaerobic environment it is reduced from its coloured oxidized form to a colourless reduced leuco-compound. (a) Fildes and Mclntosh indicator. Three stock solutions are prepared: (1) a solution of 6% glucose in distilled water; (2) 6 ml O f N/10 sodium hydroxide diluted to 100 ml with distilled water; and (3) 3 ml of 0.5% aqueous methylene blue diluted to 100 ml with distilled water (0.015%). Each time the indicator solution is required, equal parts of the three stock solutions are mixed together in a test-tube and the mixture boiled until the methylene blue is reduced. The tube of colourless indicator is immediately placed in the anaerobic jar, or attached to the external side tube, and the process of securing anaerobiosis is initiated. If anaerobic conditions are secured and maintained the indicator solution remains colourless, If, on the other hand, the blue colour returns to the solution, it has been oxidized because of a failure in the anaerobic equipment. The advantage of an external indicator over an internal one is obvious. (b) Lucas semi-solid indicator. The following formula is taken from Stokes (1960). To 5 ml of a 2% borax solution add 12 drops of 9% thioglycollic acid and 2 drops of phenol red. Then add 10 ml of methylene blue solution (one 19 mg reductase tablet (B.D.H.) to 200 ml of water) and 10 ml of melted sloppy agar. Boil the mixture in a water bath until it is colourless, transfer to ampoules and seal. When required, an ampoule is opened and is either placed within the anaerobic jar or attached with rubber tubing to the side-arm of the jar. In the presence of oxygen the indicator becomes blue to a depth of about 5 mm. With continued exposure the surface of the indicator begins to dry and it becomes ineffective. It can easily be rejuvenated by heating the top layer in a Bunsen flame, pouring it off, and leaving the

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A. T.WILLIS

moist part below exposed. This indicator is obtainable in sealed tubes ready for use (Messrs Baird and Tatlock (London) Ltd.). T h e need for indicators in anaerobic work underlines the importance of examining the anaerobic apparatus before use to ensure that it is in proper working order. Five minutes spent in testing the equipment may save many hours of futile incubation.

5. Anaerobiosis with added carbon dioxide Since the growth of some anaerobes is improved by the addition to the jar of 2-5y0 of carbon dioxide, and since no anaerobes are adversely affected by this concentration of the gas, it is a good plan to add carbon dioxide to the anaerobic jar as a routine. Like hydrogen, it is most conveniently available from a cylinder of the gas. T h e carbon dioxide is run into the jar after evacuation but before the hydrogen is run in. T h e jar is evacuated to a “negative pressure” of -35 cm of mercury; then, with the manometer still attached to the jar, carbon dioxide is run in until the “negative pressure” is reduced by approximately 1.5 cm of mercury. If atmospheric pressure is assumed to be 76 cm of mercury, the jar will now contain the equivalent of 2% of carbon dioxide.

6. Opening the anaerobic jar If the jar carries an external indicator, this is first checked to ensure that it is colourless. Such an observation should be made daily when incubation is continued for a number of days. One of the taps in the lid of the jar is then opened ;this may be accompanied by a hiss of inrushing air-evidence of an air-tight jar, but not necessarily indicative that complete anaerobiosi has been achieved. Indeed, the hiss is sometimes due to the escape of gases from the jar, especially if there has been bacterial fermentation during incubation. Similarly, the slight discoloration of the blood in fresh-blood agar plates, which is common under anaerobic conditions, is not indicative of complete anaerobiosis. After removal of cultures the jar is cleaned and dried, and stored in a warm dry place. Since all palladium catalysts are inactivated by moisture, the practice of storing jars set up and sealed is not recommended. Indeed, the life of the room-temperature catalyst may be appreciably shortened by this procedure.

B. Removal of oxygen by alkaline solutions of pyrogallol Alkaline solutions of pyrogailol have the property of absorbing large amounts of oxygen, and a variety of devices have been invented which make use of this reaction for producing anaerobic conditions. T h e method

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was first introduced by Buchner (1888). The great advantage of this method isthat it requires no special or expensiveequipment. The reagents are readily available and the method is suitable for use with any container that can be sealed. Plates for incubation are placed in the jar together with an anaerobic indicator. Pyrogallic acid is added to a solution of sodium hydroxide in a large test-tube, which is then placed inside the jar and the jar is quickly sealed. For each 100 ml of jar capacity, 1 g of pyrogallic acid and 10 ml of 2.5 N sodium hydroxide is used. Though anaerobiosis is fairly rapidly attained by this method, oxidation of the pyrogallate leads to the production of a small amount of carbon monoxide which is inhibitory towards some organisms. Moreover, excess of sodium hydroxide which is present in the pyrogalIate absorbs any carbon dioxide in the jar, and this can be a disadvantage with some organisms. I n order to overcome the problem of carbon dioxide absorption, Rockwell (1924) advocated the use of sodium bicarbonate instead of the hydroxide. This practice, however, does not lessen the amount of carbon monoxide released by the chemical reaction. Methods that utilize alkaline pyrogallol have been described by McLeod (1912-13), Wilson (1917), Spray (1930-31), Lockhart (1953), Mossel et al. (1959), Matthews and Karnauchow (1961) and Naylor (1963).

C. Removal of oxygen by chromium and sulphuric acid The chromium and sulphuric acid jar method for obtaining anaerobic conditionswas described by Rosenthal(l937). By mixing together powdered chromium and sulphuric acid, anaerobiosis is achieved by a dual chemical reaction. First, hydrogen is evolved as the metal is attacked by the acid; then the secondary reaction follows in which the chromous sulphate reduces any oxygen present. Marshall (1960) suggested an alternative method in which hydrogen and chromous ions are generated by using a mixture of sulphuric acid, pure granulated zinc and chromic sulphate. This method of producing an anaerobic atmosphere is highly effective, but it is rather time-consuming,laborious and “messy”, and sometimes fails inexplicably.

D. Use of aerobic organisms to absorb oxygen The use of aerobic organisms to absorb oxygen from the air was originally described by Fortner (1928). Organisms such as coliform bacilli and Suratiu murcescens are suitable for this purpose. Two plates of agar media of exactly the same diameter are selected. One of these is heavily inoculated with the aerobic organism and the other is streaked with the anaerobic culture. These two plates are fitted together at their rims, the junction made air-tight with plasticine, and the apparatus is placed in the incubator.

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Anaerobiosis takes some time to develop, SO that the method sometimes fails with strict anaerobes. When large numbers of anaerobic cultures must be handled, the method is clearly too laborious to be of practical value, Further reference to the use of the Fortner principle in the culture of anaerobic bacteria is made by Hobson (this Volume, p. 133).

E. Other methods for removal of oxygen from air Other methods that have been used for producing an anaerobic atmosphere include the phosphorus jar, the vegetable tissue jar, cultivation in z)ucuo, and cultivation in atmospheres of indifferent gases (McClung et a l , 1934-35; Bridges et ul., 1952; and see Willis, 1964). None of these methods is satisfactory for modern anaerobic work.

F. Shake cultures and fluid cultures Exclusion of oxygen from part of the medium is the simplest method of growing anaerobes, and is effected by growing the organisms within the culture medium, which may be either solid or fluid. I n solid media, growth is obtained as a shake culture in a tube or bottle, or as a pour-plate culture, This method of culture is favoured by some bacteriologists as an aid to purifying cultures of anaerobes. Fluid media in deep tubes are extensively used in anaerobic bacteriology since most anaerobes grow readily in this type of culture and manipulations with them are easily performed. Before use, media for shake culture and deep fluid culture must be heated in a boiling water bath or steamer to drive off any dissolved oxygen, and then cooled quickly before inoculation. There is a tendency for oxygen in the air to diffuse back into any exposed medium, and convection currents which are always present in fluid media help to distribute oxygen rapidly throughout the medium. This is largely overcome by incorporating a small amount of agar (0.05%) in fluid media, These sloppy-agar media are still fluid, but convection currents are reduced to a minimum.

G. Reducing agents Although the addition of reducing agents to fluid and shake cultures of many anaerobes is not necessary, it is essential for the satisfactory “aerobic” culture of the more fastidious anaerobes. Consequently, many bacteriologists add reducing substances routinely to all media used for anaerobic work. T h e chief value of reductants is their use in fluid cultures of anaerobes which are incubated in ordinary incubators without other anaerobic precautions being taken. T h e following are some of the reducing agents commonly used.

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1. Cooked meat particles Cooked-meat medium was introduced by Robertson (1915-16) and is probably the most widely used fluid medium for the culture of anaerobes. The cooked meat tissue contains reducing substances, particularly glutathione, which permit the growth of even strict anaerobes without the application of other anaerobic methods.

2. Glucose Glucose is usually incorporated in media in the proportion of 1%. It is a good reducing agent, is non-toxic, and it also serves as an additional nutritional agent for bacterial growth.

3. Thioglycollic acid ThioglycoIlic acid or its sodium salt may be used in media in the proportion of 0*01-0.2%, Sodium thioglycollate gradually becomes slightly toxic towards micro-organisms and should therefore be freshly prepared before use. Thioglycollate media were first introduced by Brewer (1940a, b).

4. Cysteine Cysteine is a suitable reducing agent if used in concentrations not exceeding 0.05%. Higher concentrations inhibit bacterial growth. Hirsch and Grinsted (1954) considered that cysteine was superior to thioglycollate, and they included this reductant in their reinforced clostridial medium. The chief disadvantage of cysteine is its relative insolubility.

5. Metallic iron Metallic iron may be used as a reducing agent in fluid media. Iron wire, filings, nails and steel wool are suitable for this purpose. Ordinary “tin tacks” or screws are convenient to use for they are quickly sterilized by flaming before being added to sterile stock media (Spray, 1936; Hayward and Miles, 1943).

111. CULTURAL AND OTHER TECHNIQUES A. Media The general methods of preparing media for the culture of anaerobes are the same as for other bacteriological media. The basis for most of the media used is ordinary meat-infusion-peptone broth, and clear nutrient agar, to which various enrichment substances may be added. For growth of some 6f the more demanding organisms, it may be necessary to use a richer base medium, such as one of the “digest” broths. The preparation

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A. T. WILLIS

and uses of the various media commonly used in anaerobic work are described in the Chapter by Lapage et al. (this Series, Vol 3A). Many of the media used in anaerobic bactericlogy are available commercially in Britain under the trade names of Bacto (Agents-Messrs Baird and Tatlock (London) Ltd., Chadwell Heath, Essex), and Oxoid (The Oxoid Division, 0 x 0 Ltd., London, S.E.1.).

B. Selective agents I n order to facilitate the separation of anaerobic species from mixtures with other organisms, various substances may be incorporated into fluid or solid media. Some of these selective agents are more generally useful than others, but none of them is ideal.

1. Gentian violet Gentian violet, in a concentration of 1 in 100,000 in broth media, was suggested by Hall (1919, 1920) as a means of eliminating aerobic sporeformers from anaerobic cultures. For a similar purpose, Spray (1936) recommended the use of crystal violet. Although gentian violet is effective up to a point, the growth of many anaerobes is inhibited by the dye, which, in addition, has little inhibitory effect on the growth of Gram-negative organisms. Slanetz and Rettger (1933) recommended the use of 1 in 10,000 gentian violet in plate cultures for the isolation of fusiform bacilli, and Omata and Disraely (1956) found that a mixture of crystal violet (1 in 100,000) and streptomycin sulphate (10 pg/ml) in plate cultures facilitated the isolation of oral fusiform organisms.

2. Sodium azide Johansson (1953) introduced the first selective egg-yolk medium for clostridia by the addition to it of 0.02% of sodium azide which inhibited the growth of spore-forming aerobes. Later, Wetzler et al. (1956a) used a selective blood agar medium containing sodium azide and chloral hydrate. These observations were foreshadowed by those of Lindberg et al. (1954) who incorporated sodium azide, chloral hydrate, sorbic acid and polymyxin B in their selective medium for clostridia. 3 . Sorbic acid Sorbic acid in broth and solid media was used by Emard and Vaughn (1952) and York and Vaughn (1954) for the isolation of clostridia. This agent was more selective in fluid media when polymyxin B was added (Wetzler et al., 1956b).

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4. Phenethyl ulcohol Dowell et al. (1964) and Moore et al. (1964) used 0.25% phenethyl alcohol as a selective agent for clostridia in both fluid and solid media. It inhibits the growth of Gram-negative facultative anaerobes, but not that of Grampositive species.

5. Antibiotics Antibiotics have been used extensively as selective agents for clostridia. Polymyxin B was first used by Hirsch and Grinsted (1954), and has since been employed by a number of other workers, commonly in combination with other inhibitory agents (Lindberg et al., 1954; Wetzler et al., 1956a and b; Angelotti et al., 1962; Southworth and Strong, 1964). Streptomycin has been incorporated in enrichment cultures and holding media (Willis, 1957; Smith, 1959), and sulphadiazine, in combination with polymyxin, was used by Angelotti et al. (1962) in the sulphite reduction test for the enumeration of C1. welchii in food. Neomycin sulphate was introduced by Lowbury and Lilly (1955) for the isolation of C1. welchii type A from mixtures with facultative anaerobes. Its use was later extended by Lilly (1958) to the isolation of other types of C1. welchii, and Willis and Hobbs (1959) used it for the isolation of most of the common clostridia. Used in solid media, at a concentration of 100250 pg/mI, it allows many clostridia to grow, but completely inhibits the growth of most aerobic species. C. Inoculation of media 1. Plate cultures Plate cultures are inoculated in the usual way. When dealing with very strict anaerobes it is desirable to use freshly prepared plates, since during storage the medium takes up oxygen from the atmosphere in sufficient amounts to prevent growth of these, even though complete anaerobiosis has apparently been secured in the jar. Alternatively, it may be more convenient to store a set of uninoculsted plates under anaerobic conditions, the whole jar being kept in the refrigerator. After plates have been inoculated, they should be placed under anaerobic conditions as quickly as possible. Similarly, surface cultures of such organisms must be subcultured to an anaerobic environment promptly after removal from the jar.

2. Half-antitoxinplutes For the preparation of half-antitoxin plates, the medium is first thoroughly dried in the incubator. Three or four drops of antitoxin are then pipetted

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on the plate and spread over half its surface. A convenient spreader consists of a pulled Pasteur pipette, the distal end of which has been bent in the form of a hockey stick. After antitoxin treatment, the plate is allowed to stand for a few minutes to allow the serum to be absorbed into the medium.

3 . Selective neomycin media If media containing neomycin are not available, satisfactory results are obtained by spreading a few drops of 1% aqueous neomycin sulphate solution over the surface of a neomycin-free plate before inoculation (Hobbs,

1960).

4. Fluid media Fluid media are inoculated after dissolved oxygen has been driven off by steaming or heating in a bath of boiling water. When inoculating with a Pasteur pipette, the inoculum should be pipetted gently into the depths of themedium, care being taken not to introduce any air bubbles. Very thorough flaming of the pipette before use is most important; the use of unsterile pipettes is one of the commonest causes of culture contamination. Growth develops most rapidly in fluid media which are inoculated while still warm, and are incubated in a water bath. Some strict anaerobes, such as C1. oedematiens type D, will not grow, however, unless cultures are incubated in the anaerobic jar.

D. Stock cultures of anaerobes Freeze-drying of anaerobic cultures is the most satisfactory method of preserving them. However, the preparation of freeze-dried material requires special and expensive apparatus, which is not available in many laboratories. Preservation in tubes of cooked-meat medium is a simple and quite reliable method. After incubation, the tubes are sealed in a flame, and stored at room temperature in the dark. Rejuvenation is effected by subculturing to fresh cooked-meat medium. Labelled strains of anaerobic bacteria are available from the National Collection of Type Cultures (Central Public Health Laboratory, Colindale Ave., London, N.W.9.), and from the National Collection of Industrial Bacteria (Torry Research Station, Aberdeen). I n addition to these, a large collection of organisms is held by the Wellcome Research Laboratories (Langley Court, Beckenham, Kent).

E. Study of pure cultures Before applying the various methods of identifying an anaerobic organism, it is most important to ensure that the culture in question is pure.

111. TECHNIQUES FOR

STUDYING ANAEROBIC BACTERIA

93

It must be free from both aerobic and anaerobic contaminants. When subcultures are made from a supposedly pure culture of an anaerobe, an aerobic control plate must always be inoculated, and this plate should remain sterile if aerobic contaminants are absent. Some aerotolerant organisms, however, notably CI. histolyticum and C1. tertiurn, produce small surface colonies under aerobic conditions; similarly, some organisms which grow only anaerobically on primary isolation may become oxygen-tolerant after two or three subcultures. Organisms of these types must be borne in mind when the aerobic control plate is examined. The confusion which may be caused by studying impure cultures is thoroughly understood. At the same time, however, it must be remembered that individual organisms in pure culture may appear in two or more phases from time to time, showing differences in morphology and in cultural and biochemical characters. In such cases, care must be taken not to confuse contaminationwith phase variation and vice versa. Pure cultures of anaerobes are far from easy to maintain; it is always as well to be prepared for the appearance of contaminants, and sudden changes and variations in the properties of a culture should lead one to suspect their presence. Only if it is certain that the organism is in pure culture, may the process of recording its characteristicsbegin. 1. Morphology and staining The shape, size, Gram-reaction and presence or absence of spores determine to which genus a particular anaerobe belongs. Table I summarizes the main morphological characters of the anaerobic genera.

TABLE I Some characteristics of the main anaerobic genera Organism Clostridium Fusiformis coccus

Shape

Spores

Gram-reaction

Bacillary Bacillary Coccal

+

+

-

-

+ or -

Arrangement None typical Filaments common Chainsandmasses

(a) Gram-reaction. The conventional methods of staining by Gram’s method are quite satisfactory for anaerobes, but it should be noted that

Gram-positive anaerobes not infrequently stain Gram-negative, especially in old cultures. This is especially true of strict anaerobes such as CI.tetani and Cl. oedematiens, and is probably due to the toxic effects of oxygen on these organisms. In these circumstances, however, the presence of spores will exclude anaerobes other than the clostridia.

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A . T. WILLIS

(b) Spores. I n Gram-stained preparations a spore shows as a circular or oval unstained area in the body of the bacillus. T h e size, shape and position of the spore should be noted, for sporulating cultures of some clostridia, for example C l . tetani, present a characteristic microscopic appearance on which a presumptive diagnosis may be made. Although some taxonomic systems attach importance to the size and position of the spore, these features are usually far too variable to be of any diagnostic use. A complete absence of spores in cultures should be noted, for some clostridia, notably CE. welchii, fail to sporulate readily. (c) Capsule staining. Capsule staining does not find much application, since only two of the common anaerobes are capsulated, namely C1. welchii and CZ. butyricum. Capsule formation may be demonstrated in pathological material and in cooked-meat broth cultures to which has been added a little blood or serum, I n the direct examination of pathological material from some CZ. welchii infections, capsule staining is of considerable importance (see p. 105). Muir’s capsule stain is prepared as follows-

Solution i. Carbol fuchsin (Ziehl-Neelsen). Solution ii. Saturated solution of mercuric chloride 20% solution of tannic acid Saturated solution of potassium alum Solution iii. Methylene blue

2 vols. 2 vols. 5 vols.

Technique. A thin film is fixed by heat and flooded with strong carbol fuchsin for 1 min., the preparation being gently heated. T h e film is then washed rapidly with alcohol and then with water. It is then treated with Solution ii (Muir’s mordant) for 30 sec and washed with water. Alcohol is then applied for 30 sec, the preparation becoming pale red in appearance. T h e methylene blue counterstain is applied for 1 min and the film is finally washed and dried. Organisms appear red with blue capsules.

(d) Immuno-fiuorescent staining. Development of the fluorescent labelled antibody technique by Coons et al. (1941) has made available a new approach to identifying clostridia. Of importance in this respect is the work of Batty and Walker (1963a, b, 1965), who have shown that C l . septicum and C1. chauvoei can be identified and differentiated from one another with certainty by specific staining with fluorescent-labelled vegetative “0”antisera. By this method the organisms may be identified, not only in smears of pure and mixed cultures, but also in smears of pathological material, and in tissue sections. I t seems likely that this technique will prove equally

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valuable in differentiating proteolytic members of CI. botulinum from CI. sporogenes (Batty and Walker, 1964, 1965; Boothroyd and Georgala, 1964). Fluorescent labelled antisera also act as specific stains for Cl, oedematiens and CI. tetani (Batty and Walker, 1963b, 1965). The successful application of the immunofluorescent technique depends on the sharing by all strains of the particular species of at least one common antigen that is not possessed by any other species (see Walker, Batty and Thomson, this Series, Vol. 5).

2, Motility Absence of motility is of no significance in identifying any organism; for there are non-motile variants of motile species, and it is common among the anaerobes for motility to be lost due to death of the organism or to interference with its flagellar mechanism. Hence the routine examination of anaerobes for motility is not an essential procedure. Since most anaerobes are adversely affected by the presence of free oxygen the best method for observation of motility is to use a capillary tube filled with an actively growing fluid culture (6-18 h) and sealed at both ends. Such a capillary tube is easily prepared from a Pasteur pipette. The preparation is examined under the high-power objective with reduced illumination. An alternative to this direct method of observing motility is the use of stab cultures in semi-solid agar (0.5%) (Tittsler and Sandholzer, 1936).

3. Colonial appearance Any medium which supports good surface growth of anaerobes is satisfactory for the study of colonial morphology. Media used for this purpose should not contain more than 1*0-1.5% New Zealand agar because higher concentrations appreciably alter some of the colonial characters. Freshhorse-blood agar and heated-blood agar are suitable media. For the full development of characteristic colonies some organisms require longer periods of incubation than others. Most anaerobic organisms produce semi-translucent or clear surface growths, important exceptions being the actinomycetes and some members of the Fusiformis group. The colonial morphology of the anaerobes shows considerable variation from one species to another. Thus, Cl. welchii is one of the few common clostridia which produces a circular discrete colony. CI.tetani, on the other hand, is characterized by its swarming growth. The size and shape of colonies are not static features; with continued incubation, convex colonies may become umbonate, circular colonies may become irregular, clear or semi-translucent colonies may become opaque or pigmented. Colonial appearances may also vary on different media, being affected by such factors as the concentration of the agar and the amount of

96

A . T. WILLIS

moisture present on the surface of the medium. The beautifully illustrated paper by Batty and Walker (1966) demonstrates the variety of colonial appearances that are shown by different species of clostridia.

4. Cultural characteristics (a) Fresh-blood agar. All the clostridia that are pathogenic for man will grow satisfactorily on fresh-blood agar. Like some aerobic species, some anaerobes produce a- or /3-haemolysis on this medium. Though many anaerobes are haemolytic, different strains of a single species may vary considerably in this respect; for example, some strains of C l . welchii are non-haemolytic on horse-blood agar, others produce extensive zones of haemolysis and yet others produce only a limited partial haemolysis. Thus, haemolysis, considered alone, is not of much diagnostic value. It is worth noting that the haemolysins elaborated by individual anaerobic species are not equally active against the erythrocytes of all animals. For example, the haemolysis produced by Cl. welchii on horse-blood agar is due to its @-toxin, which is relatively inactive against the red cells of the mouse, whereas Cl. welchii a-toxin haemolyses mouse red cells but is relatively inactive against those of the horse. Use has been made of such differences by Brooks et al. (1957) for determining the types of haemolysins elaborated in culture by different strains of C l . welchii. They grew their strains on the surface of anaerobically incubated horse and ox-blood agar plates, some containing a-antitoxin, some a- and &antitoxin and some without antitoxin. Identification of the haemolysins was established, not only by inhibition of haemolysins on antitoxin plates, but also by the characteristic patterns of haemolysis which they produced. On ox-blood agar, for example, the colonies of strains producing both a- and 0-toxins showed “target” haemolysis-central complete lysis (&toxin), with an outer ring of partial haemolysis (a-toxin) commonly surrounded by a zone of darkening. T h e same changes were also sometimes seen on horse-blood agar. In the absence of 0-toxin, colonies of strains forming a- and &toxins showed a similar “target” appearance on ox-blood but not on horse-blood, against the red cells of which blysin is inactive. Fusiformis melaninogenicum is an unusual organism in that its colonies blacken with prolonged incubation. This pigmentation, which takes place only on blood-containing media, begins in the centre of the colony and gradually extends to its periphery, through shades of brown, to black. T h e responsible pigment is hacmatin (Schwabacher and Lucas, 1947). (b) Human-serum and egg-yolk agar media. Seiffert (1939) and Nagler (1939) showed independently that when Cl. welchii is grown in media containing

97

111. TECHNIQUES FOR STUDYING ANAEROBIC BACTERIA

human serum a dense opalescence is produced ; this reaction-the Nagler reaction-is given by culture-filtrates of Cl. welchii as well as by the growing organisms, and is inhibited by C1. welchii antitoxin (Nagler, 1939). Later, Macfarlane et al. (1941) reported that similar but stronger opalescence was produced in egg-yolk emulsions, and showed that the reactions are due to the a-toxin of Cl. welchii. Hayward (1941, 1943), using human-serum agar plates, showed that C1. welchii, Cl. oedematiens and members of the C1. b~ermentans-sordelliigroup produced a dense diffuse opalescence on this medium; the opacity due to Cl. welchii and Cl. oedematiens was inhibited by specific antitoxic sera, whereas that due to Cl. bifermentans was inhibited by Cl. welchii antitoxin. TABLE I1 Reactions of various clostridia on egg-yolk media (Modified from Willis and Hobbs, 1958) Opalescence -----7

Organism __

Cl. welchii A-E Cl.bifermentans-sordellii CI.botulinum A-F Cl. sporogenes Cl. oedematiens Type A Type €3 Type c Type D

Produced

___ -

2

+ + + + + + +

Inhibition by Cl. welchii alpha-antitoxin

Pearly layer

~

+ +

-

-

+ + -

-

-

-

-

i-

-

Subsequent work in this field (Nagler, 1944, 1945; McClung et al., 1945; McClung and Toabe, 1947; Miles and Miles, 1947; Oakley et al., 1947) showed that various clostridia are able to produce opalescence in egg-yolk media (Table 11), and that this is sometimes due to specific lecithinases C secreted by the organisms. On egg-yolk media, some clostridia produce, in addition to an opalescence, a pearly layer which covers the colonies and may extend beyond their edge on to the surface of the medium (Nagler, 1945). This is due to lipolysis (Willis, 1960a, b, 1962). Egg-yolk media are thus of considerable value in identifying opalescence-producing clostridia and, used as half-antitoxin plates, in distinguishing between them. JVillis and Hobbs (1957, 1958, 1959) developed a lactose-egg-yolk agar medium. It is superior to other egg-yolk media in that it not only provides

98

A. T. WILLIS

more information about the organisms growing on it, but also enables a presumptive diagnosis of some anaerobes to be made from the single plate culture. T h e egg-yolk content of the medium enables appropriate organisms to produce their pearly layer or opalescence effects, or both. T h e presence of lactose in the medium provides a means of identifying lactose-fermenting organisms, which is of particular value in distinguishing between cultures of Cl. welchii and Cl. bifermentans-sordellii. Lactose-fermenting anaerobes produce diffuse red haloes in the medium around areas of growth and the colonies remain uncoloured until they are exposed to the air. T h e presence of milk in the medium enables proteolytic activity to be determined since it is attacked by proteolytic species and the result is the development of zones of clearing around areas of growth. Lactose-egg-yolk-milk agar, used as a half-antitoxin plate with a mixture of Cl. welchii type A and Cl. oedematiens type A antisera as the antitoxin, is differentially diagnostic for Cl. welchii, Cl. bifermentans-sordellii, Cl. oedematiens type A, Cl. histolyticum and non-proteolytic strains of Cl, botulinum. T h e reactions given on this medium by the commoner anaerobic species are shown in Tables I11 and IV. Anaerobes that show lecithinase C activity on egg-yolk media are Cl. welchii, Cl. bifermentans, Cl. sordellii and Cl. oedematiens types A, B and D ; anaerobes that show lipolytic activity are el. sporogenes, all types of Cl. botulinum and Cl. oedematiens type A. T h e mechanisms of the egg-yolk reactions of clostridia were studied and reviewed by Willis and Gowland (1962) and Willis (1960a, b ; 1962; 1964). (c) Gelatin and glucose-gelatin. Depending on whether or not an organism liquefies gelatin, it may be said that the organism is at least slightly proteolytic or not proteolytic at all. All strongly proteolytic anaerobes digest gelatin, but there are many gelatinase-producing anaerobes which do not break down more complex proteins. Gelatinase activity is demonstrated by growing the organism in nutrient gelatin or glucose-gelatin media. After incubation it is necessary to cool the culture at 4°C for half an hour, since gelatin is fluid at incubator temperature (37°C). (d) Inspissated serum. Loeffler’s medium is used for the determination of proteolytic properties of anaerobes. Liquefaction of inspissated serum is produced only by strongly proteolytic species, all of which are clostridia. When a clostridium is designated “proteolytic” it is generally taken to mean that it will attack complex proteins such as heated serum. In Loeffler slope cultures of proteolytic anaerobes there is first a surface erosion of the medium, followed later by its liquefaction or disintegration, accompanied by a putrefactive odour. (e) Milk agar. Milk agar was devised by Reed and Orr (1941) and serves the same purpose as inspissated serum. Plate cultures of proteolytic clost-

111. TECHNIQUES FOR STUDYING ANAEROBIC BACTERIA

99

ridia produce zones of clearing beneath and around areas of growth, due to digestion of the casein. (f) Fermentation media. Most anaerobes grow well in the ordinary peptonewater-sugar media used in the identification of enteric bacilli. The most TABLE I11

Reactions of clostridia on half-antitoxint lactose-egg-yolk-milk agar (modified from Willis and Hobbs, 1959) Lactose-egg-yolk-milk agar

,-*-( Organism

- __

Cl. welchii A-E CI. bifermentans Cl.sordellii Cl botulinum A )I B )Y

9)

c

E F

Opalescence

Produced Inhibited

____

Pearly Lactose layer fermentation Proteolysis ______I_

+ + +-

-

Ct. sporogenes Cl.oedematiens A B )) ))

C

,I

D

C1. histolyticum C1. septicum Cl. chauvoei Cl. tetani Cl. cochlearium Cl. capitovale CI.sphenoides Cl. tertium C1. tetanomorplium Cl. butysicum Cl. fallax Cl. innominatum

-

+... ...

...

... ... 1 . .

... ...

... ...

... ...

...

tMixture of C1. zuelchii type A and CI.oedematiens type A antitoxic sera,

important fermentable substances are glucose, maltose, lactose and sucrose, in that order. Strongly saccharolytic species, such as CZ.welchii ferment all these sugars; weakly saccharolytic organisms may ferment glucose or glucose and maltose only. Among the clostridia, organisms which attack

TABLE IV A scheme for the identification of clostridia using lactose-egg-ydk-mi& half-antitoxin? plates

w 0 0

Opalescence on egg-yolk agar

, I I

Produced

Not produced

I

Opalescenci inhibited by antiserumt

I La'ctose fermented

I

Proteolytic

Non-proteolytic

C1. histolyticum

CI. tetani CI. septicum Cl. tertium Cl. oedematiens

Opaiesdence not inhibited by antiserum

I

Ladtose not fermented

i

Lactose not fermented

type

c

? -I

sr: m

Cl. oedematiens type A

Cl. bifermentans C1. swdellii

I

Pearly layer

CI.sporigenes Cl. botulinum types A-F tA

mixture of Ct. welchii t y p e A and CI. oedematiens t y p e A antisera.

I

No pearly layer

I

Cl. oedbnatiens types B and D

111. TECHNIQUES FOR STUDYING ANAEROBIC BACTERIA

101

glucose usually also ferment maltose ; but organisms which do not ferment glucose do not attack any of the sugars mentioned, and constitute the group of non-saccharolytic clostridia. Another general rule that is often of value in identifying clostridia is that lactose-fermenting anaerobes are rarely strongly proteolytic. If peptone water fails to support the growth of any particular organism, the medium may be appropriately enriched. Thus, for fusiform species, the addition of serum to the medium may be necessary; some clostridia may require for growth the addition of an enrichment substance such zs Fildes’ extract of red cells or liver extract.

(g) Indoleproduction. Indole is tested for in the usual way by adding Ehrlich’s reagent to a peptone water culture of the anaerobe. The test for indole is not of great value in dealing with the common anaerobes. Some fusiform organisms produce indole; amongst the clostridia, C1. bifermentans, C1. sordellii and Cl. tetani are the only important indole-positive species. (h) Robertson’s cooked-meat broth. Practically all anaerobes grow well in cooked-meat broth. The growth of saccharolytic species is often associated with the development of a pink colour in the meat particles, probably due to the low Eh attained, with consequent reduction of the haematin. Proteolytic species attack and digest the meat particles, with the formation of a dark red or grey “sludge” in the bottom of the tube, and an accompanying putrid odour. Gas is commoqly produced in cooked-meat broth by both saccharolytic and proteolytic anaerobes. Some clostridia produce a butyrous scum on the surface of the broth, which probably indicates lipolytic activity. 5. Animal inoculation and protection tests Ideally, especially for the pathogenic clostridia, final identification of a species is made by inoculating two animals, one of which has been protected with specific antitoxin. In practice, however, animal inoculation is often unnecessary, as most of the common anaerobes are easily recognized by other means. One important exception is CZ. botulinum, some strains of which are very similar in their cultural reactions to CZ.sporogenes. Final diagnosis can then be made only on the results of animal tests. Since the pathogenic clostridia are either histotoxic or neurotoxic, some amount of toxin is usually present in the material (commonly a fluid culture) used for the inoculation of laboratory animals. If Iarge amounts of toxin are present, disease may ensue very quickly and be rapidly fatal without the typical sequence of symptoms developing. All pathogenic anaerobes require a nidus of local necrosis at the site of their inoculation into the tissues in order to ensure multiplication and toxin production will occur. It is a common practice, therefore, to add 2.5% of

102

A. T. WILLIS

calcium chloride to the culture for inoculation, the salt acting as a necrotizing agent. Since calcium chloride is toxic to micro-organisms the salt is added to the culture immediately before inoculation. Another important use of laboratory animals in anaerobic work is the demonstration of pathogenicity or toxigenicity of a known organism. Cl. tetani, for example, can easily be identified by its cultural and morphological characteristics, but its toxigenicity can be established only by animal inoculation and protection tests. Absence of toxigenicity in such a case does not contradict a diagnosis of CI. tetani, since non-toxigenic strains of this organism are not uncommon. I n addition to these procedures, the typing of some anaerobes, such as C1. welchii and Cl. oedematiens, is partly dependent on the inoculation of animals with culture-filtrates. Here, the aim is to identify specific exotoxins, the presence of which can be recognized only by in vivo methods. Similarly, the use of animals is essential for the titration of many toxin preparations and for the standardization of toxoids and antitoxins. These techniques are beyond the scope of the present text (see Batty, this Series, Vol. 5).

6. Serology of anaer7bes T h e serology of the pathogenic anaerobes was reviewed by Smith (1955). Generally speaking, the various serological reactions, such as somatic and flagellar agglutination, precipitation and complement-fixation tests, do not play an important part in the recognition or subdivision of these organisms. However, application of immuno-fluorescent staining techniques to some species of clostridia by Batty and Walker (1963a, b ; 1965; 1966) provides a new approach to the identification of these organisms (Walker, Batty and Thornson, this Series, Vol. 5).

F. Isolation and purification of anaerobes T h e techniques for isolating and purifying many anaerobes from mixtures with other organisms differ in no remarkable way from those used for other bacterial species, careful surface plating on agar media and subsequent colony selection being of prime importance. However, a number of special points deserve brief mention here, before consideration is given to the bacteriological diagnosis of clostridial infections in man. 1. Surface culture Surface culture on ordinary fresh-blood agar is an excellent method for the separation of Cl. tetani from mixtures with other organisms. The material containing Cl. tetani is inoculated on to a segment of the plate, which is then incubated anaerobically. Characteristically, Ci. tetani grows

111. TECHNIQUES FOR STUDYING ANAEROBIC BACTERIA

103

as a thin swarming film of growth across the surface of the plate, leaving unwanted contaminants behind. Subculture is then made from the outermost limit of the spreading edge of growth. Swarming anaerobic growth on the surface of plate cultures, such as that produced by Cl. tetani and C1. sporogenes, may cause difficulties in isolating other anaerobic species, a problem that may be overcome by using concentrated agar media (Miles, 1943). Alternatively, when this difficultyis encountered in the isolation of species that are relatively tolerant to oxygen, such as C1. welchii, C1. tertium and C1. histolyticum, incubation of plates under partially anaerobic conditions will allow the growth of these, but not that of more strictly anaerobic, swarming species. Suitable partial anaerobic conditions are obtained by evacuating the anaerobic jar to -60 cm of mercury and replacing the air with hydrogen; the catalyst is removed from the jar before use. Plate media for surface culture may be rendered selective for anaerobes by the addition to them of one or a number of the selective agents mentioned earlier, Neomycin sulphate is probably the most commonly used of these, as it suppresses the growth of most facultative anaerobes without interfering with anaerobic growth. Indicator media, such as egg-yolk agar and its more complex modifications, and horse-blood agar, are particularly uscful for the recognition of certain anaerobic species, and their value is greatly increased when they are made selective by the addition of neomycin.

2. Enrichment culture The most commonly used medium for enrichment purposes is Robertson’s cooked-meat broth. As in other branches of bacteriology, the aim of enrichment is to encourage particular organisms in the inoculum to increase their absolute and relative numbers. Addition of neomycin sulphate to the medium greatly favours selective growth of anaerobic organisms. Enrichment culture is followed by plating on to plain or selective agar media before and after the enrichment culture has been differentially heated. CI. welchii is the fastest growing of all anaerobes, a property that enables it to become, very rapidly, the predominant organism in cooked-meat broth cultures of mixed species. Early plating of such a culture quickly accomplishes the separation of Cl. welchii from contaminating organisms. Since Cl. welchii does not usually sporulate in artificial culture, differential heating as an aid to its isolation is quite inappropriate once growth has commenced. In isolating heat-resistant food-poisoning strains of Cl, welchii from faeces, however, primary heating of the faeces in cooked-meat broth bdore incubation is usually very successful, owing to the abundant sporulation of these strains in the intestinal tract and the high thermostability of their spores.

104

A.

T. WILLIS

3 . Diflerential heating Differential heating of material is of considerable value in destroying non-spore-forming organisms in mixtures from which clostridia are to be isolated, its success depending on the presence of heat-resistant clostridial spores. Differential heating is performed in a fluid medium, usually cookedmeat broth, and may be applied at the time of inoculation, as for some foodpoisoning strains of C1. welchii in faeces, or after the culture has been incubated. T h e success of post-incubational pasteurization clearly depends on adequate sporulation by the clostridia present in the enrichment culture. This may be expected with most species except Cl. welchii. IV. BACTERIOLOGICAL DIAGNOSIS OF CLOSTRIDIAL INFECTIONS I N MAN A. Tetanus The diagnosis of tetanus is always made on clinical grounds before it is confirmed bacteriologically. Not infrequently bacteriological confirmation is impossible, as for example in idiopathic cases, in which the presumed lesion has been so slight as to be undetectable by the time clinical tetanus develops.

1. Microscopic examination of patlzological material Microscopic examination is made for the typical "drumstick" bacilli. Their presence in material from a wound, however, is not in itself indicative of C l . tetani infection, since pathogenic strains of the organism may be present in contaminated wounds from which patients recover uneventfully without prophylactic treatment. On the other hand, in cases of clinical tetanus, material from a wound which is the obvious focus of infection may contain such small numbers of tetanus bacilli that they escape detection.

2. Culture of material from the wound Culture of wound material is much more likely to succeed; for this purpose, the best specimen is the excised wound. The material is cut up into small pieces with sterile instruments, and these are inoculated into air-free cooked-meat broth, and on to fresh-blood agar, heated-blood agar and neomycin-lactose-egg-yolk-milk agar. Only a segment of each plate is inoculated, so that the spreading edge of a tetanus culture is more easily recognized should it develop. These cultures are incubated in the anaerobic jar. Similar plates are inoculated for aerobic incubation, except that neoniycin is omitted from the egg-yolk medium. All the plate cultures are examined after 18-24 h incubation, and are re-incubated if necessary. The

111. TECHNIQUES FOR STUDYING ANAEROBIC BACTERIA

105

enrichment meat-broth culture is subcultured to similar plate media cvcry 24 h for 4 days. These procedures not only provide the greatest chance of isolating Cl. tetuni, but also enable other organisms, both aerobic and anaerobic, to be isolated and identified. T h e value of concentrated agar media for the isolation of anaerobes other than Cl. tetani is noted elsewhere (p. 103).

B. Clostridial myonecrosis (gas gangrene) As with tetanus, the diagnosis of gas gangrene is made on clinical grounds. The presence of pathogenic clostridia, especially C l . welchii, in a wound is, in itself, of no diagnostic significance. 1. Microscopic examination of pathological material Tentative confirmation of the clinical diagnosis may be aided considerably by microscopic examination of pathological material. Thus, the presence of large numbers of regularly shaped Gram-positive bacilli without spores is strongly suggestive of a Cl. welchii infection, and Butler (1945) has drawn attention to the correlation between the severity of Cl. welchii infections on the one hand, and the degree of capsulation of the organism and the extent of leucocytic damage on the other.

2. Culture from pathological material The most important step in identifying the infecting organisms is by culture from pathological material, and this should include cultures for both aerobes and anaerobes. T h e object is not merely to identify the bacterial species in the wound, but to assess their relative numbers and significance. For this reason, primary direct plating of the pathological material is of the utmost importance. Preliminary enrichment without recourse to direct plating may give a completely false impression of the relative importance of the anaerobes ultimately isolated. The following method of examination of wounds for clostridia is recommended (Table V)(1) Direct films.

(2) Inoculate exudate or tissue on to: (i) aerobic and anaerobic freshhorse-blood agar; (ii) aerobic and anaerobic heated-blood agar ; (iii) anaerobic half-antitoxin lactose-egg-yolk-milk agar, with and without neomycin ; (iv) aerobic lactose-egg-yolk-milk agar. (3) Inoculate material into four tubes of cooked-meat broth for anaerobic incubation, and heat to 100°C for 5, 10, 15 and 20 min respectively. T h e primary object here is to isolate C1. oedematiens and

106

A.

T.\VII.l.IS

Cl. septicum, the tubes bcing subculturccl after 24 and 48 h incubation to aerobic and anaerobic fresh-horse-blood agar and to aerobic and anaerobic half-antitoxin lactose-egg-yolk-milk agar plates, neomycin being present in the latter medium for anaerobic incubation. (4) Inoculate material into cooked-meat broth for anaerobic incubation. This enrichment broth is subcultured to plates as in (2) above, at 1,2,4 and 7 days. It is useful to make these subcultures in duplicate, the inoculum in one case being heated to 80°C for 10 min before transfer. It is often advisable to use concentrated agar plates for anaerobic culture in order to prevent swarming growth. These plates may be inoculated in addition to, or instead of, those mentioned above. The procedures mentioned in (2, iii) and (3) are carried out to isolate particular species. T h e half-antitoxin plates, using a mixture of C1. welchii type A and C1. oedematiens type A antitoxic sera, indicate the presence of CI. welchii, Cl. bifermentans, Cl. sordellii, Cl. oedematiens, Cl. sporogenes and Cl. histolyticum. An incubation period of 48 h is required for full development of the cultural characters of the last 3 clostridia. If possible, a separate anaerobic jar is used for examining cultures at 24 h, since a brief exposure to air at this time may prevent further development of tiny, and as yet unrecognizable, colonies of Cl. oedematiens.

C. Clostridium welchii food poisoning Bacteriological diagnosis is made by isolating heat-resistant Cl. welchii of typical toxigenic pattern from the faeces of infected patients. Isolation is usually accomplished in inoculating a little faeces into cooked-meat broth, which is then steamed for 15-60 min. If a food poisoning strain of CZ. welchii is present, it resists the heat treatment and is obtained in pure, or almost pure, culture with subsequent incubation. The organism may also be recovered by direct plating from the contaminated food. Taylor and Coetzee (1966) drew attention to the wide range of heat resistance of different strains of Cl. welchii that are associated with food poisoning. They recommended that in order to ensure the isolation of non-heat resistant food poisoning strains from faeces, unheated samples of faeces should be cultured routinely in a selective cooked-meat broth containing neomycin. This is a highly commendable method, for it ensures successful isolation, not only of non-heat-resistant strains of the classical Hobbs food poisoning variety, but also of the thermolabile haemolytic strains that are an occasional cause of food poisoning in the United Kingdom.

TABLE

v

Scheme for examination of wounds for cloatridia

Pathological material

I I

I

I I I

Film

2 c1 X

and lactose-egg-yolkmilk agar

I

agar with and without neomycin

I

Cooked-meat broth Heat to'100"C for

Plate out for Cl. oedematiensand Cl. septicurn using #-antitoxin plates

Cooked-meat broth Plate out at

1

Early subcultures far rapidy growihg clostridia. Later ones for slow-growing organisms. Use

differential heating to reduce aerobic contaminants and fastgrowing anaerobes. Use concentrated agar as indicated.

yr

0

108

A. T. WILLIS

V. TABLES O F CLASSIFICATION O F COMMON CLOSTRIDIA OF CLINICAL INTEREST Most medical and veterinary bacteriologists are not concerned with details of classification and nomenclature, but are interested primarily in the broad division of micro-organisms into pathogens and non-pathogens, the further subdivision of the pathogens into easily recognizable species also being of first importance. This attitude is at variance with that of the bacterial taxonomist who, being unbiased by pathogenicity or other properties of organisms, attempts to fit them into the general order of living creatures. This general approach to classification, which, quite naturally, is the one adopted by Breed et al. (1957), results in wide dispersion of the clinically interesting clostridia amongst the non-pathogenic species. Thus, in the key to the species of the genus Clostridium in Bergey’s Manual the 13 pathogenic species are scattered among the other 80 non-pathogenic organisms; Cl. fallax appears in the 7th position in the key, Cl. septicum in the 17th position, Cl. novyi (oedematiem) in the Zlst, CZ. perfringens (welchii) in the 42nd, Cl. tetani in the 63rd and Cl. histolyticum in the 90th position. Of course, clinical bacteriologists cannot ignore the non-pathogenic clostridia, since some of these organisms may be present as contaminants in pathological material and they must be recognized. Further, some species such as Cl. sporogenes which are non-pathogenic may increase the severity and alter the clinical appearances of infections due to recognized pathogens such as Cl. welchii. Finally, some non-pathogens resemble pathogenic species, from which they must be distinguished. T h e obvious answer to this problem is to abridge the classification of the sporulating anaerobes, leaving out the species that are not of immediate interest. Systems of this type appear in most textbooks of bacteriology, and in more specialized works such as those of Spray (1936), T h e Medical Research Council (1943), Smith (1955), Marshall el al. (1956), Kaufman and Weaver (1960), and Willis (1964). I n the following key suggested for the pathogenic clostridia, separation of species is effected chiefly by differences in their proteolytic and saccharolytic abilities, and in their egg-yolk reactions. Although a key such as this forms a useful guide to the identification of unknown organisms, its very nature limits the amount of information by which strains are recognized, so that on its own it is not definitive. Consequently, organisms should be studied more thoroughly than the key suggests, a number of confirmatory tests being used where possible (Tables 111, IV and VI). And although the toxigenic clostridia can usually be recognized by their cultural properties, the final identification of unknown strains is accomplished by animal

111. TECHNIQUES FOR STUDYING ANAEROBIC BACTERIA

109

inoculation and protection tests using specific antitoxic sera. To facilitate identification of unknown species, some of the important characters of the pathogenic and some common allied clostridia are listed in Table VI.

Key to the toxigenic and some related clostridia Feeble aerobic growthCoagulated serum liquefied, Glucose not fermented Coagulated serum not liquefied, Glucose fermented No aerobicgrowthCoagulated serum liquefied Lactose not fermented Restricted opacity and pearly layer on egg-yolk agar Diffuse opacity, no pearly layer on egg-yolk agar Salicin fermented Salicin not fermented Coagulated serum not liquefied Lactose fermented Diffuse opacity, no pearly layer on egg-yolk agar No change on egg-yolk agar Sucrose fermented, salicin not fermented Sucrose not fermented, salicin fermented Lactose not fermented Restricted opacity and pearly layer on egg-yolk agar Diffuse opacity and restricted pearly layer on egg-yolk agar Diffuse opacity, no pearly layer on egg-yolk agar Indole produced Indole not produced

C1. histolyticum

Cl. carnis Cl. tertium

C1.botulinum (types A, B, and F)

Cl. bfermentans Cl. sordellii Cl. welchii (types A-E)

Cl. cltauvoei C1.septicum

Cl. botulinum (types c,D, 8c E) Cl. oedematiens (type4 C1. oedematiens (type D) Cl. oedemntiens (tYPe B)

TABLE VI

w w 0

Reactions of the commonly encountered clostridia ~~

Pathogenicity Milk,-*-( for agar Pearly laboratory Gelatin Glucose Maltose Lactose Sucrose liquefaction Indole digestion Opalescence layer animals Egg Y O k

Organism C1. welchii (A-E) C1.butyricum Cl. muhifermentam CI. tertium CI.fallax Cl. chauvoei

f -

CI. septicum Cl. sphenoides

CI.botulimm (A,B 8~F) Cl. botulimm (C-E) CI.oedematiens (A)

(B) (C) I* (D) CI.bifermentans CI. sordellii CI.sporogenes Cl. tetanomorphuni C1. capitavale CI. cochlearium C1. tetani Cl. histolytictim $9

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111. TECHNIQUES FOR STUDYING ANAEROBIC BACTERIA

No change on egg-yolk agar Glucose not fermented Glucose fermented Gelatin liquefied Gelatin not liqueficd

111

Cl. tetani

Cl. oedematiens (type C) Cl. dzjicile

VI, APPENDIX ON MEDIA A. Fresh-blood agar The fresh-blood agar medium (Medium No. 5 ) described by Lapage et al. (this Series, Vol., 3A) is suitable for the culture of most anaerobes. For certain species it may be necessary to use a modified formula, such as that of the Cl. chauvoei medium of Batty and Walker (Medium No. 18). It is always important to indicate the species of erythrocytes used in the medium.

B. Egg-yolk agar The nutrient agar base described by Lapage et al. (Medium No. 68b) is used, except that the agar content is increased to give a final concentration in the complete medium of 1.5% (w/v). After autoclaving, cool the medium to 50-55"C, and add 25 ml of egg-yolk emulsion to each 250 ml of molten base, Gently mix, and pour plates immediately.

Egg-yolk emulsion The egg-yolk emulsion is prepared as follows: Separate the yolks of fresh eggs by the usual culinary technique, and mix them thoroughly with an equal volume (about 20 ml per yolk) of sterile normal saline solution. This emulsion should be freshly prepared before use. C. Lactose-egpyolk-milk agar (Willis and Hobbs, 1959) The nutrient agar base described by Lapage et al. (Medium No. 68b) is used, except that the agar content is increased to give a final concentration in the complete medium of 1.5% (w/v). Immediately before sterilization, add to each 100 ml of nutrient agar base, 1.2 g lactose and 0.3 ml of a 1% solution of neutral red. After the medium has been autoclaved, cool it to 5O0-55"C, and add to each 100 ml of molten base, 15 ml of sterile cow's milk and 4 ml of egg-yolk emulsion (see above). Gently mix, and pour plates immediately.

Sterilecow's milk. The sterile cow's milk is prepared by autoclaving the milk in convenient volumes after separation and removal of the cream by centrifugation.

112

A. T. WILLIS

D. Nutrient gelatin T o the nutrient broth described by Lapage et al. (Medium No. 71) add 12% of gelatin. Tube off, and sterilize by steaming for 20 min on each of three successive days. E. Glucose gelatin This consists of nutrient gelatin to which 1% of glucose is added before sterilization. F. Milk agar (Reed and Orr, 1941) T h e nutrient agar base described by Lapage et al. (Medium No. 68b) is used, except that the agar content is increased to give a final concentration in the complete medium of 1.5%. After sterilizing, cool the base to 50°-55"C and add sterile skimmed milk (see above) to a concentration of 25%. Mix, and pour plates immediately. G. Fermentation media These are prepared as described in most standard texts. T h e inclusion of a Durham's tube for the detection of gas is optional. A few drops of indicator, for example, O.lyo phenol red solution are added to the cultures aft.. incubation. This is necessary, since most indicators are reduced under anaerobic conditions to colourless leuco-forms.

H. Cooked-meat medium The cooked-meat medium described by Lapage et al. (Medium No. 24) is suitable. For routine purposes, the medium should be distributed in test tubes, not in screw-capped bottles. ACKNOWLEDGMENTS

I am greatly indebted to Baird and Tatlock (London) Ltd. for permission to reproduce Fig. 1, to Butterworth and Co. (Publishers) Ltd for permission to reproduce much copyright material, to Dr. J. W. Howie for much helpful criticism and advice, and to my wife for her secretarial assistance. REFERENCES Angelotti, R., Hall, H. E., Foter, M. J., and Lewis, K. H. (1962). Appl. Microbial., 10,193. Batty, I., and Walker, P. D. (1963a).J. Path. Bact., 85, 517. Batty, I., and Walker, P. D. (1963b). Bull. Off. int. epizoot., 59, 1499. Batty, I., and Walker, P. D. (1964). J. Path. Bact., 88, 327. Batty, I., and Walker, P. D. (1965).J. uppl. Bact., 28, 112.

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113

Batty, I., and Wiilker, 1’. D. (1966). In “Identification methods for microbiologists”. (Ed. 13. M. Gibhs and F. A. Skinner). Academic Press, London. Boothroyd, M., and Georgala, D. L. (1964). Nature, Lond., 202, 515. Breed, R. S., Murray, E. G. D., and Smith, N. R. (1957). “Bergey’s Manual of Determinitive Bacteriology, 7th ed. Baillihre, Tindall and Cox, London. Brewer, J. H. (1940a). J . Bact., 39, 10. Brewer, J. H. (1940b). J. Am. med. Ass., 115, 598. Brewer, J. H., and Allgeier, D. L. (1966). Appl. Microbiol., 14,985. Brewer, J. H., Heer, A. A., and McLaughlin, C. B. (1955). Appl. Microbiol., 3, 136. Bridges, A. E., Pepper, R. E., and Chandler, V. L. (1952).J. Bact., 64,137. Brooks, M. E., Sterne, M., and Warrack, G . H. (1957).J. Path. Bact., 74, 185. Buchner, H. (1888). Zentbl. Bakt. ParasitKde., 4, 149. Butler, H. M. (1945). Surgery Gynec. Obstet., 81, 475. Committee upon Anaerobic Bacteria and Infections (1919). Spec. Rep. Ser. med. Res. Coun., No. 39. H.M.S.O., London. Coons, A. H., Creech, H. J., and Jones, R. N. (1941). Proc. SOC. exp. Biol. Med., 47,200. Dack, G. M. (1940). Bact. Rev., 4, 227. Dowell, V. R., Hill, E. O., and Altemeier, W. A. (1964).J. Bact., 88, 1811. Emard, L. O., and Vaughn, R. H. (1952).J. Bact., 63,487. Fildes, P., and McIntosh, J. (1921). Br.3. exp. Path., 2,153. Fortner, J. (1928). Zentbl. Bakt. ParasitKde, Abt. I., 108, 1 5 5 . Hall, I. C. (1919).J. Am. med. Ass.,72,274. Hall, I. C. (1920).J. infect. Dis., 27, 576. Hall, I. C. (1922). J. infect. Dis., 30, 445. Hall, I. C. (1926). J . Bact., 11, 407. Hall, I. C. (1929). J. Bact., 17, 255. Hare, R. (1967). In “Recent advances in medical microbiology” (Ed. A. P. Waterson) p. 284. Churchill, London. Hare, R., Wildy, P., Billett, F. S., and Twort, D. N. (1952).J. Hyg., Camb., 50, 295. Hayward, N. J. (1941). Br. med.J., 1, 811, 916. Hayward, N. J. (1943). J. Path. Bact., 55, 285. Hayward, N. J. (1947). In “Recent advances in clinical pathology” (Ed. S. C. Dyke), p. 69. Churchill, London. Hayward, N. J., and Miles, A. A. (1943). Lancet, i, 645. Heller, C . L. (1954).J. appl. Bact., 17, 202. Henry, H. (1916-17). J . Path. Bact., 21, 344. Heyningen, W. E. van (1950). “Bacterial Toxins”. Blackwell, Oxford. Hirsch, A., and Grinsted, E. (1954). J . Dairy Res., 21, 101. Hobbs, B. C. (1960). I n “Recent advances in c h i c a l pathology”, Series I11 (Ed. S. C. Dyke) p. 91. Churchill, London. Johansson, K. R. (1953).J. Bact., 65,225. Kaufman, L., and Weaver, R. H. (1960). J . Bact., 79, 119. Laidlaw,P.P. (1915). Br. med.J., 1,497. Lilly, H. A. (1958).J. med. Lab. Technol., 15, 165. Lindberg, R. B., Mason, R. P., and Cutchins, E. (1954). Bact. Proc., 54, 53. Lockhart, W. R. (1953). Science, N . Y . , 118, 144. Lowbury, E. J. L., and Lilly, H. A. (1955).J. Path. Bnct., 70, 105. McClung, Id. S.(1956). A. Rev. Microhiof., 10, 173. 6

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McClung, I,. S., arid McCoy, I:. (1941). “‘l’he Anaerobic bacteria and their activities in nature and disease: A subject bibliography”, Suppl. 1. University of California Press, California. McClung, L. S., and Lindberg, R. B. (1957). In Society of American Bacteriologists, “Manual of Microbiological Methods”, McGraw-Hill, New York. McClung, L. S., andToabe, R. (1947).J. Bact., 53,139. McClung, L. S., McCoy, E., and Fred, E. B. (1934-35). Zentbl. Bakt ParasitKde., 91,225. McClung, L. S., Heidenreich, P., and Toabe, R. (1945).J. Bact., 50, 715. McCoy, E., and McClung, L. S. (1939). “The anaerobic bacteria and their activities in nature and disease: A subject bibliography”. 2 vols. University of California Press, California. Macfarlane, R. G., Oakley, C. L., and Anderson, C. G. (1941). J . Path. Bact., 52, 99. McIntosh, J. (1917). Spec. Rep. Ser. med. Res. Coun., No. 12. H.M.S.O., London. McIntosh, J., and Fildes, P. (1916). Lancet, i, 768. MacLennan, J. D. (1962). Bact. Rev., 26, 177. McLeod, J. W. (1912-13)J. Path. Bact., 17, 454. McVay, L. V., and Sprunt, D. H. (1952). Ann. intern. Med., 36, 56. Marshall, J. D., Wetzler, T. F., and Kawatomari, T. (1956). US.arm. Forces med. J . , 7,1445. Marshall, J. H. (1960). J . gen. Microbiol., 22, 645. Matthews, A. D., and Karnauchow, P. N. (1961). Can. med. Ass.J., 84, 793. Medical Research Council (1943). M.R.C. War Memor. No. 2. 2nd ed. H.M.S.O., London. Miles, A. A. (1943). Army Path. Lab. Serv., Current Notes, No. 9, 3 . Miles, E. M., and Miles, A. A. (1947).J.gen. Microbiol., 1, 385. Moore, M. L., Engwall, C., and Moskal, P. A. (1964). Am.J. med. Technol.,30, 385. Mossel, D. A. A,, Goldstein Brouwers, G. W. M. v., and de Bruin, A. S. (1959). J . Path. Bact., 78, 290. Nagler, F. P. 0. (1939). BY.J . exp. Path., 20, 473. Nagler, F. P. 0. (1944). Nature, Lond., 153, 496. Nagler, F. P. 0. (1945). Aust. J . exp. Biol. med. Sci., 23, 59. Naylor, P. G. D. (1963). J. appl. Bact., 26, 219. Oakley, C. L., Warrack, G. H., and Clarke, P. H. (1947).J. gen. Microbiol., 1, 91. Omata, R. R., and Disraely, M. N. (1956).J. Bact., 72, 677. Parish, H. J., and Cannon, D. A. (1961). “Antisera, toxoids, vaccines and tuberculins in prophylaxis and treatment”. 5th ed. Livingstone, Edinburgh. Reed, G. B., and Orr, J. H. (1941). War Med., Chicago, 1,493. Reed, G. B., and Orr, J. H. (1943). Am.J. med. Sci., 206, 379. Robertson, M. (1915-16). J. Path Bact., 20, 327. Rockwell, G. E. (1924).J. infect. Dis., 35, 581. Rosenthal, L. (1937).J. Bact., 34, 317. Schwabacher, H., and Lucas, D. R. (1947). J. gen. Microbiol., 1, 109. Seiffert, G. (1939). 2. ImmunForsch. exp. Ther., 96, 515. Slanetz, L. W., and Rettger, L. F. (1933). J . Bact., 26, 599. Smith, H. W. (1959).J. Path. Bact., 77, 79. Smith, L. DS. (1955). “Introduction to the pathogenic anaerobes”. University of Chicago Press, Chicago. Spray, R. S. (1930-31).J. Lab. din. Med., 16, 203.

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Spray, R. S. (1936). J. Bact., 32, 135. Southworth, J . M. L., and Strong, D. H. (1964). J. Milk Fd Technol.,27, 205. Stokes, E. J. (1960). “Clinical bacteriology”, 2nd ed. Arnold, London. Taylor, C. E. D., and Coetzee, E. F. C. (1966). Mon. Bull. Minist. Hlth, 25, 142. Tittsler, R. P., and Sandholzer, L. A. (1936).J. Bact., 31, 575. Weiss, J. E., and Rettger, L. F. (1937).J. Bact., 33,423. Wetzler, T. F., Marshall, J. D., and Cardella, M. A. (1956a). Am. J. clin. Path., 26,345. Wetzler, T. F., Marshall, J. D., and Cardella, M. A. (1956b). Am. J. din. Path., 26,418. Willis, A. T. (1957).J. Path. Bact., 74, 113. Willis, A. T. (1960a). Nature, Lond., 185, 943. Willis, A. T. (1960b). J. Path. Bact., 80, 379. Willis, A. T. (1962). Lab. Pract., 11, 526. Willis, A. T. (1964). “Anaerobic Bacteriology in Clinical Medicine.” 2nd ed. Butterworth, London. Willis, A. T. (1965). Lab. Pract., 14, 690. Willis, A. T., and Gowland, G . (1962).J. Path. Bact., 83, 219. Willis, A. T., and Hobbs, G. (1957). Nature, Lond., 180,92. Willis, A. T., and Hobbs, G. (1958).J. Path. Bact., 75, 299. Willis, A. T., and Hobbs, G. (1959).J. Path. Bact., 77, 511. Wilson, W. J . (1917). Lancet, i, 724. Wright, B. M. (1943). Army Path. Lab. Serv., Current Notes No. 9, 3 . York, G. K., and Vaughn, R. H. (1954).J. Bact., 68, 739.

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CHAPTER IV

A Roll Tube Method for Cultivation of Strict Anaerobes R. E. HUNGATE Department of Bacteriology, University of Calqornia,Davis, California, U.S.A. I. Theoretical Considerations 11. Procedures . A. Avoidance of oxygen during media preparation and storage R. The nature of the culture medium . C. Sterilization D. Obtaining pure cultures . E. Storage of cultures . F. Applications of anaerobic methods to various media . G. Other applications .

.

References

.

.

117

. . . . . . . . .

120 120 126 127

128 131 132 132 132

I. THEORETICAL CONSIDERATIONS Categories such as micro-aerophilic, moderately anaerobic, strictly anaerobic, and extremely anaerobic lack precision in defining the degree of anaerobiosis. Oxidation-reduction potential can bg: a more precise index of anaerobiosis and can provide a long and continuous scale within which the range of oxygen relationships between the most aerobic and most anaerobic conditions can be compared. Oxygen exerts its effect through chemical reaction with constituents of the cells or of the medium. Following this reaction these constituents are more oxidized and the oxygen atom is more reduced. The degree of oxidation or reduction of a chemical system can be defined by the redox potential. Aerobes contain many systems at low redox potentials (i.e., the systems are anaerobic), but these are not subJect to irreversible injury by a high potential and in the active cell are maintained at low potentials by continuous interaction with reducing systems. Some anaerobes are not killed by high potentials, but cannot grow until the medium is more reduced ; others are killed, certain essential systems being irreversibly destroyed. In nature, oxygen is the almost universal cause of high redox potential. The question has been raised, does the oxygen or the potential inhibit

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the anaerobes? Oxygen cannot exist apart from its potential, though similar potentials can be induced by chemicals other than oxygen. These other agents with high potentials will also affect reduced systems within thecell if they can react with them, either spontaneously or aided by enzymes. An oxidant not reacting with metabolic systems within the cell may affect an electrode inserted to measure the potential, but such a measurement provides little indication of the potential of cell systems not reacting with the electrode T h e present discussion refers to systems which are free to react with each other. Oxygen oxidizes those biochemical systems in the cell which have an oxidation potential less than that of the oxygen. T h e redox potential of oxygen is a function of the concentration of oxygen and its reduced form according to the equationconc. 0 2 Eoxygen = Eo of oxygen + RT In nF conc. reduced form of02 Since n = 4 in the reaction 02-+2 0 2 - , for each tenfold decrease in the concentration of oxygen there is a decrease of 0.015 volts in the oxygen potential, provided the concentration of the reduced product (assumed to be water) remains constant. T h e potential of the reaction 0 2 4 2 0 2 - i s +0.81 V at pH 7.0 and at an oxygen concentration in equilibrium with one atmosphere pressure of 0 2 , or 0.80 V at the concentration in air. Some anaerobes such as the methane-forming bacteria cannot initiate growth at potentials greater than -0.33 V (Smith and Hungate, 1958). T h e oxygen concentration at - 0.33 V would be 10-(0'80+0~33)~(0'015) = 10-75 of the concentration of oxygen in the atmosphere. T h e amount of oxygen in one litre of water at 30°C in equilibrium,with air at one atmosphere pressure is0.02608 x 1000 x 0.21 mmoles,litre

22.4 = 0.245 mmoles/litre = 2.45 x moles/litre = (2-45 x 10-4) x (6.06 x 1023) molecules/litre since 1 mole of oxygen contains 6.06 x 1 0 2 3 molecules. = 1.48 x 1019 molecuIes/litre of water in equilibrium with

air at 1 atm. pressure. At a potential of -0.33 V this becomes1.48 x 1019 x 10-75 (see above). = 1-48x 10-56 molecules/litre. This calculation strikingly illustrates: (a) that it is difficult to obtain low potentials for cultivation of strict anaerobes, (b) that the permissible concentration of oxygen in solution becomes a statistical function (10-55

IV. ROLL TUBE ANAEROBIC TECHNIQUE

119

molecules/litre), rather than a finite number of molecules, (c) that it is impossible to obtain low potentials simply by removing oxygen, and, the corollary, (d) that to obtain the needed low potential some reduced system ata lower potential must be added. T h e concentration of the added reducing system should be minimal, to avoid toxicity and to simulate the concentrations in most natural environments. Since a stable potential in a system is the expression of equilibrium or flux between all interacting oxidation-reduction systems, addition of an oxidizing agent will raise the potential, and additional reducing agent will be needed to lower it to the former level. But this leaves some of the originally reducing material in an oxidized form, and the greater the proportion of the oxidized form, the higher the potential in the system. T h e greater the concentration of oxidized materials in the system the greater the concentration of reducing agents needed to bring the system again to a fixed low potential. This is the reason why exposure to oxygen during preparation of media must be avoided, and the addition of reducing agent must be the terminal step in preparation, i.e., must occur just before inoculation. Petri plates are unsuited for work with obligate anaerobes though their many advantages have led to numerous adaptations for anaerobic work. These have been successfully used with many bacteria only moderately sensitive to oxygen, but they are entirely unsuited for work with extreme anaerobes. A roll tube method was developed in which agar medium was distributed as a thin layer over the internal surface of test tubes charged with an anaerobic atmosphere (Hungate, 1947) for the isolation of obligately anaerobic bacteria of the rumen. This technique has undergone numerous modifications and improvements (Hungate, 1966) since the original description (Hungate, 1950). Experience has shown a considerable superiority of these methods over most anaerobic methods in common use. It is the aim of this chapter to indicate some of the factors to be considered in developing stringently anaerobic techniques and to describe in detail the procedure and rationale of the roll tube method. Since air is the primary source of oxygen, air must be excluded from cultures of delicate anaerobes. T h e low solubility of oxygen in aqueous solutions makes it generally more common in any gas phase than in contiguous aqueous solutions. Large gas spaces are therefore preferably avoided in strictly anaerobic procedures. A high ratio of liquid to gas volume is desirable. In the roll tube method exposure of bacteria and culture medium to air is avoided by displacing the air in the culture vessel with an oxygen-free gas such as carbon dioxide, hydrogen, nitrogen, or mixtures of these gases. Carbon dioxide is the gas of choice because it is heavier than air, relatively

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cheap, and valuable in buffering. Any oxygen in the gas must be completely absorbed. Vessels are stoppered under conditions preventing access of air. T h e cultures require no special incubators and can be removed and examined with no anaerobic precautions if kept stoppered. If opened, anaerobiosis can be continuously maintained during necessary manipulations, and the culture again closed without exposure to oxygen.

11. PROCEDURES A. Avoidance of oxygen during media preparation and storage 1. Removal of 0 2 from gases A vertical column of heated copper (in the form of coarse copper filings) is satisfactory and economical for removing 0 2 from gases. Various methods have been developed by many workers. I n our experience it has been most satisfactory to pack the copper in a Pyrex glass column (Fig. 1) 25 mm inside diameter, narrowed at top and bottom to permit attachment of rubber tubing or ground glass connections. T h e column is heated electrically to about 350°C by a coil of nichrome wire wrapped around the column. T h e wattage needed will vary according to the extent of insulation. T h e gas stream enters at the bottom and leaves at the top of the column. T h e glass above the copper surface is not covered with asbestos. This permits visual inspection to determine whether the copper is reduced, i.e., with a bright copper colour rather than the dark colour of the CuO. Reduction is achieved by passing hydrogen gas through the column. T o avoid an explosion, the tube must be filled with carbon dioxide or nitrogen to displace any oxygen before the hydrogen is introduced. Transition from one gas to another without entrance of any oxygen is easily accomplished by having a shortened Pasteur pipette (Fig. 1) introduced in the line prior to the copper column. T h e carbon dioxide or nitrogen is adjusted to flow at a slow rate and the conduit is opened by disconnecting at A as the rubber tube at B is closed by constricting it between thumb and forefinger and kept closed while the tube is disconnected. A rubber tube carrying a slow flow of hydrogen terminates also in a Pasteur pipette. This pipette is inserted in the disconnected open end of the rubber tube at A and air displaced by holding it within the open end a moment before the pipette is inserted tightly at A and the tube again opened at B. After the copper is reduced, the same procedure is used to change back to carbon dioxide without entrance of air. T h e electric current in the nichrome coil is operated continuously. T h e expenditure of electricity and the heat evolved are modest if the insulation is sufficient. In wiring the column, a thin asbestos sheet is first wrapped around the

IV. ROLL TUBE ANAEROBIC TECHNIQUE

121

glass tube, extending from the base to the upper level of the copper. Plumbers’ powdered asbestos is moistened and packed to the desired thickness around the asbestos sheet and nichrome coil. Each end of the nichrome wire can be attached firmly between two iron washers held between the nut and head of a 1; in. by Q in. iron bolt threaded throughout its length. The head and connections are buried in the asbestos, with the other end

From cylinder

FIG.1. Heated copper column for freeing gases from traces of oxygen. See text for explanation.

projecting at right angles from the asbestos surface. Two nuts with intervening washers are used to attach the electrical power supply leads to the external ends of the bolts, and additional asbestos can be added to cover and insulate these connections. T h e powdered asbestos dries and becomes sufficientlyrigid to support the coil and leads. T h e glass column and asbestos at the base should be supported and the column also held near the top.

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R. E. HUNGATE

2. Removal of

0 2 from media and other solutions T h e easiest way to remove oxygen from a heat-stable solution is to boil it vigorously for about one minute. It can then be kept anaerobic by replacing the vapour with oxygen-free gas as the container cools. Agar to be included in the medium can be added prior to boiling.

(4

(b)

FIG.2. (a) Method of holding pipetting tube to permit adjustment of level. (b) Method of gassing the tube during the pipetting process.

T h e tube delivering the gas into the vessel should extend down near to the liquid (Fig. 2b) but it is not necessary to bubble gas through the liquid if the latter is already oxygen-free. I t is sufficient to exclude air. If the solution is heat labile it can be freed of oxygen by bubbling oxygen-free gas through it but the rate of removal is slow unless the gas bubbles are very abundant. Oxygen escapes by diffusion through the gas-liquid interface, making the rate of removal a function of the interface area. Thirty minutes to an hour are usually required to rid a solution of most of the dissolved oxygen, but even more extended bubbling is necessary to equal the effect of boiling. One of the best methods to ensure a low oxygen concentration in a medium is to allow sufficient growth of an aerobic micro-organism in the unsterilized stoppered tube to remove all the dissolved oxygen. T h e oxygen in the gas phase can also be absorbed if the tube is thoroughly shaken several times during a 30-minute incubation period prior to sterilization. This method is applicable only if the metabolic products or the bodies of the bacteria employed to remove the oxygen (which are killed later by autoclaving) do not interfere with the use of the culture. As mentioned earlier, the concentration of dissolved oxygen is small in comparison to that in the equilibrated gas phase and in many instances the reducing agent can effectively dispose of the amounts in solution. As a precaution, however, it is much better to store ingredients such as sugar

123

IV. ROLL TUBE ANAEROBIC TECHNIQUE

solutions, bicarbonate solutions, and other dissolved materials in closed containers from which air has been excluded. The reducing agent must always be kept under oxygen-free conditions.

3. Dispensing anaerobic media It is most convenient to prepare media in large batches, transferring to the desired culture containers before or after autoclaving (Figs. 2b, 3a). Transfer prior to sterilization is more desirable because heat accelerates combination of 0 2 with ingredients of the medium and removes some of the oxygen which almost always remains despite elaborate precautions.

(a)

(b)

FIG. 3. (a) Disposition of tube, rubber stopper, pipette, and gassing needle during transfers with the pipette. (b) Withdrawal of the gassing needle as the nibber stopper is slipped into place.

Culture tubes are 16 x 150 mm, made of heat-resistant glass, with a thickened constricted neck taking a No. 00 rubber stopper (Fig. 4a).

(4 (b) FIG.4. (a) Method for holding tube, stopper, gassing needle, and bent capillary pipette for picking colonies. (b) Displacing the dead air space prior to use of a syringe.

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R. E. IIUNGATE

An &-free gas stream (Fig. 2b) terminating in a sterile tube must flow into both the dispensing and the receiving vessel at a rate sufficient to exclude air. The rubber tube conducting each gas stream is attached distally to a conduit which can easily be flame sterilized and which is of a diameter small enough to permit partial insertion of the rubber-stopper closure while the conduit is still in the neck of the container. A 5-inch 20-gauge syringe needle locked to the cotton-filled barrel of a I-ml Luer-Lok syringe serves as a suitable conduit (Fig. 3a). T h e flange at the open end of the barrel is cut off to allow attachment of the rubber tube. T h e syringe and needle are dry sterilized prior to initial use and thereafter the needle portion is flamed as necessary. Shorter needles are more convenient with shallow vessels, but it is important that the end of each needle be inserted well into the vessel to avoid a vortex at the neck which could draw in air. The distance below the open neck should be about five times the diameter of the neck. T h e flow of gas should be sufficient for the purpose at hand but excess should be avoided, not only for economy but also because the gas is so often a source of oxygen. I t is useful to verify the rate of gas flow either by testing in a beaker of water prior to flame sterilization, by noting its force in the flame as the needle is sterilized, or by inserting the sterile end momentarily into the liquid in the gassing container. T o displace air from a 16x 150 mm culture tube containing 5 to 10 ml of medium, a flow rate of about 5 ml of gas/second, continued for 5 seconds, is satisfactory. For simply excluding air a slower rate is adequate. T h e medium is best transferred with a serological pipette to which a rubber mouth tube about 2 feet long is attached (Fig. 2b). Manipulation of the pipette by the mouth tube permits better exclusion of oxygen than do customary techniques. The pipette is inserted in the dispensing container, and 02-free gas drawn through it before the liquid is sucked in. T h e rate at which gas is sucked into the pipette should not exceed the rate at which the gas enters the vessel. With C02, the taste discloses when the air has been displaced. T h e pipette is closed by squeezing the rubber tube between the thumb and the first joint of the forefinger (Fig. 2a). The pipette is held by the other three fingers. After a volume slightly in excess of that required has been drawn in, the quantity of solution in the closed pipette can be easily adjusted to the desired mark by changing the angle between thumb and forefinger. With practice, a culture tube, a gassing needle and the pipette can all be simultaneously and appropriately manipulated (Figs. 3a, b).

4. Closing the vessel (Fig. 3b) This is done by inserting the stopper with the thumb and finger of one hand, holding the tube with the other three fingers, with the gassing needle

125

IV. ROLL TUBE ANAEROBIC TECHNIQUE

still in place. The stopper should be loose enough to let the gassing needle be withdrawn with the other hand without moving the stopper. If the stopper is inserted too tightly, as the needle is pulled out the pressure against the needle may break the glass of culture tubes without a thickened neck. The stopper and vessel neck are held firmly with one hand so that friction of the needle as it is withdrawn does not change the position of the stopper in the neck. Gassing is continued for a moment, then the needle is quickly pulled out, as light but firm pressure with the other hand pushes the stopper into place, closing the tube. Both hands can then be used to seat the stopper more firmly. The stopper seats readily if a little of the medium is tipped up to moisten the inner end and a twisting motion is used to force the stopper into its final position with the basal solid portion just inside the narrowest part of the neck of the tube. The twisting motion gives less tube breakage and if a break occurs the fingers are less likely to be cut. The removal of the needle should be rapid, as the end comes by the stopper. A slow removal can inject air into the vessel. Similarly, in opening an anaerobic vessel, the needle should be rapidly plunged to the optimal depth as soon as the stopper is removed. The stopper may first be loosened with both hands but not opened. Then it is pulled out with one hand as the gassing needle is slipped in with the other. Butyl rubber stoppers are used to close the vessels (Hungate et al., 1966). Butyl rubber is quite impermeable to oxygen and the permeability

(a)

( b)

FIG.5. (a) Inserting the needle into the recess in the rubber stopper. (b) Removal of sample from anaerobic tube into anaerobic syringe.

to other gases is less than with most types of rubber. Recessed (Fig. 5a) butyl rubber stoppers, No. 00, can be obtained from Arthur H. Thomas

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H. E. IIIJNGATK

Co., l’hiIddphi;i, U.S.A. Solid butyl-rubber stoppers can be obtained at reasonable price on spccial ordcr from most manufacturers. When asepsis is critical, it is well to place a cap of aluminium or other material over the top of culture tubes prior to sterilization, and to maintain it in place except when the tube is opened.

B. The nature of the culture medium T h e composition of the medium is determined as for any other cultures, with the exception that the buffering capacity must be increased to take care of acid fermentation products, and reducing agents must be included. I t should be clear that in designing the culture medium simulation of the natural habitat is often essential for success if organisms of unknown nutritional requirements are to be isolated directly from nature (Hungate, 1963). Ingredients of the medium can be added before heat sterilization except for reducing agents and heat-labile materials. Inorganic salts are usually provided to balance the high concentration of sodium bicarbonate in the buffer. In practice it has been convenient to store a three-times concentrated balanced inorganic salts solution in two separate flasks; one, the A solution containing the acid phosphate plus the other salts-Solution A

NaCl KH2P04

6 3 3 0.6

g g g g 0.6 g

(NH4)2S04 MgS04 CaClz Water

1 litre

Solution B contains 3 g K2HP04 per litre. One part of A and one part of B together compose one third of the culture medium. Carbon dioxide-bicarbonate is a more common buffer in nature than is phosphate. Concentrations up to one per cent are not usually toxic. Unless other buffers are abundant the p H of the medium is determined by the relative concentrations of carbon dioxide and bicarbonate. A solution of 0.5% sodium bicarbonate, equilibrated with an atmosphere of carbon dioxide, has a p H of about 6.7. Other desired acidities can be obtained by using carbon dioxide and bicarbonate concentrations calculated from the Henderson-Hasselbach equationpH

=

6*52+10g

molar conc. bicarb. molar conc, dissolved COz

Sodium bicarbonate in a 10% aqueous solution is sterilized by filtration because heating converts it to sodium carbonate, H20, and CO2.

IV. ROLL TUBE ANAEROBIC TECHNIQUE

127

This raises the pressure within the culture medium container, increasing the risk of stoppers blowing out. A period of equilibration after sterilization is necessary to allow reconversion of the ( 2 0 2 , H e 0 and carbonate to bicarbonate. Conversion of bicarbonate to carbonate also increases the alkalinity of the medium and the extent to which salts of phosphate and sulphate precipitate. If no bicarbonate is added prior to autoclaving, and the vessels are filled with carbon dioxide, the salts do not precipitate. In our laboratory, hydrogen (Mylroie and Hungate, 1954), sodium thioglycolate, cysteine, sodium sulphide, hydrogen sulphide and dithionite have been used as reducing agents. T h e hydrogen is used with a platinum catalyst. The sodium thioglycolate and cysteine can be heat sterilized and stored under carbon dioxide or other gases. A 17; solution of dithionite is sterile. T h e sodium sulphide must be stored under hydrogen or nitrogen since it is very alkaline, and carbon dioxide is rapidly absorbed, creating a vacuum which facilitates the entrance of air. Acidification to pH 6.7 converts sodium sulphide to hydrogen sulphide. T h e Eo’ at p H 7 is -0.571 for the reaction HzS+S+2H++Ze. Sterile hydrogen sulphide gas can be drawn off in a syringe from a Kipp generator and is perhaps the best and most convenient reducing agent for stringent anaerobes, though in most cases also cysteine is needed. Final concentrations of reducing agents can be 0.02 to 0.05% ; with higher concentrations inhibitory effects may be encountered, particularly with dithionite. For routine use we have employed chiefly a combination of 0.03% (w/v) cysteine hydrochloride and 0.25 ml of H2S gas per 10 ml of medium. The bicarbonate and the reducing solutions are added to the medium just before it is inoculated. Heat-labile nutrients can also be added just before inoculation. The initial volume of the medium before autoclaving should be diminished by the volume of reagents to be added later. A final volume of 4.5 ml per tube containing 1.2% agar gives a usable thickness of agar after the tube is rolled, and is convenient for serial dilutions. A low concentration (0.0001%) of resazurin is included in the culture medium to indicate oxidation-reduction potentials above -0.042. Others (Bryant, 1963) have used phenosafranine, with a redox potential (Eo‘, pH 7.0 = 0.252) better suited for registering low potentials. Benzyl viologen (Eo’, pH 7.0 = -0.359) has an appropriate potential but is toxic at the concentrations needed to give a detectabIe colour.

C. Sterilization 1. Steam pressure sterilization Large batches of O r f r e e medium can be sterilized in any container strong enough to withstand the pressure within the vessel as the autoclave

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K. E. IIUNCATE

cools. For quantities up to 5 litres, round-bottom boiling flasks can be used. T h e stopper must be securely wired in place. In many cases large quantities of medium can also bc sterilized in open containers which are removed from the autoclave and gassed as soon as removal is safe, I t is easier to maintain anaerobiosis in large rather than in small liquid volumes. During sterilization, culture tubes containing medium can be held in a vertical rack with a lid which must not be removed until the tubes have cooled to a safe temperature. 'The test tube racks of the Labtool Specialties Co. of Ypsilanti, Michigan, U.S.A., can be used. Stoppered tubes containing sterile medium can be stored for long periods without oxidation, but they may gradually oxidize due to slight permeability or leaks in the seal. Gradual diffusion of COz or Hz out of the container lowers the internal pressure and increases the chance of penetration of air.

2. Filtration Filtration can be accomplished anaerobically by using filters designed for application of pressure rather than of vacuum. Air is first eliminated by passing 02-free gas in either direction through the filter assembly and then gassing both inlet and outlet orifices during transfer of the medium to the filter. If the 02-free gas source is under sufficient pressure it can be used to supply the pressure for filtration, but it is necessary to be sure that the gas conduit between the gas cylinder and the filter will withstand the pressure to be applied.

D. Obtaining pure cultures 1. Inoculation T h e roll tube method cannot be used quantitatively for surface inoculation because of the vertical surface during incubation. It can be used to obtain growth of surface colonies by inoculating a liquid suspension of cells and rotating the tube to spread them over the surface. T h e excess liquid and cells drain to the base of the tube and do not affect the growth of cells left above on the agar surface, provided the tube is never tipped too far toward a horizontal position. T h e roll tube method is best suited for inoculation just prior ta solidification of the melted medium. Most of the colonies develop in the agar but as they increase in size many reach the gas interface and continue development as surface colonies. They may also reach the wall of the culture tube and grow as a film between agar and glass. T h e appearance of the colonies in each of these locations differs markedly from the appear-

IV. ROLL TUBE ANAEROBIC TECHNIQUE

129

ance in the other locations and may give the inexperienced worker a false impression of impurity. Careful cxamination under a binocular dissecting microscope can reveal the exact location of colonies and assist the interpretation of their form.

2. Picking colonies Colonies can be picked by touching with an inoculating needle and transferring the adhering cells quickly to the subculture or other preparation. This transfer exposes the cells to oxygen and since they are in a very thin layer on the needle, sensitive cells may be killed. It is better to transfer some of the agar along with the cells. This may shield them and increase the viability. A platinum-iridium inoculating needle can be flattened at the end to form a microspade or spatula. The wire can be bent several mm from the flattened end to an angle which allows the flattened end to be pushed under a colony to lift it out. A little agar block containing the colony should first be cut out with the sharp edges of the microspade. A steady hand is needed during the cutting and also as the block is removed. If the freed block touches the agar or glass after it has been taken up it usually adheres and comes off the microspade. The microspade technique is useful and economical for routine transfers of pure colonies and others which are separated from contaminants by several mm. Transfer with a Pasteur pipette is better for difficult isolations and for protecting the inoculum from air during transfer. The capillary of a sterile cotton-plugged Pasteur pipette is drawn out to a finer capillary at a distance from the main shank which will permit reaching the colony when the pipette is inserted into the tube (Fig. 4a). The diameter of the finer capillary should be about equal to that of the colony to be picked. This capillary is cut off about two inches from the main capillary shank and is bent at a right angle by momentarily placing it horizontally in a very small flame (the flame of a match is enough). With a piece of broken porcelain the capillary is scratched about 5 mm distal from the bend and is cleanly broken at the scratch. It is lightly flamed, i.e., enough to sterilize but not enough to melt the tip, and the plugged end attached to the rubber mouth tube. The capillary portion of the pipette is inserted into the culture tube alongside the gassing needle, and some 02-free gas is sucked in. The tip is then placed on the agar surface over the colony to be picked, and forced down into the agar as gentle mouth suction is applied. The colony is drawn into the capillary together with a little agar and some liquid that squeezes out of the agar when it is broken. The picked colony is rapidly introduced into the subculture tube and a little molten agar (cooled to 45°C) drawn in. This, with the colony, is then '7

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R. E. HUNGATE

extruded onto the dry wall of the culture tube and is drawn in and out of the capillary tip to disperse the cells, but no gas is blown out of the capillary. A few gas bubbles do little damage, too many can introduce some oxygen. Additional molten agar is drawn up and ejected onto the inoculum on the wall in order to avoid solidification. T h e pipette is withdrawn, the stopper is held in place and inserted as the gassing needle is withdrawn, and the tube is tilted back and forth a few times to mix the inoculum. Too rapid tilting forms gas bubbles which may persist during solidification of the agar and confuse the identification of colonies. T h e inoculated agar is solidified in cold water either in a mechanical tube roller (such as that supplied by Astell Laboratory Service Co., Catford, England) or by hand. Ice water is preferable for obtaining a firm agar, and the speed of the mechanical roller holds the agar in an even film until it is hard. If rolling is done by hand it is necessary to stop the motion just before solidification occurs, since otherwise the forming gel is broken and the resulting solidified agar is weak and ragged. During incubation, water of syneresis drains down from within the thin agar on the vertical walls of the tube. It does not usually disturb the surface colonies but the accumulation of liquid in the base of the tube makes it necessary to avoid spreading the bacteria in the basal liquid by undue tilting of the tube during examination and handling.

3. Dilution The concentration of colonies in the tube receiving the inoculum is usually not low enough to permit picking of an isolated colony. It is necessary to inoculate several dilutions of the material. This dilution can be directly in the tubes of molten agar at a temperature of about 45"C, or special dilution medium can be made up and used for dilution prior to inoculation into the culture tubes. Such dilution medium, if used, should be prepared with all the anaerobic precautions described for the culture media, and small volumes (0.1 ml) should be used for inoculation in order to maintain agar concentrations in the culture tubes. Once the inoculum has been placed in the first tube, further dilutions are advantageously made by means of a hypodermic syringe. T o permit this, we use recessed No. 00 butyl rubber stoppers (Catalogue No. 8820-B of the Arthur H. Thomas Co., Philadelphia, U.S.A.) or ordinary No. 00 butyl rubber stoppers in which a recess about 3 mm in diameter has been bored with a high speed drill in the centre of the top, extending about two-thirds of the length of the stopper. T h e 21-gauge needle easily penetrates the reduced thickness of rubber, and if locked to the syringe it does not pull off as the needle is withdrawn. In non-quantitative transfers to

IV. ROLL TUBE ANAEROBIC TECHNIQUE

131

agar tubes, the dead space in the end of the syringe can be transferred and the same syringe used for all the dilutions. Usually four tubes serially inoculated in this way will give sufficiently well isolated colonies in one of the dilutions. The syringe method is very rapid and convenient also for transfers of broth cultures (Fig. 5b), using the relatively inexpensive plastic syringes now available. Just before the syringe is used, the air in the dead space should be eliminated by inserting the needle in a stream of &-free gas (Fig. 4b) issuing from the orifice of a sterile Pasteur pipette with most of the capillary portion removed.

4. Criteria of purity As with almost all aerobic bacteria growing on the surface of agar, colonies of different anaerobic bacterial species almost always differ in size, texture, colour, transparency, margin, shape, consistency, or other characteristics. Close examination under the dissecting microscope can usually detect contaminants. Keen observation is needed to avoid conclusions of impurity based on differences due to location of the colony in the agar. I n the roll tube the proportion of colonies on the surface or between agar and glass is usually greater than in the similarly inoculated Petri dish because the agar layer is thinner. The history of a culture is the best index of its purity. If all cultures derived from a single colony exhibit only colonies of one type, the number of polonies in the tubes of a dilution series decrease in accordance with the dilution, and there is no microscopic evidence of impurity, then the culture can be assumed to be pure. When speed of isolation is critical, as in clinical examination of specimens, a single colony picked from a high dilution is often pure and can be subjected immediately to further tests, but its isolation should be continued to ensure that it actually was pure. With methanogenic, some cellulolytic, and some other anaerobic bacteria difficult to grow, it is as well to satisfy the above criteria of purity rigorously at an early stage of the investigation.

E. Storage of cultures This has not been studied systematically and extensively for anaerobes grown with the roll-tube technique. The problem of preservation is no different from that of other culture methods, except that the stoppered tubes can be stored almost indefinitely without drying out or becoming oxidized. The same advantages apply also to storage of sterile uninoculated tubes of media. Many strains of anaerobic bacteria remain viable for months if the culture is frozen quickly and held at the temperature of dry ice.

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F. Applications of anaerobic methods to various media Most media can be used to study anaerobes with little modification from aerobic methods. Reducing agents and some carbon dioxide and bicarbonate should usually be added. Some media, if prepared and inoculated with care for anaerobiosis, may be sufficiently reduced to start growth without adding reducing agents, particularly if inoculated heavily, but if nutritional requirements are under study it is well to subculture enough times to ensure that growth is not due to materials transferred with the inoculum. Many anaerobes require carbon dioxide. When acid production is to be measured it may be desirable\ to reduce the concentrations of carbon dioxide and sodium bicarbonate. G. Other applications The roll tube method may readily be adapted for the cultivation of micro-organisms requiring special gas atmospheres, such as hydrogen/ oxygenlcarbon dioxide mixtures or methane/air mixtures. REFERENCES Bryant, M. P. (1963).J. Anim. Sci., 22, 801-813. Hungate, R. E. (194n.J. Bact., 55, 631-645. Hungate, R. E. (1950). Buct. Rew., 14, 1-49. Hungate, R. E. (1963). In “The Bacteria” (Ed. Gunsalus, J. C., and Stanier, R. Y,), Vol. IV, Chap. 3, pp. 95-119. Academic Press, New York. Hungate, R. E. (1966). In “The Rumen and Its Microbes”, pp. 23-30. Academic Press, New York. Hungate, R. E. (1966). J. Bact., 91, 908-909. Mylroie, R. L., and Hangate, R. E. (1954). Can?. Microbial., 1, 55-64. Smith, P. H., and Hungate, R. E. (1958).J. Bact., 75, 713-718.

Rumen Bacteria P. N. HOBSON The Rowett Research Institute, Bucksburn, Aberdeen, Scotland I. Introduction: The Rumen Environment

. . .

.

11. Media for Isolating and Counting Rumen Bacteria . A. Habitat-simulating media . B. Media for isolating and counting bacteria with special biochemical properties . C. Non-habitat-simulating media . . 111. Media for Identifying and Determining SpecificProperties A. Morphology and growth ranges . B. Biochemical tests . C. Growth factors .

.

IV. Defined Media for Growth of Specific Strains of Rumen Bacteria V. Maintenance of Cultures

VI.

.

Continuous Cultures and “Artificial Rumens ”

References

.

133

135 135 138 143

. . . . .

147

.

147

.

148

*

144 144

145 146 146

1. INTRODUCTION: T H E RUMEN ENVIRONMENT

All herbivorous animals depend on micro-organisms for digestion of plant constituents which are not attacked by the digestive enzymes secreted into the mammalian gut; cellulose, pectic materials, hemicelluloses, etc. In herbivores the gut has developed in such a way as to form a fermentation chamber in which proliferation of micro-organisms and digestion of food can take place. This fermentation may precede the true gastric digestion or it may take place after such digestion. Ruminants have the former type of digestive system, in which the stomach is made up of four compartments, the first and largest, the rumen, being the one in which microbial digestion takes place. Because of the economic importance of ruminants the rumen has been the principal site for study of food-digesting bacteria, and, again because of their economic importance, the principal ruminants studied have been cattle and sheep. However, bacteria like those of the rumen occur in the intestines of ruminants and non-ruminant herbivores,

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and in omnivorous animals such as man; and their detailed study awaits only the application of the techniques of rumen microbiology to these sites. A recent review of microbial digestion of foodstuffs in ruminants and non-ruminants is that of Howard (1967). A much more detailed description of food digestion in the rumen and the bacteria and protozoa concerned is given in the book by Mungate (1966). This book is recommended as an introduction to the subject and a source of references to original papers, and only a brief description of the rumen environment will be given here. T h e rumen is essentially a continuous culture of long turnover time, about a day, in which micro-organisms are mixed with incoming foodstuffs by contraction and expansion of the rumen wail and by rumination. Flow through the system is provided by the ingress of food taken by the host animal and the large volumes of saliva secreted by the animal, and by rumen movements which move on the mass of organisms and undigested food to the omasum and abomasum. Although some air must be swallowed by the animal during feeding and oxygen may diffuse through the rumen wall from the blood stream the rumen environment is essentially anaerobic. T h e mass of digesta has a relatively large capacity for oxygen uptake (Broberg, 1957), most likely owing to the action of facultative anaerobes and some anaerobic bacteria contained in the rumen flora, and the Eh of the digesta is usually about -150 mv. T h e gas phase above the liquid digesta is composed principally of carbon dioxide, from the bicarbonate of the saliva and from bacterial fermentation products, and methane, which is a bacterial fermentation product. A typical analysis of rumen gas is (yo): hydrogen, 0.18; hydrogen sulphide, 0.01 ; oxygen, 0.56; nitrogen, 7.00; methane, 26.76; carbon dioxide, 65.35 (McArthur and Miltimore, 1961). Ruminant saliva provides a large amount of bicarbonate, as well as other ions, and a typical analysis of sheep parotid saliva is (rneq/litre)-Na+, 177; K+ 8; Ca++, 0-4; Mg++, 0-6; P, 52; C1-, 17; HC03-, 104 (McDougall, 1948). Microbial fermentation of foodstuffs produces acids and these are neutralized by the saliva, and in the conventionally-fed ruminant the rumen p H varies from about 6 to 7, being generally about 6.5. Most rumen bacteria grow best in this p H range, but some methods of feeding, principally those where large amounts of starchy materials are given, may produce rumen p H values below 6 and the bacteria present tend to be of the more aciduric types. T h e instantaneous concentrations of soluble substances in the rumen reflect a balance between their formation (e.g., soluble sugars by microbial hydrolysis of insoluble starch or cellulose, amino-acids by hydrolysis of food proteins, ammonia by deamination of amino-acids, acids by fermentation of carbohydrates) and their removal either by microbial growth, passage

V. RUMEN BACTERIA

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with the ingesta to the intestines, or absorption through the rumen epithelium into the blood-stream of the host animal. The concentration of soluble sugars and polysaccharides in the rumen fluid is generally low and many of the bacteria are attached to insoluble plant particles (largely cellulose) or starch granules. The Auid is also low in amino-acids, and ammonia is generally the microbial nitrogen source present in excess. A relatively large concentration (about 0.1 M) of the three lower volatile fatty acids, acetic, propionic and butyric is present, together with small concentrations of Cg and C6 acids and branched chain isomers of these and butyric acid. Formic, lactic and succinic acids are also primary products of foodstuff fermentations, but these are rapidly utilized in secondary fermentations and converted to methane and acetic and propionic acids. Similarly any hydrogen formed as a primary fermentation product is utilized in secondary reactions, principally in the formation of methane. The rumen fluid contains about 1010-1011 bacteria per ml and many of these bacteria exist in a symbiotic relationship with others, on which they depend for providing primary nutrients, such as sugars from polysaccharides, or for growth factors such as the volatile fatty acids or vitamins. Any system of culturing rumen bacteria must take this into account, as well as the general conditions appertaining to the rumen environment. ’No successful cultures of the important rumen bacteria were obtained until the late 1940’s when Sijpestcin, Gall and her co-workers, and Hungate successfully cultured some of the bacteria. The “habitat-simulating” media and the anaerobic techniques used by Hungate (1950) have proved most useful in culturing rumen bacteria and media based on his general principles have been used in most subsequent studies. Many of the media to be described fall into this category. ’ A complete list of media or papers dealing with relevant topics is not given. A selection of media and references is given, but further references may be found in the papers and books cited.

11. MEDIA FOR ISOLATING AND COUNTING RUMEN BACTERIA A, Habitat-simulating media

These media are designed principally to culture the anaerobic bacteria not cultured by using the usual anaerobic techniques or media, see Willis

(this Volume, p. 79). However, they will support the growth of other anaerobic and facultatively anaerobic bacteria normally present in the rumen. Such media can be divided into two groups: those designed for

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viable counts of all the bacteria present in rumen samples, and those for viable counts of groups with specific biochemical properties. Certain morphologically distinct rumen bacteria cannot be cultured and there may be many non-culturable bacteria in the mass of small bacteria usually making up the background of any slide of rumen contents; it is a general observation that viable counts are generally only a small proportion (less than 10%) of total counts.

1. General techniques for prefaring media, diluting fluids and dilutions All habitat-simulating media are prepared, dispensed and inoculated under oxygen-free gas streams using the techniques described by Hungate (thisVolume, p. 117). T h e basal medium constituents are sterilized by autoclaving and sodium bicarbonate, sodium sulphide, cysteine and sugars are usually added as filter-sterilized concentrated solutions under anaerobic conditions. Cysteine is usually used as the hydrochloride and concentrations given generally refer to this salt. For full details of medium preparation the original papers should be consulted. Diluting fluids used are generally similar to the basal medium composition, although in some cases dilutions have been made in the actual medium. Diluting fluids are discussed in the original papers and also by Hungate (this Volume, p. 117). For observations and sub-culturing, colonies can be picked from solid media under a stream of COz. Suitable portions of liquid media can be rediluted to obtain pure cultures.

2. Media for complete viable counts Only two of the many media used will be described here; both are similar, but were developed independently and tested in America or in the author’s laboratories over a number of years. Each supports the growth of a wide variety of rumen bacteria, but will not necessarily grow the methanogenic bacteria unless the bacterial fermentation produces some hydrogen or formic acid. Although many types of rumen bacteria have now been extensively studied and a number of growth factors (principally volatile fatty acid mixtures) are now known, rumen fluid is the most convenient source of such factors, known and unknown, for inclusion in a medium for counts of rumen bacteria. In some media rumen fluid is described as “strained”. I n this case only the larger feed particles have been filtered off on surgical gauze. This fluid gives a medium containing many small particles and colonies may be difficult to see. T h e rumen fluid described as “clarified” has been centrifuged at high speed to remove bacterial cells and gives a clear medium. T h e clarified is as good nutritionally as strained fluid. Since the nitrogen requirements

137

V. RUMEN BACTERIA

of the bacteria can be ammonia alone, ammonia plus amino-acids, or only amino-acids, a mixed nitrogen source is included in the medium (the rumen fluid acts as a nitrogen source). Almost all cultured rumen bacteria need a carbohydrate energy source and glucose is the most generally utilizable sugar. However, some of the cellulolytic and amylolytic bacteria (e.g., ruminococci and Bacteroides amylophilus) will ferment only the relevant disaccharide, cellobiose or maltose, and these are also included, but all sugar concentrations should be kept low. Some species such as VeilZmella alcalescens (V. gazogenes) will not ferment carbohydrates and lactate is included as substrate for these. Salts based on the mixture originally used by Hungate (1950), which in turn is similar in composition to ruminant saliva, are included. Sodium bicarbonate is added as buffer and also to simulate ruminant saliva. A reducing agent and redox indicator are also included and the medium is equilibrated with, and incubated under, a carbon dioxide atmosphere. Carbon dioxide is an essential growth factor for many types of rumen bacteria; it is fixed into fermentation products or cell components such as the carboxyl groups of amino acids.

Medium 1. This was devised by Bryant and Robinson (1961a)Glucose Cellobiose Soluble starch Agar Resazurin Minerals 1 Minerals 2 NazC03 Cysteine HCI-NasS (of a solution containing2 * 5 "/ow/v of each) Centrifuged rumen fluid Distilled water

0.25 g 0.25 g 0.05 g 2.0 g 0~0001g 3 -75ml 3 -75 ml 0.2g 1 ml 20 ml to 100 ml

The medium is equilibrated with, and incubated under, a gas phase of SOY0 COz, 50% H2, but by increasing the NaHC03 concentration to 0.4% a gas phase of 100% COz may be used (see Hungate, this Volume, p. 117 for techniques). Minerals 1 is (% w/v)-KzHP04, 0.6. Minerals 2 isKH2P04, 0.6; (NH4)zS04, 1.2; NaCl, 1.2; MgS04.7Hz0, 0.25; CaC12, 0.6. Medium 2. This was developed by Mann in the author's laboratories and has been in use for some years. It is referred to by Kurihara et al. (1967), and Eadie et al. (1967). The medium is equilibrated with, and incubated under, 100% coz.

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Bacto Casitone Bacto yeast extract Minerals (a) Minerals (b) Centrifuged (clarified)rumen fluid Agar (Difco) Resazurin Sodium lactate (70% w/v) Glucose Maltose Cellobiose Cysteine HCI Sodium bicarbonate Distilled water

1.og 0.25 g 15 ml 15 ml 20 ml 2.0 g 0.0001 g 1.og 0.2g 0 - 2g 0.2 g 0.05 g 0.4g to 100 ml

Minerals (a) contains (per 1000 ml)-K2HP04, 3.0 g. Minerals (b) contains (per 100 ml)-KH~P04, 3.0 g; (NH4)2S04, 6.0 g ; NaC1, 6.0 g; MgS04.7H20, 0.6 g; CaC12, 0.6 g.

Medium 3. Observations over a number of years have shown that the starch medium (Medium 6) can be used as a medium for total viable counts, since at least as wide, and often a wider, variety of bacteria grows in this medium as in Medium 2 (above). Why this should be so is not at present known. T h e rumen fluid used is usually obtained from a cow or sheep fed on a mainly grass or hay diet. This material is easier to centrifuge than that from animals on high concentrate diets. There is no definite evidence that rumen fluid from the animal species being studied is necessary, although there may be exceptions. For instance our own (unpublished) work showed that the same counts were obtained on Medium 2 from rumen samples from hill sheep and deer when media containing deer or sheep rumen fluids were used in culturing each sample.

B. Media for isolating and counting bacteria with special biochemical properties Although the media described below are intended to be differential and to enable bacteria with specific properties to be counted and isolated, in all cases rumen bacteria without these properties grow either in the basal medium or on breakdown products of the selectivc substrate, so that further tests, preferably in which a chemical determination of the property can be made, are needed to identify the bacteria. In addition, bacteria exhibiting the property in a weak form may not be identified in some of the media described.

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V. RUMEN BACTERIA

1. Cellulolytic bucteria (a) Solid media. In these a finely divided cellulose preparation is suspended in agar in a roll-tube culture. Cellulolytic bacteria are identified by zones of clearing around colonies. A number of cellulose preparations have been used, e.g., ball-milled filter paper, fiiter paper powders commercially prepared for chromatography, ball-milled cotton, acid-treated and ball-milled cotton. None of these substrates are the same as naturally occurring cellulose and their use may tend to over-estimate, if anything, the number of cellulolytic bacteria present in the rumen fluid. Halliwell and Bryant (1963) used a medium containing cotton fibres in testing for true cellulolytic activity in pure strains of rumen bacteria, but this medium could not easily be used for counting bacteria (see Part 111).

Medium 4. This was devised by Hungate (1957), and is made up as followsStrained (not clarified) &men fluid KH2P04 K2HPO4 MgS04 CaClz NaCl (NH4)2S04 Cysteine HCI Sodium bicarbonate Resazurin Cellulosepowder Agar Distilled water

30 ml 0.02 g 0.03 g

0.01 g 0.01 g 0.1 g

0.1 g 0.02 g 0.5 g 0~0001g 1g 2.0g to 100 ml

Equilibrated with, and incubated under, 100% COz (Hungate, this Volume, p. 117). The cellulose is filter paper finely wet-ground in a pebble mill. This medium is typical of the solid, cellulose powder media which have been used for isolating and counting cellulolytic bacteria. Commercial cellulose powder (MN 300, Camlab Ltd., Cambridge) has also been used (Kurihara et al., 1967). Incubation times may be up to 3 or 4 weeks. (b) “Liquid” media. A difficulty which arises with solid media is that the cellulose particles are comparatively large compared with the size of colonies and may not be entirely digested. This leads to indeterminate zones of cellulolysis and an indeterminate count of cellulolytic bacteria. The following medium has been extensively used with ten-fold dilutions of rumen contents and cellulolysis can easily be seen. Subcultures can be made from the liquid or from portions of the filter paper.

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Medium 5 . This was deviscd by Mann (1968), and is made up as followsBacto casitone Bacto yeast extract Centrifuged (clarified)rumen fluid Minerals (a) Minerals (b) Resazurin Cellobiose Cysteine HC1 Sodium bicarbonate DistilIed water

0.25 g 0.06 g 10 ml 15 ml 1 5 ml 0~0001g

0.025 g 0.05 $3 0 . 4g to 100 ml

T h e cellulose is added as a sterile filter paper strip suspended in the medium and this shows pitting and finally disintegration where cellulolytic organisms are active. Minerals (a) and (b) are as in Medium 2. Incubation time is 21 days and the medium is equilibrated with, and incubated under, 100% CO2.

2. Amylolytic bacteria Starch hydrolysing bacteria may be counted in anaerobic roll tubes containing starch after the tube is flooded with iodine-potassium iodide solution, but starch hydrolysis is a very rapid reaction compared with cellulolysis and amylases appear to diffuse rapidly from the colonies, so that the cultures should be tested after only a short incubation and nonamylolytic colonies may be found in the zones of starch hydrolysis. A suitable medium (devised by Mann) which has been in use for some years in the author’s laboratory is given below, but many non-amylolytic, as well as the amylolytic, bacteria grow in the cultures.

Medium 6 (Kurihara et al., 1967), is basically the same as Medium 2, (above) but with the lactate, glucose, cellobiose and maltose replaced by 0-5 g soluble starch added as a solution in 15 ml of water (dissolved and sterilized by autoclaving at 5 lb/sq in. for 30 min). Incubation time is up to 3 days, but less time may be needed to get distinct zones of starch hydrolysis, as these rapidly spread.

3. Proteolytic 6acteria I n a casein-containing medium such as that described below proteolytic colonies can be recognized by changes in the casein surrounding the colony, although some bacteria which produced no change in the casein agar were found to be proteolytic when tested in a liquid medium (Abou Akkada and Blackburn, 1963).

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Medium 7 . This was devised by Blackburn and llobson (1962), and by Abou Akkada and Blackburn (1963)Minerals (a) Minerals (b) Cysteine HCl Casein Centrifuged(clarified)rumen fluid Tryptose Agar (Difco) Resazurin Sodium bicarbonate Distilled water

15 ml 15 ml 0.05 g 0.5 g 10 ml 0.3 g 2.5 g 0.0001 g 0.5 g to 100 ml

Minerals a and b are as described for Medium 2 (above). T h e medium is equilibrated with, and incubated under, 100% C02. T h e sugars added are either glucose, or xylose, maltose and cellobiose (0.1% (w/v) each). Casein is Glaxo Casein C, acid precipitated, washed with water, dissolved in dilute NaOH, neutralized, and freeze-dried to give an easily-soluble powder.

4. L+olytic bacteria Only one species of anaerobic, lipolytic, rumen bacterium has so far been described. It may be counted in the medium described below, but the colonies are extremely small. Zones of clearing of the oil emulsion are also very small and may only be seen under magnification.

Medium 8. This was devised by Hobson and Mann (1961), and is made up as follows-

Minerals (a) Minerals (b) Centrifuged (clarified) rumen fluid Cysteine HCl NaHC03 Resazurin Linseed oil Agar (Difco) Distilled water

15 ml 15 ml 40 ml 0.05 g

0.4g 0'0001 g 1.0ml

2E to IOOml

The medium is equilibrated with, and incubated under, 100% C02. Minerals (a) and (b) are as in Medium 2, above. T h e linseed oil is added as 2 ml of a SO% (v/v) sterile emulsion in rumen fluid to the medium at 70°, and the medium is shaken to disperse the oil before dispensing.

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5 . Methanogenic bacteria Methanobacterium ruminantium was the first methanogenic bacterium to be shown to occur in sufficient numbers to be of importance in the rumen. Viable counts and isolations are made from dilutions of rumen contents in the following medium. T h e usual anaerobic techniques (Hungate, this Volume, p. 117) are used. In the first tests the medium was reduced with cysteine or sodium dithionite. However, growth of methanogenic bacteria was erratic because the poising capacity of the medium with dithionite was low, and the methanogenic bacteria need a low Eh for growth. A better method of ensuring a low Eh in the medium was found to be to grow Escherichia coli in the cultures before inoculation with rumen contents. In the final method the E. coli were heat-killed before the rumen contents inoculum was added. Medium 9. This was devised by Smith and Hungate (1958) and contains the followingK2HP04 KHzPOi NaCl NHiCl MgS04-7HzO CaCla Resazurin NaHC03 Rumen fluid (clarifie:d) Agar Sodium pyruvate Sodium dithionite Distilled water

0.1 .g 0.1 g 0.2 g 0.1 g 0.01g 0.01 g 0.0001 g

0.6 g 30 ml 2g 0.1 g 0.003 g to 100 ml

The medium is equilibrated with, and incubated under, an atmosphere of 80% hydrogen, 20% carbon dioxide. After tubing in 5 ml amounts an inoculum of 0.1 ml of a 48 h culture of E. coli in Brucella Broth (Albimi) is added and the tubes incubated at 39" for 12 h. T h e E. coli is then killed by heating the tubes at 75" for 45 min, the agar melted and the tubes held at 45" until inoculated (through the bung with a hypodermic syringe and thin needle) with a dilution of rumen contents. On incubation, methanogenic colonies continue to increase in size for a week or two whilst nonmethanogenic bacteria attain a maximum colony size in a few days. Methanogenisis also leads to a pressure decrease in the atmosphere of the tubes.

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6. Media for other isolations A number of solid media containing specific sugars have been used to try selectively to isolate bacteria fermenting these sugars. The media have been basically similar to those described for total viable counts, but with only one sugar added. Examples of such media are the one used by Hobson and Mann (1961) containing glycerol, the xylan-containing media of Hobson and Purdom (1961)’ a pectin-containing medium used in the author’s laboratories, and the saponin-containing medium of Gutierrez et al. (1958). In each case these media cultured bacteria fermenting the chosen substrate, but non-fermenting bacteria also grew in the media and as fermentation of the substrates could not be definitely detected by any means applicable to the roll-tube cultures, colonies from the higher dilutions had to be subcultured into liquid media containing the same substrate in which fermentation could be tested by physical (pH change) or chemical (loss of substrate, formation of fermentation products) means.

C. Non-habitat-simulatingmedia As well as the anaerobic bacteria needing specialized growth conditions the rumen contains, especially under certain feeding regimes or conditions of malfunction such as bloat, bacteria which are commonly cultured from other habitats and these may be counted and identified from growth in media developed for investigation of these other habitats, using wellestablished anaerobic methods. Such bacteria will also grow in many of the specialized anaerobic media described above. Dilutions used to inoculate the previous media may also be used for the following media. 1. Lactobacilli Media that have been used successfully in rumen work are described by Rogosa (1951a, b), (the SL medium) and by de Man and Sharpe (1960).

2. Streptococci Suitable media are described by Medrek and Barnes (1962) and Barnes (1956). Streptococcus bovis is the principal Gram-positive streptococcus so far found in rumen contents. 3. Veillonellas A suitable medium for growth of veillonellas is described by Rogosa (1956); this will also support the growth of Peptostreptococcus elsdenii and

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the two genera can be distinguished by colonial form. The medium was later modified by Rogosa (1964) and this paper should also be consulted.

4. Peptostreptococci These will grow in Medium 3 above, or in Medium 10 below. Medium 10. This semi-solid thioglycollate medium was devised by Hobson et al. (1958) and contains the following-

I

Bacto casitone Bacto yeast extract L-cystine Thioglycollic acid NaCl Agar (Difco) Sodium lactate solution (70%) Resazurin, 0 * 1% (w/v) solution Distilled water

sl? 2g

0.3 g 0.12 ml 2g 0.3 g 5 . 7 ml 0.4 ml 400 ml

pH 6-9-7.0

The medium is sterilized by autoclaving at 121" for 15 min.

111. MEDIA FOR IDENTIFYING AND DETERMINING SPECIFIC PROPERTIES The bacteria growing in the non-habitat-simulating media (Section IIC) and presumptively identified thereby may be further tested using the appropriate standard media described elsewhere. T h e specialized bacteria growing in the habitat-simulating media may be further tested using modifications of the media described in Section 11. Classification of the rumen bacteria is not very definitive at the moment. However, the properties of species so far described may be found in Hungate's (1966) book or Bryant's earlier review (1959), as well as in original papers. A. Morphology and growth ranges Morphology may be determined in liquid media used for isolation or fermentation tests, or after growth on slope cultures. Motility and flagellation may often be best determined from the syneresis liquid at the bottom of slope cultures or young cultures in a liquid medium can be used. Formation of a starch-type polysaccharide is a feature of some rumen bacteria and this can be tested for in iodine-stained wet preparations at different

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stages of growth. Growth at various temperatures can be tested using the habitat-simulating media, but alteration of the initial culture pH from the usual value of about 6.8 is difficult because of the presence of bicarbonate and a 100% COz atmosphere.

B, Biochemical tests Fermentation of sugars may be tested for by observing growth and fall in pH in media such as Media 1 or 2 above, in which the mixed sugars are replaced by one sugar, usually at a concentration of 0.5%. A medium without sugar may be used as control, but this will generally show some growth, and with certain bacteria sugar fermentation may only depress the p H to about 5.8, Changes in p H may be determined by an indicator in the medium or, better, by p H meter. Fermentation products may also be determined in such media and products of fermentation of specific substrates may be used in classifying the bacteria. Loss of substrate may be determined by chemical methods. An example of this was in isolating xylan-fermenting bacteria (Hobson and Purdom, 1961). Extent of hydrolysis of cotton cellulose and paper cellulose was determined by a gravimetric method by Halliwell and Bryant (1963). Pectin hydrolysis was tested by observing disintegration of potato slices placed on slopes of anaerobic media by Purdom (1965) after initial isolation of bacteria in media containing a soluble pectin preparation. Chemical determination of loss of uronic acid in media containing a polygalacturonic acid was also used in examination of pectinolytic bacteria. Hydrolysis of protein in liquid cultures was determined by chemical estimation by Abou Akkada and Blackburn (1963) and Blackburn and Hobson (1962) using casein-containing media. Liquefaction of gelatin may be tested in a medium modified by including geIatin-charcoal tablets (Hobson and Mann, 1961, based on Kohn, 1953). Deaminative activity is sometimes difficult to determine as ammonia is a constituent of the medium for many rumen bacteria. Net change in ammonia concentration in the medium, which contains amino-acids, usually in the form of a protein hydrolysate, is measured chemically. Examples of this are given in the papers by Abou Akkada and Blackburn (1963) and Bladen et al. (1961). Lipolysis of linseed oil can be tested for by ether extraction and titration of the higher fatty acids produced from a linseed oil emulsion liquid medium (Hobson and Mann, 1961). This differentiates higher fatty acids from the fermentation products of the glycerol moiety of the oil. Other properties such as production of HzS and reduction of nitrate

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may be tested in the habitat-simulating media, if necessary with additional substrates added.

C. Growth factors Specific growth factors of the bacteria cultured in the non-habitat-simulating media, i.e., those belonging to well-defined families, are determined using standard procedures. Media for determining specific growth factors of the bacteria cultured only in the habitat-simulating media are based on these media. One of the first tests is that of growth without rumen fluid. This could be tested with a medium such as Medium 2 above less the rumen fluid. Growth without COz can be tested in a similar medium lacking NaHC03 and C02, but buffered by phosphate and incubated under nitrogen. A further standard test for these bacteria is growth with ammonia as sole nitrogen source. Further details will not be given here as schema for testing for bacterial growth factors in media of the habitat-simulating type are given in a number of the papers already quoted in this Chapter and in others dealing with rumen bacteria. Amongst these are those of Abou Akkada and Blackburn (1963), Bryant and Robinson (1961b, c; 1962), Pittman and Bryant (1964), Bryant et al. (1959), but many other papers, especially those describing new species of rumen bacteria, give details of media with which tests for growth factors were determined.

IV. DEFINED MEDIA FOR GROWTH O F SPECIFIC STRAINS O F RUMEN BACTERIA Many of the rumen bacteria isolated in the habitat-simulating media have only been grown in media such as those described for their isolation. This may have been because they need undefined growth factors or because they have not yet been investigated in detail. Some bacteria have been investigated in detail because of their specific properties, and this is especiially the case with the cellulolytic bacteria. For a number of strains of bacteria, defined (amino-acid), or semi-defined (protein hydrolysate) media have been described. “Strains” is used here as there is variation in growth requirements, as in other properties, amongst different isolates of the same species. This may be a reflection of the present state of classification of the rumen bacteria. These media will not be described here, but some papers giving details are as follows-Bryant and Robinson, 1961b, c; White et al., 1962; Pittman and Bryant, 1964; Blackburn, 1968; Hobson et al., 1967.

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V. MAINTENANCE OF CULTURES The lactobacilli, streptococci, etc., isolated from the rumen may be maintained under standard conditions. For instance, Peptostreptococcus elfdeniimay be maintained in Robertson’s Cooked Meat Broth stored at room temperature and sub-cultured about once a month. We have always used this medium prepared in the laboratory (see Mackie and McCartney, 1953), but dehydrated medium could be used. Many of the bacteria isolated in the habitat-simulating media may be maintained in media similar to the isolation media described and stored at 2’C. Sub-culture may be needed only at monthly or longer intervals, but with some strains weekly culture may be needed. No definite rules can be laid down. In general sub-cultures kept at -20°C have no longer a life than those kept at 2” C, and in many cases the life may be shorter. However, cultures rapidly frozen to -60°C may be kept for periods of siimonths or longer between sub-cultures, and this is one of the best methods of maintaining cultures. Some enzymic activities, such as cellulolysis or proteolysis may be lost on continued sub-culture even if the specific substrate is included in the medium. Freeze-drying of cultures of bacteria isolated on habitat-simulating media has been successful in. some cases. Suspension in glucose-serum or milk has been used (for example, Hobson and Mann, 1961). The methods of freeze-drying generally used may be applied to rumen bacteria (see Lapage et al., this Series, Vol. 3A).

VI. CONTINUOUS CULTURES AND “ARTIFICIAL RUMENS” As the rumen is a form of continuous culture a number of workers have attempted to establish in vitro continuous cultures of mixed rumen organisms with a volume of more cr less whole rumen contents as the starting “inoculum”. Unlike the chemostat type of culture a complete growth medium is not fed continually, but nutrients, usually ground animal feeds, are added at intervals, while a regular flow of “artificial saliva” serves to wash over excess cells and fermentation products. The culture vessel also may be made of a dialysis membrane so that products of microbial action may be dialysed to an external salts solution, thus simulating the absorption of microbial products by the rumen epithelium in vivo. Difficulties have been found in the mechanics of this type of culture, especially the addition of solid nutrients, and apparatuses have varied in their degree of automated working. The cultures have also varied in their success in keeping the balance of the fforaof the original inoculum of rumen contents, and it may be said that in all cases some change in the bacterial and protozoal population has

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taken placc after somc hours or days. In many cases some readily enumerated munbers of the microbial population (such as protozoa) have completely disappeared, Nevertheless some of these cultures have simulated the rurnen function in such things as volatile fatty acids production to a high degree. One of the latest papers in which bacteriological investigations were made on a continuous culture artificial rumen is that of Slyter et al. (1966). No description of these cultures will be given here, but a number of references to papers in which artificial rumens are described is given by Hungate (1966). Chemostat types of continuous cultures of pure strains of anaerobic rumen bacteria have been used in a few cases not to simulate rumen conditions but to investigate the metabolism of these bacteria. T h e difficulty in prolonged cultures is keeping the redox potential at a low value by adapting the specialized culture conditions of the habitat-simulating media (above) to continuous flow operation. Hobson and Smith (1963), Hobson (1965 a, b), Hobson and Summers (1966, 1967) have described apparatus used for cultures ot' up to about 2000 hours duration and some results of cultures of different species of anaerobic bacteria. The main problems associated with this work are in the attainment of anaerobic conditions. Medium reservoirs are constructed so that media may be prepared and kept under an oxygenfree carbon dioxide atmosphere and the apparatus is thoroughly purged with carbon dioxide before the medium is pumped in. Piston-type pumps are used, as the flexible tubing used in the usual peristaltic pumps is permeable to oxygen. T o cut down oxygen-transfer through rubber tubing (which can oxidize the medium sufficiently to prevent bacterial growth) rubber connections are made as short as possible by using metal (copper for gas lines, stainless steel for medium), or glass, tubing. Initially, selected heavy wall red-rubber tubing was used. Neoprene was found to be better, but the tubing tends to split on autoclaving where it is under tension, as on joints to glass tubing. Butyl rubber tubing is now available in a number of sizes and this has proved satisfactory both from the point of view of oxygen transfer and durability. Hungate (1963) described some results of a continuous culture of Ruminococcus albus and Kistner and van Zyl (1967) have recently described the adaptation of the cyclone column culture apparatus of Dawson (1963) to the culture of a Butyrivibrio and a strain of R. aibus. REFERENCES Abou Akkada, A. R., and Blackburn, T. H. (1963).J, gen. Microbiol., 31,461. Barnes, E. M. (1956).J. uppl. Buct., 19, 193. Blackburn, T. H. (1968).J. gen. Microbiol., 53, 27. Blackbum, T. H., and Hobson, P. N. (1962).J.gen. Microbiol., 29,69.

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Bladen, H. A., Bryant, M. P., and Doetsch, R. N. (1961). Appl. Microbiol.,9, 175. Broberg, G. (1957). Nord. VetMed., 9, 57. Bryant, M. P. (1959). Bact. Rew., 23, 125. Bryant, M. P., and Robinson, I. M. (1961a).J. Dairy Sci., 4,1446. Bryant, M. P., and Robinson, I. M. (1961b). Appl. Microbiol., 9, 91. Bryant, M. P., and Robinson, I. M. (1961~).Appl. Microbiol., 9, 96. Bryant, M. P., and Robinson, I. M. (1962). J. Bact., 84, 605. Bryant, M. P., Robinson, I. M., and Chou, H. (1959).J, Dairy Sci., 42,1831. Dawson, P. S. S. (1963). Can.J. Microbiol., 9,671. Eadie, J. M., Hobson, P. N., and Mann, S. 0. (1967). Anim. Prod., 9,247. Gutierrez, J., Davis, R. E., and Lindhal, I. L. (1958). Science, N.Y.,127, 335. Halliwell, G., and Bryant, M. P. (1963). J . gen. Microbiol., 32,441. Hobson, P. N. (1965a).J. gen. Microbiol., 38, 161. Hobson, P. N. (1965b).J. gen. Microbiol., 38, 167. Hobson, P. N., McDougall, I. E., and Summers, R (1967).J. gen. Microbiol., SO, i. Hobson, P. N., and Mann, S. 0. (1961). J. gen. Microbiol., 25, 227. Hobson, P. N., Mann, S. O., and oxford, A. E. (1958).J.gen. Microbiol., 19,462. Hobson, P. N., and Purdom, M. R. (1961).J. appl. Bact., 24,188. Hobson, P. N., and Smith, W. (1963). Nature, Lond., 200, 607. Hobson, P. N., and Summers, R. (1966). Nature. Lond., 209,736. Hobson, P. N., and Summers, R. (1967). J. gen. Mictibiol., 47, 5 3 . Howard, B. H. (1967). In “Symbiosis”, Volume I1 (Ed. S. R4. Henry). Academic Press, New York and London. Hungate, R. E. (1950). Bact. Rew., 14, 1. Hungate, R. E. (1957). Can. J. Microbiol., 3, 289. Hungate, R. E. (1966). “The Rumen and Its Microbes”, Academic Press, New Yorkand London. Kistner, A., and van Zyl, J. G. (1967). Can. J. Microbiol., 13, 455. Kohn, J . (1953). J. din. Path., 6, 249. Kurihara, Y . ,Eadie, J. M., Hobson, P. N., and Mann, S. 0.(1967).J. gen. Microbiol., in the press. McArthur, J. M., and Miltimore, J. E. (1961). Can.3. Anim. Sci., 41, 187. McDougall, E. I. (1948). Biochem.J., 43, 99. Mackie, T. J., and McCartney, J. E. (1953) “Handbook of practical bacteriology”, p. 192. E. and S. Livingstone, Ltd., Edinburgh. Medrek, T. F., and Barnes, E. M. (1962). J. appl. Bact., 25, 159. Mann, S. 0. (196Q.J. appl. Bact., 31, 241-244. de Man, J. C., and Sharpe, M. E. (1960).J. appl. Bact., 23,130. Pittman, K. A., and Bryant, M. P. (1964). J. Bact., 88,401. Purdom, M. R. (1965). Private communication. See Howard, B. H. (1967). Rogosa, M. (1951a).J. dent Res., 30,682. Rogosa, M. (1951b).J. Bact., 62,132. Rogosa, M. (1956).J. Bact., 72,533. Rogosa, M. (1964). J. Bact., 87, 162. Slyter, L. L., Bryant, M. P., and Wolin, M. J. (1966). Appl. Microbiol., 14, 573. Smith, P. H., and Hungate, R. E. (1958).J. Bact., 75, 713. White, D. C., Bryant, M. P., and Caldweli, D. R.(1962). J. Bact., 84, 822.

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CHAPTER V I

Methods for the Gram-negative Non- sporing Anaerobes ELLAM. BARNES Food Research Institute, Norwich, Norfolk, England I. Methods . A. General precautions to be taken . , 3. Media . C. Growth tests, morphology, etc. . D. Biochemical tests . E. Isolation techniques . F. Isolation media , References

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. . . . . .

152 152 153 153 154 158 158

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159

The anaerobic non-sporing Gram-negative bacteria which are included within the family Bacteroidaceae are amongst the least studied of the bacteria associated with man and other animals. They occur in large numbers in the alimentary tract and are frequently associated with necrotic lesions. There is at present no generally recognized system of classification, identification or nomenclature. This situation has arisen partly because these anaerobes have been studied by quite different groups of microbiologists, notably dentists studying the oral flora, pathologists isolating organisms from necrotic lesions, and agricultural microbiologists studying the flora of the rumen and, more recently, the intestine. Thus there have been a number of detailed studies of particular organisms but little attempt has been made until recently to relate these organisms to one another. T h e techniques used for their growth have also varied from the strictly anaerobic techniques developed by Hungate (this Volume, p. 117) for the rumen strains, to the more traditional methods discussed by Willis (this Volume, p. 79). So far, it has been found that all of these anaerobes can be grown by the Hungate roll-tube technique, providing that the medium is suitable; but there are a number of anaerobes, such as those isoiated from the rumen, which cannot be grown using the methods described below. Evidence suggests that there are organisms in the intestinal flora which resemble the rumen strains and they will also only be isolated by using the Hungate technique. Anyone embarking on a study of the intestinal flora should

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include this particular technique together with any other methods used. I n many cases, however, particularly in medical laboratories where all types of organisms are being handled, it is often easier to adapt standard procedures than to use an entirely different technique. Also, where possible the simplest method should be used and if an organism can be grown easily without using the Hungate technique then it seems unnecessary to take the extra precautions. T h e methods described below have been developed for the isolation and growth of strains of Fusobacterium, Sphaerophorus and Bacteroides. Typical strains are Fusobacterium polymorphum as described by Baird-Parker (1960)) Sphaerophorus necrophorus as described by Fievez (1963) and Bacteroides fragilis which has been described by Beerens et al. (1963) under the name of Eggerthellu convexa. Other important species are Bacteroides melaninogenicus (Bergey’s Manual (7th ed. 1957)) and Fusiformis nodosus (Beveridge, 1934 and 1967).

I. METHODS

A. General precautions to be taken 1. Growth on agar plates All the plates used for isolations or purification must be poured on the day of use or stored in an anaerobic jar. A suitable type of anaerobic jar is the Baird and Tatlock jar fitted with the catalyst as described by Willis (this Volume, p. 79). Many of these anaerobes will not grow without carbon dioxide and it is recommended that 10% carbon dioxide is included with the hydrogen. It is important that the last traces of oxygen are removed rapidly; thus the presence of a catalyst is essential. Incubation in an atmosphere of hydrogen plus 10% carbon dioxide without taking this precaution is not sufficient for growth in many cases. Plates must be placed in the anaerobic jar within 10 min of inoculation.

2.Growth in liquid media This can be carried out without using an anaerobic jar providing that the following precautions are taken(1) A semi-solid reducing medium should be used such as RCM or VL (described below). T h e agar is added at 0.05-0-1% to prevent convection currents and to reduce the adsorption of oxygen from the air. (2) It should be distributed in tubes or small bottles which can be tightly sealed. For example the 1 oz screw-capped Universal container (28 ml) made by the United Glass Bottle Manufacturers Ltd., Great Britain. (3) Only a small head space should be left in the bottle.

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153

(4)The inedium should be held in a boiling watcr bath for at least 20 min to remove oxygen, then cooled, and inoculated iinmediatcly. (5) A large inoculum of about 0.25 in1 should be used.

B. Media Reinforced Clostridial Agar or Broth ( R C M ) (Hirsch &’ Grinsted, 1954) Peptone (Evans Medical Ltd., Speke, 10 R Liverpool) Beef extract (Lab-Lemco, Oxoid) 10 g Yeast extract (Oxoid) 3g Cysteine hydrochloride 0.5 g Glucose 5g 5g Sodium acetate (anhyd) Soluble starch Ig Agar (New Zealand) 0.5 g Distilled water 1 litre pH 7’2-7.4

For the solid medium use agar (New Zealand) 12 @re. V L medium (modified from Beerens et al., 1963) Tryptone (Oxoid) NaCl Beef extract (Lab-Lemco, Oxoid) Yeast extract (Difco) Cysteine hydrochloride Glucose Agar (New Zealand) Distilled water pH 7.2-7’4

10 €! 5g 3g 5g

0.4 g

2.5 g 0.6 g 1 litre

For the solid medium use agar (New Zealand) 12 g/litre. VL blood agar. Supplement with sterile horse blood, 5% (Burroughs Wellcome Ltd., 183-193 Euston Road, London, N.W. 1). Baial medium. Use VL medium without glucose. B.G.P. medium. (For maintenance of stock cultures), Basal medium with glucose 0.1%, and Na2HP04 0.4%. All of the liquid media are sterilized at a pressure of 15 lb/sq. in. for 15 min in 19 ml lots in screw-capped 1 oz bottles. C. Growth tests, morphology, etc. Most strains of Fusobacterium, Sphaerophorus or Bacteroides will grow on VL or RCM agar or in VL or RCM broth. However, the growth of Bactevoidesfragilis and other strains is much better on VL blood agar, so this is

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the agar medium of choice. With a few exceptions blood or serum supplements are not generally required in the liquid media but the addition of haemin (1.0 ,ug/ml) to either VL or RCM broth stimulates the growth of the Bact. fragilis strains. For Bact. metaninogenicus all media, whether liquid or solid, are supplemented with laked blood 5% (prepared by freezing and thawing whole blood) and menadione 0.5 ,ug/ml. Fusformis nodosus will not grow in the media described above unless it is supplemented with dried powdered sheep or ox hoof 2%. For details, reference should be made to Thomas (1958). T h e media should be inoculated with 0.25 ml of a 24-48 h culture and incubated at 37" for 1-7 days.

D. Biochemical tests There are certain tests which are considered particularly important for the differentiation of Fusobacterium, Sphaerophorus, and Bacteroides (Barnes and Goldberg, 1968). These are the terminal pH in glucose broth, the production of butyric acid from glucose, and the presence of threonine deaminase. Stimulation by bile is also considered important for the identification of many of the Bacteroides strains but care must be taken over the bile used for this test, as not all batches (particularly of the dried product) are effective (Beerens, personal communication). Additions are made to the Basal medium for the various biochemical tests. After holding in a boiling water bath for 20rnin to remove oxygen and cooling, the media are inoculated with 0.25 ml of a 24-48 h culture and incubated for 1-7 days according to the test and the organism. 1. Carbohydrate fermentation All carbohydrates are sterilized by filtration and added to the basal medium to give a concentration of 0.25% (1% is frequently used by other workers). Change in p H is determined by use of the capillator technique (B.D.H. Ltd).

2. Detection of volatile f a t t y acids T h e volatile fatty acids produced from glucose fermentation may be determined by the method of Guillaume et al. (1956), as modified by Charles and Barrett (1963). I n this test the organism is grown for 7 days in 20 ml of basal medium with added glucose (0.25%). T h e culture is then centrifuged, the supernatant adjusted to p H 2.0 with normal HCl, and the non-ionized volatile fatty acids extracted by shaking with 30 ml ether. The extraction is repeated twice more and the bulked ether extracts are washed

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155

with an equal volume of distilled water. 1.5 ml of morpholine (33% v/v) is added to the washed ether after which the ether is removed at room temperature by evaporation under vacuum. The 1-2 mi containing the morpholine salts of the volatile fatty acids are then analysed chromatographically, the solvent used being that of Guillaume et al. (1956), i.e., butanol, 30 parts; cyclohexane, 30 parts; propanediol (1 : 2), 10 parts; ammonia, 0.7 parts; morpholine, 0.07 parts; distilled water 3.5 parts. After separation for 22-26 h, the paper is allowed to dry at room temperature for 20 min before spraying with the indicator solution (Universal indicator (B.D.H.), 10 ml, ~ / 1 NaOH, 0 2 ml). Gas chromatographic techniques are also being used (Moore et al., 1966).

3, Threonine deaminase test Guillaume et al. (1957) showed that certain of the Gram-negative anaerobes utilized threonine. The threonine was deaminated with the production of a-keto butyrate and ammonia; the a-keto butyrate could then be converted to propionic acid, hydrogen and carbon dioxide. There are three methods of performing this test at the present time; by the detection of the propionic acid produced from the threonine (Beerens et al., 1959), by the detection of ammonia (Suzuki et al., 1966) and by the nile-blue reduction test of Beerens and Tahon-Castel(1965). This latter test is the least specific as reduction of the nile blue does not necessarily indicate that the threonine deaminase pathway is involved. (a) Detection of propionic acid

Reagents Threonine solution. DL-threonine loo/,, in phosphate buffer (pH 7.0). Sterilize by filtration. Store in a refrigerator. Dilute to 1% using boiled and cooled phosphate buffer (pH 7.0) immediately before use. Phosphate buflerpti 7.0. Solution 1, KHzP04 9.075 g/lOOO ml. Solution 2, Na~HP04.2Hz0,11.870 g/1000 ml. Mix 370 ml of Solution 1 with 630 ml of Solution 2. Sterilize in 150 ml lots at 15 lb psi for 20 min. Before use, boil for 1 h in water bath. Method Inoculate 200 ml of boiled and cooled VL broth with 10-15 ml of an actively growing culture in RCM or VL broth. Cover with 20-25 ml sterile liquid paraffin (specific gravity 0.830-0.870). Incubate for 45-48 h at 37°C. Centrifuge to separate the cells. The cells are washed with 150 ml of boiled and cooled phosphate buffer and reccntrifuged. They are then transferred to a screw capped 1 oz bottle using 6-8 ml buffer. After centrifuging, the

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buffer is poured off and 10 ml 1% threonine are added to the cells and the culture incubated at 37°C for about 40 h. After incubation the culture is centrifuged and the supernatant analysed for propionic acid as follows. Using a Markham still for the steam distillation (Markham, 1942), 8 ml are pipetted into the apparatus followed by 1 ml conc. sulphuric acid, About 50 ml of distillate are collected and titrated with ~ / 1 NaOH 0 using cresol red (1% aqueous) as indicator. A further 1 ml of ~ / 1 NaOH 0 is then added and the solution concentrated under vacuum to 1-2 ml using a rotary film evaporator in a water bath at 70°C. T h e concentrate is put through a small column of cation exchange resin (BIO-RAD, A G SOW-X8, 200-400 mesh, Hydrogen form). T h e size of column is calculated according to the total amount of ~ / 1 NaOH 0 used (1 g of resin is sufficient for about 17 ml ~ / 1 NaOH). 0 T h e eluate is collected in sufficient N-morpholine to neutralize all the acid present and the fatty acids are thus present as the morpholine salts. T h e fatty acids are then determined chromatographically as described above and the presence or absence of propionic acid recorded. (b) Detection of ammonia and nile-blue reduction Reagents Threonine solution. DL-threonine 10% in phosphate buffer (pH 7.6). Sterilize by filtration. Store in a refrigerator. Dilute in buffer to give a concentration of 2% for the detection of ammonia. Phosphate b u f k p H 7.6. Prepare Solutions 1 and 2 as above. Mix 132 ml Solution 1 with 868 ml Solution 2. Sterilize by autoclaving in 20 ml lots in 1 oz bottles. Nile B h e . Dissolve 0.005 g/ml in boiled and cooled phosphate buffer (pH 7.6). Use immediately. Nessler’s reagent. Dissolve 8 g potassium iodide and 11.5 g mercuric iodide in 20 ml water and adjust to 50 ml. Add 50 ml6N NaOH. Mix and allow to stand for 24 h. N.B. T h e water must be ammonia free. Allow the reagent to settle before use. Protect from the light.

Method T h e organism is grown in 2 x 1 oz bottles each containing about 20 ml VL broth (or in 40ml in a suitable container). When the organism has grown sufficiently, i.e., is at the end of the log phase (this may vary between 18 and 40 h), centrifuge and wash with phosphate buffer (pI4 7.6), combining the cells into one 1-02 bottle. T h e culture is again centrifuged and further washed with buffer. Finally resuspend with 1 ml buffer (pH 7.6).

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Into a plugged sterile test tube 180x 9 mm introduce the following solutionsDetection of anznronia Control (1) Phosphate buffer (2) Test organism Test (1) Threonine 2% in buffer pH 7.6 (2) Test organism

0.5 ml 0.5 ml 0.5 ml 0.5 ml

Seal with a stopper or cover with foil. Incubate in a water bath at 37°C for 3 h. After incubation add 0.05 ml20% NaOH and 0.1 ml Nessler’s reagent. Mix gently. A red colour or reddish-brown sediment in the tube indicates threonine deamination, whilst the control tube remains colourless. Nile-blue reduction Control (1) Nile blue (2) Phosphate buffer (3) Test organism Test (1) Nile blue (2) Threonine 10% in buffer 7.6 (3) Test organism

0.5 ml 0.5 ml 0.5 ml 0-5 ml 0.5 ml 0.5 ml

The tubes are sealed immediately with 2 mi agar. Incubate in a water bath at 37°C. Read the tests at intervals up to 24 h. A positive reaction is shown by the decoloration of the nile blue, to a yellowish colour. T h e control tube should stay blue.

4. Bile stimulation Bile 10% is added to BCP medium (Beerens and Castel, 1960). 5. Hydrogen sulphide Hydrogen sulphide is detected by the development of a black colour in either the basal medium or BGP to which has been added ferrous sulphate 0.02%, and sodium thiosulphate 0.03%.

6. Indole Indole is tested for in cultures grown in the basal medium or, where growth is poor, in BGP. Ether (1 ml) is added to the culture which is shaken vigorously to extract the indole, 0.5 ml of Ehrlich’s reagent paradimethplamidobenzaldehyde,4 g ; absolute alcohol, 38 ml; E-ICI, 80 ml, is added to the ether layer, followed by a saturated solution of potassium persulphate, a red colour indicating the presence of indole.

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7. S k i m milk medium Skim milk medium containing cysteine hydrochloride O * 0 8 ~(pH o 7-4) is used to detect acid (by use of litmus paper), digestion, and clot.

8. Gelatin liquefaction Gelatin liquefaction is determined in BGP containing 15% gelatin.

9. Antibiotic sensitivities Antibiotic solutions are added to one of the growth media to give the required concentrations.

E. Isolation techniques I n the isolation of these organisms all operations should be carried out as quickly as possible to avoid prolonged contact with oxygen. Dilutions should be prepared in a medium such as RCM or VL which has been held in a boiling water bath to remove any dissolved oxygen. Surface inoculation of the required medium is usually carried out by rapidly spreading 0.05 ml of the diluted sample over one half of the agar plate, two dilutions being spread on each plate. T h e plates are placed in the anaerobic jar as described above. Organisms within the family Bacteroidaceae differ markedly in their reactions to dyes and inhibitors, so different media have been developed for their isolation. Whilst strains of Fusobacterium and Sphaerophorus are resistant to dyes such as crystal violet and brilliant green, the strains of Bucteroides are inhibited by them. On the other hand the Bacteroides are resistant to antibiotics such as neomycin and kanamycin which may be inhibitory for some of the other organisms. For example, the chicken isolates of Goldberg et al. (1964), now designated “group 4”,grow well in the presence of ethyl violet or brilliant green but are inhibited by neomycin at levels well below those normally used for the isolation of the Bacteroides.

F. Isolation media Fusobacterium Medium (Omata and Disraely, 1956)

15 g 5g

Casitone (Difco) Yeast extract (Difco) Glucose NaCl L-cysteine Crystal violet Streptomycin Agar (Difco) Distilled water

5g 59

0.75 g 0.01 g 0.01 g 15 g 1 litre pH 7.2

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159

The streptomycin solution (final concentration 10 pg/ml) is added to the agar immediately before pouring the plates, together with serum 5% (Burroughs Wellcome Ltd., 183-1 93 Euston Rd., London, N.W.1, England). A somewhat similar medium has been described by Baird-Parker

(1957). Sphaerophorus. VL blood agar to which has been added sodium azide 0.05 mg/ml and ethyl violet 0.05 mg/ml or brilliant green 0.02 mg/ml (Fievez, 1963). The inhibitors are usually prepared as 1% aqueous solutions. The required concentration may be added to the medium hefore autoclaving. Bacteroides (Finegold et al., 1965). VL blood agar to which has been added 100yglml kanamycin or 100 pg/ml neomycin. Vancomycin 7-5 pg/ml may also be added to make the medium more selective. Bacteroides melaninogenicus (Finegold et al., 1965). VL medium containing laked blood 5% and menadione 0.5 pg/ml together with kanamycin 100 yglml. Unnamed isolates from poultry caeca. Amongst the unnamed Gram-negative anaerobes which are present in large numbers in the caecal flora of chickens and turkeys, the “group 4” isolates of Goldberg et al. (1964) may be isolated using RCM agar containing ethyl violet 0.05 mg/ml and sodium azide 0.05 mg/ml. It is essential to use a high level of carbon dioxide (10%) in the anaerobic jar. Failure to do this possibly explains previous isolation difficulties. Addition of 5% blood to this medium improves the isolation of certain strains but reduces the selectivity of the medium. REFERENCES I

Baird-Parker, A. C. (1957). Nature, Lond., 180, 1056-1057. Baird-Parker, A. C. (1960). J. gen. Microbiol., 22, 458-469. Barnes, E. M., and Goldberg, H. S. (1968). J. gen. Microbiol., 51, 313-324. Beerens, H., and Castel, M. M. (1960). Annls Inst. Pasteur, Paris, 99, 454-456. Beerens, H., Guillaume, J., and Petit, H. (1959). Annls Inst. Pasteur, Paris, 96, 211-216. Beerens, H., Schaffner, Y., Guillaume, J., and Castel, M. M. (1963). Annls Inst. Pasteur Lille, 14, 5 4 8 . Beerens, H., and Tahon-Castel, M. M. (1965). Annls Inst. Pasteur, Paris, 108, 682-684. Bergey’s Manual of Determinative Bacteriology (1957), 7th ed. (Ed. R. S. Breed, E. G. D. Murray, and N. R. Smith). The Williams and Wilkins Co., Baltimore, U.S.A. Beveridge, W. I. B. (1934). J. Path. Bact., 38, 467-491. Beveridge, W. I. B. (1967). Bull. 08.int. Epizoot., 67, 7. Charles, A. B., and Barrett, F. C. (1963). J . med. Lab. Technol., 20, 263-268. Fievez, L. (1963). “Etude comparbe des souches de Sphaerophorus necrophorus

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isolkes chez l’homme et chez l’animal”. Presses Academiques Europkennes, Bruxelles. Finegold, S. M., Miller, A. B., and Posnick, D. J. (1965). Ernuhihrungsforschung, 10, 517. Goldberg, H. S., Barnes, E. M., and Charles, A. B. (1964). J. B Q C ~87, . , 737-742. Guillaume, J., Beerens, H., and Osteux, R. (1956). Annls Inst. Pusteur Lille, 8,1342, Guillaume, J., Petit, H., and Beerens, H. (1957). Annls Inst. Pusteur Lille, 9, 68. Hirsch, A., and Grinsted, E. (1954). J. Dairy Res., 21, 101-110. Markham, R. (1942). Biochem. J., 36, 790-791. Moore, W. E. C., Cato, E. P., and Holdeman, L. V. (1966). Int. J. Syst. Buct., 16, 383. Omata, R. R., and Disraely, M. N. (1956). J. Buct., 72, 677-680. Suzuki, S., Ushijima, T., and Ichinose, H. (1966). rap. Microbial., 10,193-200. Thomas, J. H. (1958). Aust. wet. J.,34, 411.

r.

CHAPTER VII

1

Psychrophiles and Thermophiles T. D. BROCK

Department of Microbiology, Indiana University, Bloomington, Indiana, U.S.A. AND A.

H. ROSE

School of Biological Sciences, Bath University, Bath, England I. Introduction

.

11. Psychrophiles

.

A. Isolation . B. Maintenance of stock cultures C . Cultivationmethods

.

.

111. Thermophiles . A. Introduction . B. Heat exchangers . C. Aeration and evaporation control D. Miscellaneoustechniques

References

.

.

. . . . . . . . .

161

.

164 164 165 166 166

.

167

162 162 163 163

I. INTRODUCTION It is common practice among microbiologists to subdivide microorganisms into three categories based on the effects of temperature on growth; these categories are known as psychrophiles, mesophiles and thermophiles (Farrell and Rose, 1967a, b). The basis for this subdivision is largely historical. A great deal of the pioneer work in microbiology was done with organisms, particularly pathogenic or potentially pathogenic strains, that grow optimally between about 25" and 40°C. These microbes are referred to as mesophiles. When microbes which grow well at temperatures below and above the mesophile range were iso"lted, they were considered to be unusual and often exotic. Further research has shown quite clearly, however, that psychrophiles and thermophiles are certainly not uncommon and probably not exotic. There has never been, and probably never will be, any great measure of agreement on definitions for psychrophiles and thermophiles (Farrell and 8

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Rose, 1967a, b). Nevertheless, these remain useful subdivisions of the microbial world if, as Ingraham (1962) has pointed out, one asks no more of them than that they refer to microbes that grow well at low temperatures (psychrophiles), intermediate temperatures (mesophiles) and high tempera. tures (thermophiles). Detailed accounts of the physiology of psychrophiles and thermophiles have been made by Brock (1967), Farrell and Rose (1967a, b), and Farrell and Campbell (1969). This article is confined to a brief discussion of the methodology associated with isolation and study of psychrophiles and t her mophiles.

11. PSYCHROPHILES A. Isolation The most widely accepted definition of psychrophiles is that they are micro-organisms capable of growing well at 0°C within 1-2 weeks (Ingraham and Stokes, 1958; Stokes, 1963). The psychrophilic habit is widely distributed among micro-organisms. Many microbiologists recognize two classes of psychrophiles. The first of these-the “obligate” psychrophiles-not only have a lower minimum temperature for growth compared with mesophiles but also a lower maximum temperature for growth, which may be below 20°C. “Facultative” psychrophiles, on the other hand, have maximum temperatures for growth in the same range as those for mesophilic microbes, namely 30”-45”C. If psychrophilic microbes are to be isolated from naturally occurring materials, it is obvious that a temperature of 0”-2°C will, after a suitable incubation period, allow these microbes to grow and, at the same time, prevent growth of mesophiles. It is necessary, too, to ensure that the material never experiences a temperature of about 15°C or above if obligate psychrophiles in the sample are not to be killed. Over the years, many microbiologists interested in the isolation of psychrophiles have failed to take the precaution of keeping the temperature of the sample low. This applies particularly to the isolation of psychrophiles from oceans and to samples collected in polar regions of the world. Some microbiologists persist in using the pour-plate technique which, of necessity, exposes the microbes in the sample to a temperature of about 4O”C, although procedures have been published which lower this setting temperature (Parker et al., 1968). Unless they are examined in situ, samples from polar regions present many problems, particularly as the tendency is often to have the samples transported to laboratories in more temperate climates, a journey that invariably exposes the samples to high temperatures.

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163

For examining water samples for psychrophiles, undoubtedly the best technique is to pass the sample through a membrane filter that will retain the organisms and then to incubate the filter at a low temperature on suitablemedia. (See Mulvaney, this Series, Volume 1for details of membrane filtration techniques.) The ease with which these filtrations can be carried out in the field or aboard ship now makes this technique de rigeur. Using the technique, Stanley and Rose (1967) found obligate psychrophiles to be very common in lakes on Deception Island in Antarctica. The impression is that, once suitable precautions are taken in the handling of samples, obligate psychrophiles will be found to be very much more widespread than is presently thought.

B. Maintenance of stock cultures Stock cultures of micro-organisms are normally stored in or on solidified nutrient media at temperatures below the minimum for growth so that further growth of the organisms is prevented, thereby ensuring that metabolic waste products do not accumulate in high concentrations in the culture. Thus stock cultures of mesophilic and thermophilic microbes are usually stored in laboratory refrigerators (2'4°C) ;thermophiles can also be stored at laboratory temperature (18"-22°C). In our experience, stock cultures of psychrophiles are best stored in a laboratory refrigerator. As these are at temperatures above the minimum for growth of psychrophiles, organisms in stock cultures may grow quite extensively in a relatively short time. It is advisable, therefore, to transfer stock cultures of psychrophiles fairly frequently. If stock cultures of obligate psychrophiles are maintained at laboratory temperature even for a brief period, there can be a dramatic fall in viability because these temperatures are above the maximum for growth. In general, when working with psychrophilic microbes, it is advisable not to expose cultures for long periods to temperatures much above 5".

C. Cultivation methods Psychrophilic microbes are usually isolated and cultured on solidified nutrient media and, for this purpose, low-temperature anhydric incubators suffice. The review by Patching and Rose (this Series, Volume 2) describes the advantages and disadvantages of the various types of low-temperature incubator that are available. The main drawback in using these incubators is that it is difficult to avoid temperature gradients being set up inside the cabinet. The variatiens of temperature inside the cabinet may be as great as +, 2"-3"C, especially when the incubators are operated at near-zero temperatures and, under these conditions, it is impossible to obtain accurate data on the effect of temperature on growth of organisms.

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Such data can only be obtained using thermostatically controlled baths containing water or other liquid such as odourless kerosene which is kept stirred or agitated. Patching and Rose (this Series, Volume 2) have discussed in detail the different types of constant-temperature bath which can be used for incubating cultures of psychrophilic micro-organisms.

111. THERMOPHILES

A. Introduction The old idea that a thermophile is any organism capable of growing at

55°Cis rapidly disappearing. It is now quite clear that there is a continuum of organisms from those with optima near the mesophilic range to those with optima of 7Oo-75"C and perhaps even higher (Brock, 1967; Brock and Brock, 1968). When isolating and studying thermophiles, it is important to consider first the temperature of the natural environment where the organism is growing, and to reproduce this temperature in the laboratory. When this is done, organisms growing optimally at the temperature selected will usually be isolated. If 55°C is used as the isolation temperature, organisms growing optimally at 55°C will be isolated. However, it is not enough to consider only the temperature factor of the environment. Nutrient quality and quantity, pH value, salinity, and perhaps other factors, must all be properly adjusted for the habitat from which inocula are being taken. By basing isolation procedures on these principles, a whole new group of thermophilic bacteria has been discovered (Brock and Freeze, 1969), and more new groups perhaps await discovery. The past dominance of Bacillus stearothermophilus in thermophile research is due to two factors: use of 55°C as incubation temperature, and use of media fairly rich in organic constituents. By rejecting traditional isolation methods, our understanding of thermophiles should broaden considerably. The unique methodology involved in the study of thermophiles concerns temperature control, and the rest of this article will deal with that and certain ancillary problems. It is much easier to heat something up than to cool it down, and a wide range of devices of various sizes and price ranges can be obtained for this purpose. For much work, precision temperature control is not needed. However, when working at temperatures near the maximum temperature for growth of an organism, precise control is necessary, as an excursion of one degree above the upper limit may cause death. At the optimum temperature or lower, variation of f 1°C can easily be tolerated unless precise growth studies or physiological tests are being done.

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165

B. Heat exchangers Heat exchangers commonly used are metal (or plastic), water, oil and air. Temperature blocks using metal or plastic heat exchangers are now widely available commercially, and are the cheapest way of getting a wide range of incubation temperatures. Temperature control is not precise but is good enough for many purposes. The temperature blocks manufactured by some companies can be used through a range from about 30" to 100°C, although not all blocks have this wide a range and the specifications should be examined carefully before ordering. One disadvantage of temperature blocks is that a different metal block must be used for each size of test tube. However, this disadvantage is offset to some extent by the fact that the metal blocks are interchangeable, so that several sizes can be used in one heater. A variant of the temperature block is the temperature-gradient block, which provides a range of temperatures in one long bar. The advantages of a gradient block are minimal, since all temperatures available on the gradient block can also be obtained in separate temperature blocks, usually at lower cost. For instance, one temperature-gradient block sells for $900 and holds two tubes at each temperature. Individual temperature blocks, holding 10 tubes each, sell for about $35 each. Thus one can buy over 25 temperature blocks for the cost of one gradient block and have increased capacity and greater versatility. Analysed in these terms, a gradient block seems more like a gimmick than a useful device. Water baths are the mainstay of the thermal biologist, especially when precise temperature control is necessary or when flasks, bottles, and other odd-shaped containers.are to be incubated. A variety of types is available. The usual precautions to ensure adequate stirring should be made if it is important that all the vessels in the same bath are at the same temperature. At high temperatures, the greatest problem is evaporation of water from the bath. At temperatures of 70°C or higher, an uncovered bath will lose all of its water overnight. If an automatic water-level device is used, distilled or deionized water must be used to avoid a build-up of salts. A cover reduces evaporation greatly, but may make difficult the installation of aeration devices or other attachments. An alternative is to place a thin layer of paraffin oil on top of the water. This very effectively cuts down evaporation but can only be used in an unstirred bath. Some commercial water baths are available with automatic water-control devices, or a device can be built out of stock components. One company (New Brunswick Scientific Co., New Brunswick, N.J., U.S.A.) manufactures a water-bath shaker with automatic waterlevel control. One of us (T.D.B.) operated such a shaker satisfactorily for a year at 70°C before it developed a variety of problems, most of which have been difficultfor the company to rectify. As this shaker costs over $800, it should be bought for thermophile work only as a last resort (see p. 168). Water

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T. D. BROCK AND A. H. ROSE

is the most useful heat exchanger for large fermenters and can be provided directly from the tap. Unfortunately, most hot-tap waters run at temperatures of 60"-65°Cso that, if higher temperatures are needed, the temperature of the water will have to be further raised with an auxiliary heater; alternatively live steam may be circulated. However, there is no technical reason why thermophiles cannot be grown at high temperatures in fermenters. Recently in the Department of Microbiology at Indiana University, a run in a 200 litre fermenter was carried out at 70°C. At temperatures above S O T , water baths cannot easily be used due to excessive evaporation, and oil baths come into use. We have had little experience with these, perhaps being put off by the alleged carcinogenicity of many of the commercial oils and by the fact that, since oils do have a finite vapour pressure, their use in an enclosed room for a long period of time might be dangerous. Air incubators provide less precise temperature control but offer the advantages of cheapness and simplicity. They are especially useful for incubating Petri plates.

C. Aeration and evaporation control Thermophilic anaerobes are technically easier to culture than aerobes, but have been studied only rarely. With aerobes, special attention to aeration efficiency is necessary because at high temperatures the solubility of oxygen drops drastically. However, at high aeration efficiencies, evaporation of water from the culture medium becomes a problem. If the upper portion of the vessel is cooled, either by room-temperature air or by a water-cooled reflux condenser, evaporation can be controlled fairly well. For small vessels and short-term incubation (2-3 days), air cooling may be sufficient, but for long-term incubation, especially at high aeration efficiency, refluxing is the best. We have found that a very suitable arrangement for incubating 500-1000 ml volumes at 95"-1OO0Cis to use a round-bottom boiling flask with standard taper connections. If a three-holed flask is used, the central hole can be used for the reflux condenser, one of the side holes can be used for aeration, and the other for a thermometer. Heat can be supplied by a standard flask heater. Any evaporation is made up by addition of sterile distilled water.

D. Miscellaneous techniques 1. Agar plates The concentration of agar for use in Petri plates should be increased to 2.5-3.0% for use at temperatures of 60"-75"C, if undue spreading of colonies is to be avoided. Evaporation can be avoided by placing plates in a sealed

VII. PSYCHROPHILES AND THERMOPHILES

167

jar, but sufficient air space should be left. Such jars can be Tncubated either in air incubators or submerged in water baths. Evaporation from individual plates can be controlled by wrapping them tightly with plastic film such as Saran (Dow Chemical Co.).

2. Contamination Contamination by air-borne organisms is of little consequence in thermophile research, since most such contaminants are rapidly killed. However, there are a few potential contaminants in the laboratory. In addition, if a variety of thermophiles are being studied in the same laboratory, the usual aseptic precautions should be taken to avoid cross contamination. We make this point because one of us (T.D.B.) has noted that the quality of aseptic technique of people working with thermophiles deteriorates with time, sometimes with disappointing results.

3. Photosynthetic thermophiles Water is essential as a heat exchanger for culturing photosynthetic thermophiles to avoid localized heating effects from the lights. Glasswalled aquaria make the best water baths, since lights can be directed from the sides towards cultures immersed in various parts of the bath. Agar plates should also be incubated submerged in water baths. For growing large volumes of organisms, high intensity submersible lights which can be inserted into fermenters are available. A water-bath shaker fitted with a Lucite cover and light assembly is useful for growing photosynthetic thermophiles in shaker flasks. 4. Sample collection Although there is no current evidence for cold shock of thermophiles, this should be kept in mind as a possibility. If difficulty is experienced in obtaining successful cultures from a given thermal environment, it might be profitable to transport the sample to the laboratory in a vacuum flask, or to inoculate preheated culture media at the collection site. REFERENCES hock, T. D. (1967). Science, N . Y., 158, 1012-1019. Brock, T. D., and Brock, M. L. (1968). J . uppl. Buct., 31, 54-58. Brock, T. D., and Freeze, H. (1969).J. B a t . , 98, 289-297. Farrell, J., and Rose, A. H. (1967a). I n “Thermobiology” (Ed. A. H. Rose), pp. 147-218. Academic Press, London and New York. Farrell, J., and Rose, A. H. (1967b). A. Rev. Microbiol., 21, 101-120. Farrell, J., and Campbell, L. I,. (1969). Adv. microbial Physiol., 3, 83-109.

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Ingraham, J. L. (1962). In “The Bacteria” (Eds. I. C. Gunsalus and R. Y. Stanier), Vol. 4, pp. 265-296. Academic Press, New York. Ingraham, J. L., and Stokes, J. L. (1959). Bact. Rev.,23, 97-108. Parker, E. T., Bernsteinas, J. P., and Green, J. H. (1968). Appl. Microbiol., 16, 1794. Stanley, S. O., and Rose, A. H. (1967). Phil. Trans. Ry. SOC. B., 252, 199-207. Stokes, J. L. (1963). In “Recent Progress in Microbiology” (Ed. N. E. Gibbons) Vol. 8, pp. 187-192. University of Toronto Press, Toronto.

A design of water-bath shaker more satisfactory than that of the New Brunswick instrument (see this Volume, p. 165) has recently been placed on the market by Fermentation Designs (U.S.A.). This uses a magnetic drive and so avoids the problem, often experienced with the New Brunswick instrument, of leaks around the drive shaft. NOTE ADDED IN PROOF:

CHAPTER V I I I

Isolation, Growth and Requirements of Halophilic Bacteria N. E. GIBBONS Division of Biosciences, National Research Council, Ottawa, Canada I. 11.

Introduction

.

Extreme Halophiles . A. Occurrenceand economic aspects B. Growth requirements . C. Media D. Isolation and enumeration E. Large-scale production . F. Maintenance .

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.

111.

Moderate Halophiles . A. Occurrence . B. Ionic and growth requirements C. Media . D. Pathogenic halophiles .

References

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. . . . . . . .

169

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. .

.

180 180 180 181 182

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182

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170 170 171 174 177 178 179

I. INTRODUCTION Sodium chloride is needed by all warm-blooded animals, and its availability or scarcity has affected the patterns of settlement of vast areas (Block, 1963) and influenced the course of history. For centuries it provided the main means of preserving food and it remains an important preservative in many areas. Much salt for this purpose was obtained by evaporating sea water, and many centuries ago bacteria, presumably from the sea, adapted to life in the strong brines of the salt ponds (cf. Baas-Becking, 193 1). Marine bacteria have been distinguished from terrestrial bacteria by their sodium requirement (MacLeod and Onofrey, 1957) and they might thus be considered the beginning of a series of organisms with increasing sodium and/or chloride requirements. Although one definition of halophiles is that they are organisms that require more than the usual physiological concentration of salt (0.9%) for growth and survival, in this paper halophiles are organisms that need more than 3% NaCl.

170

N. E. GIBBONS

Various classifications have been proposed for halophiles, based on their salt requirements (Ingram, 1957; Larsen, 1962). Until more is known of the specific bases for their salt requirements, these must be regarded as classifications of convenience. The definitions in this paper are based on the upper and lower limits of salt required for growth, without regard for underlying metabolic mechanisms. Using this criterion, we regard moderate halophiles as organisms which grow in the presence of concentrations of salt ranging from 3 to IS%, the extreme halophiles grow only in media containing 15% or more NaCl. It is well to emphasize the definite requirement, which distinguishes the halophiles (salt-loving) from the salt-tolerant organisms able to grow in media without added salt and also in the presence of quite high concentrations of NaCl, as do many of the micrococci. Although several ions are involved in the nutrition of these organisms their foremost requirement for both growth and maintenance of stability is NaCl and, unless otherwise stated, throughout this paper salt and NaCl are synonymous. Halophilic yeasts and fungi are known but only bacteria will be considered here.

11. EXTREME HALOPHILES A. Occurrence and economic aspects Reddening of heavily salted products, such as fish, intestines (sausage casings) and hides, has been recognized probably since salting as a means of preservation began. Although references to what may be red halophiles occur from ancient times (cf. Baas-Becking, 1931), the biological nature of the problem was first mentioned by Farlow (1880). Bacteriological examinations using salt-containing media began with Hoye (1906), followed closely by the more detailed studies of Kellerman (1915), Klebahn (1919) and Harrison and Kennedy (1922). By this time the requirements of the microorganisms concerned for a high salt concentration, aerobic conditions and a temperature optimum around 42°C were recognized. With the decline of salt as a preservative, much of the economic incentive for studying these organisms disappeared. Work, however, continued in a few laboratories on the biochemistry and physiology of the halophiles and has been reviewed by Larsen (1962, 1967). The present interest taken in the unique properties of these organisms by bacterial physiologists, electron microscopists and space scientists is encouraging, The halophiles are examples of adaptation to a severe environment involving many interesting biochemical changes, of growth at low water activities and in such complex concentrated mixture of salts that little is known of the state of the ions or of the water. The best known organisms are the rods : Halobacterium salinarium, H . cutirubrum, H. halobium, H . trapanicum and the cocci, Micrococcus litwalk,

VIII. REQUIREMENTS OF HALOPHILIC BACTERIA

171

M. morrhuae. There is some question as to whether the cocci are to be regarded as micrococci or as sarcinae or whether both exist, and further study is required. In fact, more work is required on the classification of the whole group. Biochemical studies indicate that the rod forms at least are a distinct group. The primary source of these organisms is solar salt, prepared by evaporating sea water in open ponds (salterns). The reddening of the brine as it approaches saturation has been taken as an indication of incipient crystallization for thousands of years (cf. Baas-Becking, 1931). However, the reddening of salt ponds may be caused by other pigmented brine organisms. Because of the exposure to high intensities of light in the salterns, Larsen speculated that the red carotenoid pigments constituted a protective mechanism and that colourless forms were likely to be non-existent or rare. Dundas and Larsen (1962, 1963) showed that a wild red strain was more resistant to ultraviolet and sunlight than a colourless mutant. Recently, A. C. Allison (personal communication) has shown that H . cutirubrum is as resistant to ultraviolet and ionizing radiations as Micrococcus radiodurans. I have isolated colourless forms from the edge of saline lakes and from cultures of red halophiles received in this laboratory. However, little has been done with the white and other organisms that can be isolated from salt and salted products. Halophiles are also found in meat-curing brines, but these are mostly the moderate type. However, Hornsey and Mallows (1954, 1955) found in beef-curing brines organisms which grew well at 15-20% NaCl, and similar organisms were found by Ingram et al. (1957) in bacon-curing brines. These organisms grew only if unheated blood plasma or muscle press juice was supplied, and even then slowly to form very small colonies. They were present in large numbers (106/ml) and may have a rale in the curing process. However, they have not been studied further to my knowledge. B. Growthrequirements 1. Ionic The NaCl requirement of these organisms has been known for many years; Brown and Gibbons (6955) were probably the first to show that the sodium requirement was specific and NaCl could not be replaced by other salts. Growth did not occur when the NaCl concentration fell below 3.0 M. This limit could be lowered to 1-5 M if the NaCl is replaced by KCI, but no growth occurred with 1.0 M NaCl in saturated KCI. A requirement for C1 became apparent when other sodium salts would not support growth. This has been studied in more detail by Boring et al. (1963). Potassium was also essential, and Brown and Gibbons found 50-100 ppm allowed optimum growth. Rubidium could replace potassium, but

172

N. E. GIBBONS

concentrations of 10 to 100 times that of potassium were required for equivalent growth, and the colour of the organisms was pale, suggesting that potassium is required for normal pigmentation. It should be remembered that this work was done with static cultures in shallow layers, rather than shaken or aerated cultures, and the above amounts are probably suboptimal for growth under well aerated conditions. Magnesium was another essential ion; optimal growth was obtained with concentrations of 0.1-0.5 M Mg2+. At these concentrations, more of the rods appeared "normal", with fewer of the bizarre forms usually mentioned in morphological descriptions. At lower concentrations the organisms grew as spheres and seemed to adapt to the low concentrations of Mg2+. Oddly enough the spherical form was retained when these cells were returned to media containing the original concentration of magnesium. Using shake cultures, Sehgal and Gibbons (1960) found that 10 ppm of Fez+ increased the yield of cells. It also allowed better growth at concentrations approaching the lower limits of the NaCl requirement. Small amounts of manganese (0.05 ppm) provided some stimulation of growth and a marked increase in pigmentation.

2. Temperature Early workers had noted that the red halophiles grew best at temperatures around 40°C. Christian (1956) found that most rapid growth occurred between 37" and 45°C. Gibbons and Payne (1961) reported growth curves of five species of halophiles in aerated cultures at temperatures of 30"-55°C. T h e maximal rate of growth occurred at about W"C, but in some cultures maximum turbidity was obtained at slightly lower temperatures. Under the conditions used, Sarcina litoralis did not grow at 50°C; the three red rodshaped organisms did not grow at 55"C, but a colourless rod did. T h e rate and amount of growth depended on both temperature and salt concentration. As a rule, growth, as measured by turbidity, was better at the higher temperatures in concentrations of 20% or more salt. At the less favourable conditions, changes in morphology were noted which suggests that perhaps turbidity might not be a reliable method of estimating growth under a variety of conditions (see discussion below).

3 . Oxygen Practical men noticed that fish and other products did not redden as long as covered by brine, but that exposed portions often became coloured. In early studies, growth was obtained on agar, but not in broth, unless a solid interface was provided. Using sparged air, growth can be obtained in large volumes of liquid media and has enabled us to produce the large number of cells required for

VIII. REgUIREMENTS OF IIALOPIIILIC BACTERIA

173

chemical and biochemical studies (Kushner, 1966). However, nothing is known of the actual oxygen requirements of these organisms and, as their growth requirements seem paradoxical-aerobic organisms growing under conditions of temperature and salt concentration which markedly reduce the solubility of oxygen-further study seemed desirable. Dr S. M. Martin in our laboratories has found that in a 7.5 litre fermenter (4 litres of culture with stirring at 400 rev/min and air injection at the rate of 4 litres per min) the oxygen concentrations in the medium are practically zero throughout the log phase of growth, i.e., ca. 2% of saturation level or < 0.05 mg/litre. The solubility of oxygen in water at 40°C is 6.1 mg/litre, in 25% NaCl solution about 2 mg/litre and in the medium used probably even less. The extremely halophilic red rods are very susceptible to disruption by physical forces, particularly shear. There is thus a limit to stirring, pumping, etc., as a means of improving aeration. The effect of increased oxygen tensions is being examined. There is considerable interest in the cell envelopes of the red halophilic rods and for a time there was doubt that a true wall existed. The envelopes lack DAP and muramic acid, and there has been speculation as to exactly how the walls are held together (reviewed by Larsen, 1967). Recent work indicates that a wall can be distinguished by electron microscopy (Stoeckenius and Rowen, 1967; Cho et al., 1967) but it begins to disintegrate when salt concentration is lowered below 1.6 M. These chemical and physical characteristics of the walls cast doubts upon the present classification and one wonders whether these organisms can legitimately be grouped with the Eubacteria. 4. Anaerobes

To my knowledge, only one anaerobic halophile has been described. Baumgartner (1937) described as a strict anaerobe a non-motile rod, which would not grow in less than 4% NaCl, and had a requirement for optimal growth of 12-15% salt. He also mentioned facultatively anaerobic halophiles. While one might expect such organisms to occur because of the general low solubility of gases in brines, little information is available. Volcani (194-4) isolated several anaerobic bacteria from bottom sediments of the Dead Sea. These included lactose fermenters, methane producers, denitrifiers, sulphate reducers and cellulose decomposers. However, the information is scanty and apparently the organisms are no longer available (B. E. Volcani, personal communication). Reference has also been made by Etchells et al. (1947) to spoilage of salted corn by a non-sporing rod, which required at least 15% NaCl in the medium and grew in a chopped liver medium under a petrolatum seal.

174

N. E. GIBBONS

These organisms seemed to require reduced oxygen tension at least. They do not appear to have been studied further.

C. Media Once the salt requirement of these organisms was recognized, several media were devised. The high concentration of salt produced difficulties with some grades of gelatin and agar and other solidifying agents were used, e.g., starch and silica gel. With a good grade of agar, no difficulty should be experienced. At salt concentrations of 20% or more only halophiles grow and if uncontaminated salt is used an unheated medium is suitable for some purposes. This eliminates some of the difficulty commonly encountered in preparing media containing ingredients such as milk. However, for pure culture work and most bacteriological studies a sterilized medium is required. Excellent results can be obtained using fish broths. Lochhead (1934) devised a skimmed-milk medium, which gives good growth and provides a pure white background for the red colonies. The milk medium was modified by Dussault and Lachance (1952) to include magnesium, iron and glycerol-

Milk medium of Dussault and Lachance (1952) MgS04.7H20 M g ( N 0 3 ) ~6H2O . FeC13.7HaQ Proteose peptone No. 3 (Difco) Glycerol (chemically pure) NaCl (chemically pure) Distilled water

5g 1g 0.025 g 5g 10 g 200 g 1litre

For an unsterihed medium, divide into two equal parts, to one add 50 g of skimmed-milk powder, to the other 30 g agar. Heat the second to melt the agar then cool to around 60'-70°C and add the milk mixture which has been warmed to pouring temperature, mix and pour. For a sterilized medium, dissolve the ingredients listed (less the salt) in 500 ml of water, add 30 g of agar and sterilize. Sterilize the slightly dampened salt and 50 g of skimmed-milk powder dissolved in 5 0 0 ml water in separate vessels. Add the hot agar solution to the salt and dissolve as much of the salt as possible. Add the milk, mix to dissolve the remaining salt, and pour plates or distribute to tubes aseptically.

T o avoid the difficulties of making protein-containing media, hydrolysates were used. Webber (1949) devised a complex salt mixture, plus arginine, glutamate, succinate and gelatin hydrolysate (Table I). Katznelson and Lochhead (1952) used a similar medium in their study of growth factors,

175

VIII. REQUIREMENTS OF 14ALOPIIII.IC BACTERIA

' TABLE I Composition of some complex media used for growing extreme halophiles.

Components

Casamino-acids VF Casamino-acidst Yeast extractt Proteose peptonet Tryptonef Gelatin hydrolysate Arginine Cysteine Tryptophane Na citrate Nasuccinate NaHglutamate Glutamic acid KsHP04 KHaP04 KNOs KCl MgSOi.7Hz0 MgCla. 6HgO NaNOa FeS04.7Hzo FeCla NHnCl &Cia. 2H20 NaCl Other salts pHfinaladjustment

?&? &Katznelson Lockhead (1952)

.. ..

1.5

..

.. .. ..

..

..

0.1

..

0: 01 0.01

0:06 0.2 0.3

0.05

..

0.1

0105

,.

0.05 0.01

..

2:0

510 2% 0.002

+

..

.. .. ..

..

..

..

0:j

.. 0:is

.. ..

0:i 2.5 *.

0:000s

.. ..

22L26

22

.. .. .. .. ..

(1960)

0.5' 1.0

Sehgal& Dundas Eimhjellen Gibbons ct 01. (1960) (1963) (1965)

0:;s 1.0

..

.. .. ..

..

..

0.5

..

0.3.

0:3

.. .. ..

.. ..

..

..

*.

20-30

3 Oxoid

7.0

7-2-7.4

..

.. .. .. .. .. .. ..

.. .. ..

.. .. ..

..

.. .. .. ..

0:i

015

..

2.0

*.

..

0.005

..

.. 0:s

..

..

0.2' 2.0

.. 1:0

..

..

0:0023

..

25

..

4-

6.5-6.8 6-5-k.8

Expressed as g/lOO ml. t Difco

1:s

0.2

..

22%

(1955)

..

o:i

.. ..

Brown & Abram & Gibbons Gibbons

..

0:s

.. ..

015

..

25

..

.. .. .. .. .. 210

..

.. .. .*

0:s

25§

7-4

4 Solar salt

They substituted vitamin-free casein hydrolysate for gelatin and made a few minor changes in the salts. The red pigmented extreme halophiles did not respond to vitamins, nucleotides, etc., but were stimulated by yeast extract.

In this laboratory a simpler medium has been used for routine production of cells, several slight modifications being made as more information was collected (Table I). All versions allow good growth in aerated or shake cultures when incubated at 3 7 ' 4 ° C . Most of these media are adjusted to a pH of about 7.5, heated to boiling or autoclaved 5 min, filtered and readjusted to the pH shown. Some phosphate is lost, probably as ammonium magnesium phosphate, and clarification is necessary if growth is to be measured by optical methods. Laboratory-grade salts may be used if desired, but for routine culture of cells a good commercial grade, such as used in dairies, issuitable and cheaper. Eimhjellen used solar salt, which is dissolved in tap water, boiled and filtered to remove solid debris. The other salts (+ agar if required) are then

176

N. E. GIBBONS

added and the whole sterilized. A solution containing 10% each of tryptone and yeast extract is autoclaved separately and the required amount added aseptically to the mineral medium (Table I). Eimhjellen usually uses Trapani salt, which contains sufficient magnesium and calcium so that further addition of these ions is unnecessary. Studies have also been under way on a synthetic medium. For synthetic media all salts should be of analytical grade and all ingredients as pure as possible. Dundas et al. (1963) proposed the following (g/lOO ml; all salts analytical grade)NaCl KGl MgSOl(anhyd.) MgClz .6H20 CaC12.2HzO NH4Cl KzHPO4 FeC13.6HzO Cytidylic acid

24 0.5 0.5 0* 5 0.01 0.5 0.5 0 *0005 0 020

L-Lysine L-Arginine L-Proline t L-Valine t L-Methionine tL-Isoleucine (salt free) tL-Leucine (purified) L-Tyrosine L-Phenylalanine L-Glutamine

0.025 0.050 0.025 0.025 0.010 0.025 0.025 0.010 0.005 1.5

-f Essential amino-acids.

This medium supported growth of several strains of H. salinarium, H. cutirubrum, and H . halobium, although not as well as the complex medium (Dundas et al., 1963; Table I). It was less satisfactory for the coccal forms. A synthetic medium, suggested by Onishi et al. (1965), is based on the amino-acids present in casein plus the salts found necessary in more complex media. Its formula is as follows (all figures are per 100 ml and mg unless otherwise noted)1 5 Amino-acids DL- Alanine

L- Arginine

L-Cysteine L-Glutamic acid (or DL-aspartic acid Glycine nL- Isoleucine L-Leucine L-Lysine 2 Nucleotides Adenylic acid Salts NaCl MgS04.7HzO NH4Cl KN03 KzHP04 KHzP04 (see below)

43 40 5 130 45) 6 44 80

DL-Methionine DL-Phenylalanine L-Proline DL-Serine DL-Threonine (see below) L-Tyrosine DL-Valine

37 26 5 61 50

20 100

85 10

25 g 2g 0.5 g 10 5 5

Uridylic acid

10

50 0.03 0.7 0.044 0.23 0.1 g

VIII. REQUIREMENTS OF HALOPHILIC BACTERIA

177

Stock solutions of each amino-acid and of the nucleotides are prepared and combined as required. T h e minor elements are also prepared as a stock solution. The final medium is adjusted to pH 6.2 with KOH and the volume adjusted to 100 ml. It can be sterilized at 120°C for 20 min without affecting itsgrowth-promoting properties. Dundas et al. used relatively large amounts of glutamine in their medium. We found the ammonium ion as useful and used it in place of the amides. Subsequent work (Onishi and Gibbons, 1965) indicated that the ammonium ion was possibly replacing threonine and as a result it was suggested the threonine content of the above medium be reduced from 50 to 5 mg per 100ml. The amount of potassium in the Onishi et al. medium (76 ppm) is considerably lower than in the complex medium of Sehgal and Gibbons (a.1000 pprn). In this laboratory the medium was neutralized with KOH, which adds 100-120 ppm K. We have now found that there is little growth if the medium is neutralized with NaOH, and that even when neutralized with K O H the potassium is probably limiting. M. Gochnauer (personal communication) has found that much better growth is obtained by adding 0.2-0.5 g of KCI per 100 ml (ca. 1000-2500 ppm K). In the Onishi et al. medium modified in this way, she finds that in the presence of glycerol or lactate or pyruvate, vitamins have little if any effecton growth. However, if glucose is used, vitamins are stimulatory and the final pH is 7.5, rather than the usual 8-3-8.5. I t is assumed that some glucose may be utilized under these conditions, although glucose and other sugars are utilized little, if at all, in complex organic media. The amount of iron in the synthetic medium is about a tenth of that added in the organic medium. T h e phosphate requirements also need critical examination. Citrate has been added to minimize losses, but the loss due to sterilization and p H changes during growth in the presence of large amounts of magnesium and calcium is not known. T h e above media are not ideal, and further work is needed before all of the requirements of these organisms are elucidated. Further work is also needed on the relation of absorbance to numbers of colony-forming organisms and changes in cell morphology (see discussion in Section E).

D. Isolation and enumeration Red halophiles have been found throughout the world wherever heavily salted products are prepared or used. Only a few species occur commonly. The organisms under consideration in this Section are those usually found on salted proteinaceous materials and solar salts. For isolation, one of the complex media solidified with agar is usually used. The material is streaked or distributed on the surface. Incubation may be at 37"or 40°C with protection

178

N. E. GIBBONS

against excessive drying; plastic bags are very useful. ’I’he organisms are often embedded in the salt crystals and grinding of coarse salt may be helpful. Longer incubation is usually required with crystals than with cultures, brines or contaminated products. It may be well to point out here that in any work on the extreme halophiles one must think “salt” at all times. Technicians have been known to omit the salt from media, dilute the organisms in water or normal saline, etc., with disastrous results. It must be stressed that the red rod-shaped extreme halophiles will not survive exposure to salt concentrations of less than 15% even for short periods. T h e coccus forms are more resistant to changes in salt concentration, but they have not been studied sufficiently to assess adequately the effects of exposure to water and low concentrations of salt. For quantitative work a weighed sample of the salt or other material is dissolved or shaken in a sterile 15% salt solution. After suitable dilution in 15 or 20% NaCl solution, viable organisms may be estimated by the dropplate technique. Dussault (1954) described the technique for these organisms and showed that reproducible results can be obtained. We have used the method for many years. As with qualitative work the plates are incubated at 37°C in plastic bags. T h e time required will vary and must be determined for the particular study. Estimates on growing cultures may often be obtained in 3 4 days; some strains newly isolated from salts may require 10-14 days to form colonies. T h e most probable number method may also be used (see Postgate, this Series, Volume 1). Because of the aerobic nature of the organisms, incubation is in flasks on a shaker. T h e enzymes of these organisms are also inactivated in low salt concentrations. Sudden exposure to water or buffers inactivates the enzymes irreversibly. Holmes and Halvorson (1965) demonstrated that if the salt concentration was reduced gradually, the malic dehydrogenase of H. salinarium could be purified at low salt concentrations. If the salt concentration was then restored gradually, some 50 to 70% of the activity was retained depending on the amount of salt present during purification and the time the enzyme was exposed to the low concentration.

E. Large-scale production Kushner (1966) has described a system for producing large quantities of cells that was in use here for some time. A 25 gallon polyethylene tank is filled 2/3 full (ca. 70 litres) with one of the culture media described and inoculated with 1 litre of a 3 day old culture. I t is equipped with a glassenclosed heater and thermostat to provide a temperature of 37°C. Glass and plastic are used throughout, as the salt solutions are corrosive to most metals; stainless steel may be used. Mixing and aeration are accomplished

VIII. REQUIREMENTS OF HALOPHILIC BACTERIA

179

with a sparger at the bottom of the tank and an air flow of about 1.5 litres per min. T h e sparger must be cleaned at intervals as it becomes plugged with salt crystals and bacteria. There is considerable foaming, which can be controlled by an antifoam agent, such as Dow-Corning Antifoam compound A. If care is taken the medium need not be sterilized and cultures do not become contaminated in the 3-5 days required for maximum yield of cells. After use, the jar and equipment are cleaned and residual halophiles killed by thorough washing with water. As these cultures generate an offensive odour the apparatus is best installed in a fume hood. Currently, 100 litres of media (Sehgal and Gibbons) are incubated in a 150 litre stainless-steel fermenter at 37°C with aeration at 2-2.5 cu. ft/min through a sparger, with agitation at 150-250 rev/min. Under these conditions and with an inoculum of 5%, yields of 5-6 g wet weight of cells per litre are obtained in 65 h. In the course of work employing 150 and 7.5 litre fermenters, S. M. Martin (unpublished results) has noted that although the absorbance (&60) of cultures of H . cufirubrurn correlates well with the total cell nitrogen and the cell mass, it is not correlated with the number of colony-forming cells, as determined by either drop-plate counts or most probable number counts in liquid medium. T h e mean doubling time of the cell mass (as measured by absorbance) is approximately twice that of the viable cells. T h e cells are in the log phase of division well before there is an appreciable increase in the absorbance of the culture. Microscopic observations suggest that during growth, in stirred fermenters at least, the cells undergo gross morphological changes. T h e finding (S. T. Bayley, personal communication) that mitochondrial activity is greatest in cells taken about the time the absorbance begins to increase (about mid log phase on the basis of viable cell counts) suggests that, although the media and conditions outlined produce cells in relatively large quantities, there is still much to be learned about the growth patterns and requirements of these organisms.

F. Maintenance Our collection of extreme halophiles is maintained on agar slants of the Sehgal and Gibbons media in screw-capped tubes held at 5°C. With transfers at intervals (ca. 4-5 months) it has been kept with little loss and no noticeable change for some 10 years. The organisms have so far not been freeze-dried successfully. However, the organisms do survive drying for long periods in and on relatively d r y salt. Also, organisms can usually be recovered from the crystals of salt in dried-out cultures. On learning that a fish curer, who always used salt at least 3 years old, never had trouble with reddening, Dussault (1953) stored solar salt con-

180

N. E. GII313ONS

taining ca. 106 organisms/g a t 37‘ and 22 C for 2 months and found a 99% reduction a t both temperatures. The moisture content fell from 4.15% to 1.5 % during that time. Eimhjellen (1965) indicates colonies may be obtained from solar salt stored for 2-5 years, and we have isolated organisms from salt stored several years in the laboratory. T h e reduction noted by Dussault is comparable to that often found in freeze-drying of non-halophiles, and it is possible that the isolations from old salt are from a small percentage of survivors. Dussault (1957) also showed that the number of red halophiles may increase on salt stored at relative humidities of 75% and over. However, sufficient quantitative work has not been done to warrant definite conclusions 111. MOIIEKAI‘E HALOPHILES

A. Occurrence The moderate halophiles are arbitrarily taken as bacteria that will not grow without added salt, the optimum salt concentration for growth being between 3 and 5%. Brines range from the weak 2-30/, solutions produced on sauerkraut to the saturated solutions used for meat and fish. T h e weaker brines are usually used for plant materials and their main flora is usually halotolerant rather than halophilic. Only rarely is mention of halophiles made in the literature on pickle-making and vegetable salting (Etchells et al., 1947). T h e organisms considered here are some of the more extensively studied types from fish and meat brines. Papers presented at a Symposium held in 1957 (Microbiology of Fish and Meat Curing Brines, HMSO, London, 1958) dealt with the organisms in brines, their source, their properties and requirements, and whether or not they were necessary in the curing process. Since that time a few studies have been made on some specific groups or types, but the information on the moderate groups is not as extensive as on the more “glamorous” extreme halophiles. Organisms which meet this definition have been studied extensively in Japan. Many papers have appeared on the physiology of Pseudomonas strain No. 101 and on Micrococcus sp. No. 203. Although these organisms are referred to by various authors as halotolerant and halophilic they are usually grown at 10% salt concentration and are probably moderate halophiles. T h e same applies to the organisms from soy sauce mash, although these include anaerobic, halotolerant and moderately halophilic bacteria (Ueno and Omato, 1961).

B. Ionic and growth requirements Some of the moderate halophiles are NaCl specific. Micrococcus halodentri$cans, isolated from Wiltshire bacon-curing brine, does not grow at

salt cctnccntrations of 1.7.S':,, or lower, grows optimally in 4-57;, salt, but grows well but a t a reduced rate up to 23'%,salt. It will not grow in other salts such as NaHr, NaNO3, 1,iCI and KCl although it will survive in their presence for varying periods depending on the salt used and its concentration (Robinson and Gibbons, 1952). It has been suggested that this organism is a halophilic variety of M. denitriJicans (Kocur and Martinec, 1962). In contrast, another organism isolated by Robinson (1950) from the same source and now known as Vibrio costicolust grows in various salts (Flannery et al., 1952). In NaCl it requires a concentration of at least 2.5%) to grow; the optimum is around 7-8'%,, the maximum around 20'1/,. With sodium sulphate the minimum, optimum and maximum areabout 1.4,8.7 and 17.50/, respectively. Similar results were obtained with sodium molybdate, sodium phosphate and sodium bromide. No growth occurred with sodium iodide, potassium nitrate or sodium nitrate, although these salts allowed growth provided l*2yoNaCl was present. Substituting magnesium, potassium and lithium for sodium allowed some growth, but apparently the cation present ismore important than the anion.

C. Media M . halodentrij5cans and V. costicolus grow in usual laboratory media plus salt. We use 0.5% each of proteose peptone and tryptone (Difco) + 3.5-4°/0 NaCl. Flannery et a2 (1952) suggested 0.5yoyeast extract, 1% trypticase and 5.8% NaCl for V. costicolus. Incubation at 2530°C is satisfactory. The media used by Japanese workers vary from a simple 1% peptone broth for Bacterium No. 101 (Shiio et al., 1956) to a peptone, meat extract, K N 0 3 (1%, each) -t MgSO4.7HzO (0.02%) (Kono and Taniguchi, 1960) for Micrococcus 203, to a complex mixture of polypeptone (1%,), meat extract (lye), yeast extract (0.4%), liver extract (5%), soy sauce (lfY%,), rice Roji extract (2WX) used by Ueno and Omata (1961) for soy sauce organisms. All media contained 10%)NaCl. I cio not know of ary synthetic media that have been developed specifically from these organisms and considerably more work is required on their growth requirement and nutrition. We have noticed, for instance, that M. halodentrij'icans did not denitrify unless magnesium was present in small amounts.

t

I t might be mentioned that this organism was not designated V. costicojtts in these laboratorics. Robinson (1950) indicated it resembled V . costicolrrs described by Smith (1938), a culture of which was not preserved. According t o the test of Shewan c't (11. (1954) (Vibriostatic agent 0/129), it is not a vihrio. Also, the specific name should hc rostico/u.rather than c,rtiroltts, so both thc taxonomy ;Ind nomenclature is in doiil>t. Similar ciirvcd org;inisms h;i\.c hcen follntl in rntaat-curiny hrincs(1 Icnryc~tol., 19.57; Iiutti;iilx, 1057).

D. Pathogenic halopkiles Ily definition, i t is doiil)tful i f iiny 1 i i i i n ; i n o r i i n i i i i ; i l ~xitli~igcn i s a true ha1ol)hilc. I Iowc\~cr,some h;ivc I~ecndcscriI)cti. A micrococcus thiit grcw poorly in ordinary media, but wcll in media containing 3--S(,’{)salt wits isolated from infected lesions on the fingers of seal hiintcrs (‘l’hj6tta anti Knittingcn, 1949). ‘l’his is probably a marine organism that can grow in human serum. In Japan, several outbreaks of food poisoning have been attributed to halophiles. Various salted products have been implicated, principally sea fish. Although the organism was first isolated on blood agar, its later isolasalt agar, under the assumption that staphylococci were involved, tion on 4(;{’ resulted in its “halophilic” character being stressed. How organisms meeting the definition of halophiles could multiply in the human intestine was not clear. Sakazaki et nl. (1963) have reviewed the whole question and, after testing numerous strains, identified the causal organism as Vibrio parahaemol3~ticz~s. ‘rhey statc that the organism grows well in brain heart infusion broth or on t h o c l agar without added salt, but poorly on peptone water or on plain agar with ordinary salt concentration (presumably 0.5%). This raises the whole question of halophilism in moderate halophiles, or at lcast in some organisms that h w e been placed in this category. In my opinion, sufficient is known to place the extreme halophilic rods in a separate category, quite apart from other bacteria. T h e position of the moderate halophiles is still in doubt. More information is needed on their nutrition and on thcir physiology in relation to their ionic requirements. Further information on their chemistry and biochemistry may reveal whether they have unique properties similar to the extreme halophiles. REFERENCES Abram, D., and Gibbons, N. E. (1960). Can. J . Microbid., 6, 536--543. Baas-Uecking, L. G. M.(1931). Scient. Mon., 32, 434-446. Baumgartner, J. C . (1937). Fd. Res., 2, 321-329. Block, M. R. (1963). Scient. Am., 209, 89-98. Boring, J., Kushner, D. J., and Gibbons, N. E. (1963). Can. J . Microbid., 9, 143154. Brown, PI. J., an d Gibbons, N. E. (1955). Cun.J. bi‘icrobiol., 1,486--494. Buttiaux, 11. (1 957). In “The Microbiology of Fish and hleat-Curing Brines”, pp. 137-148. Pi.,lI. Stationery Oflice., Idondon. Cho, K. Y . , Doy, C. H., and Mercer, 1:. 13. (1967).,T Uact., 94, 196-201. Christian, J . €3. 13. (1956). D.Phi1. Thesis, University of C‘amhridgc.. Dundas, I. D., and Larsen, 1-1. (1962). Arch. Microbird., 44, 233 -239. Dundas, I. I)., and Larsen, 11. (1963). Arch. Mic.robio/.,46, 19--22. Dundas, I . I)., Sribivasan, V. It., and Ilnl\orson, l I. 0 . (19hB). (:an. J. iWicvobio[., 9,619-624. I)ussault, 1 I . 1’. (19S3). /’MI,y. /<(,/I. :?t/ant. (,’.st. s t . , No. 55, 7-9.

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Dussault, 11.1’. (19S4)..7. Fi.sft.R e s . Ud.Cm.,11,261-266. Dussault, €1. P. (1957). In “1Llicrobiology of Fish and Meat-Curing Brines”, pp. 13-19. H.M. Stationery Office London. Dussault, E-I. P., and Lachance, R. A. (1952).J. Fish. Res. Bd. Can., 9, 157-163. Eimhjellen, I(. (1965). In “Anreicherungskultur und Mutantenauslese”. Zentbl. Bakt. Parasitkde, Abt. 1., Suppl., 1, 126-1 38. Etchells, J. L. Jones, I. D., and Lewis, W. M. (1947). U S D A Tech. Bull., No. 947, 1-64. Farlow, W. G. (1880). U S . Fish C o ~ mR. e p . for 1878, p. 969. Flannery, W. L., Doetsch, R. N., and Hansen, P. A. (1952).J. Bact., 64,713-717. Gibbons, N. E., and Payne, J. I. (1961) Can.J. Microbiol., 7,483489. Harrison, F. C., and Kennedy, M. E. (1922). Trans. R. Soc. Can., 3rd Ser., 16, 101-1 52. Henry, M., Joubert, L., Renault, I,., and Goret, P. (1957). In “Microbiology of Fish and Meat-Curing Brines”, pp. 21 3-222. H.M. Stationery Office. London. Holmes, P. K., and Halvorson, H. 0. (1965). J. Bact., 90,312-3 15, 3 16-326. Hornsey, H. C., and Mallows, J. €1. (1954). J. Sci. Fd. Agric., 5 , 573-583. Hornsey, H. C., and Mallows, J. H. (1955). J . Sci. Fd Agric., 6, 705-712, 712-715. Hoye, Kr. (1906). Hergens Mus. Arb., 1906, No. 12,64 pp. Ingram, M. (1957). In “Microbial Ecology” (Ed. R. E. 0.Williamsand C. C. Spicer), pp. 93-1 33. university Press, Chmbridge. Ingram, M., Kitchell, A. G., and Ingram, G. C. (1957). In “1LZicrobiology of Fish and Meat-Curing Brines”, pp. 205-212. H. M. Stationery Office, London. Katznelson, H., and L,ochhead, A. G. (1952). J. Bact., 64, 97-103. Kellerman, K. F. (1 915). ZentbZ. Bakt. Parisitkde, Abt. II,42, 398-402. Klebahn, H. (1919). Mitt. Inst. ailg. Bot. Hamb., 4, 11-69. Kocur, M. and Martinec, T. (1962). Folia prirodovedeckefak. Univ.J. E. Purkyne v Brne, 3(3), 1-1 21. Kono, M., and Taniguchi, S. (1960). Uiochim. biophys. Actu., 43,419-430. Kushner, D. J. (1966). Blotech. Bioengng., 8 , 237-245. Larsen, H. (1962). In “The Bacteria” (Ed. I. C. Gunsalus and R. Y. Stanier), Vol. 4, pp. 297-342. Academic Press, New York. Larsen, H. (1967). Adz!. microb. Physiol., 1, 97-132. Lochhead, A. G . (1934). Can.J. Nes., 10, 275-286. MacLeod, R. A., and Onofrey, E. (1957).J. cell. comp. Physiol., 50, 389401. Onishi, H., and Gibbons, N. E. (1965). Cun.J. Microbiol., 11, 1032-1033. Onishi, H., McCance, M. E., and Gibbons, N. E. (1965). Can. J. Microbiol., 11, 365-3 73. Robinson, J. (1950). Ph.l). Thesis, McGill University, 92 pp. Robinson, J., and Gibbons, N. E. (1952). Can.J. Rot., 30, 147-154. Sakazaki, R., Iwanami, S.,andFukumi, H. (1963).Ja~..7.nzcd.Sci. fliol.,16,161-188. Sehgal, S. N., and Gibbons, N. E. (1960). Can. J . Microbiol., 6, 165 -169. Shewan, J. M., Hodgkins, and Idiston, J. (19.54). Nature, b n d . , 173, 208. Shiio, I., Maruo, B., and Akabori, S. (1956). J. fliochenr.43(6), 779-784. Smith, F. B. (1938). Proc. A. Soc. Qd., 49(3), 29-52. Stoeckenius, W., and Rowen, R. (1967).J. Cell Uiol., 34, 365-393. Thjotta, Th., and KnittinZen, J. (1949). Acta path. microbiol. scund., 26, 4117-41 1. Ueno, T. and Omata, S. (1961).J. Ferment. Techno/.,Osaka, 39, 52-60, 360-370. Volcani, R. E. (1944). In “Papers Collected to Commemorate the 70th Anniversary of Dr Chaim Weizmann”, pp. 71-85. Collective Volume, Daniel Sieff Research Institute, Rehovoth, Israel. Wehher, RS. hS. (1949). Biol. Rev. Cy Cdl. N. Y., 11, 9--14.

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CHAPTER I X

Isolation, Cultivation and Maintenance of the Myxobacter ia JOHN

E. PETERSON

Department of Botany, University of Missouri, Columbia, Missouri, U.S.A. I. Introduction: The Myxobacteria

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11. The Isolation of Myxobacteria . A. Soil as a source of myxobacteria B. The bark of living trees as a source C. Dung and plant-debris sources D. Sources of cellulolytic forms E. Sources of bacteriolytic forms . F. Parasitic forms G. Aquatic sources

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111. The Cultivation of Myxobacteria . A. Media . B. Purificatioq . C. Culture conditions . D. Liquid culture . E. Problems encountered during cultivation

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IV. The Maintenance of Myxobacterial Cultures . A. Maintenance as living cultures . B. Maintenance by freeze drying . C. Maintenance by freezing . D. Maintenance of desiccated resistant structures References

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I. INTRODUCTION: T H E MYXOBACTERIA ’The myxobacteria generally cannot be handled successfully in the same fashion as the true bacteria. Further, though they are a small group of organisms, there is considerable diversity within the order; not even the same techniques of isolation and culture can be applied to all the myxobacteria. For these reasons, the methods necessary for handling the myxobacteria are worthy of special attention, although, in general, they are neither particularly unique nor exotic. The most recent comprehensive treatment of the myxobacteria is that found in “Bergey’s Manual of Determinative Bacteriology” (Breed et al.,

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1957). T h e Order Myxobacterales, as recognized in “Bergey’s Manual”, consists of five families, the Cytophagaceae, the Myxococcaceae, the Archangiaceae, the Sorangiaceae and the Polyangiaceae. T h e members of the monogeneric family Cytophagaceae and the three recognized species of the genus Sporocytophaga of the family Myxococcaceae do not produce fruiting bodies; all the other myxobacteria do. It is now quite clear that these non-fruiting forms are not myxobacteria. They are more closely related to the flexibacteria than to the myxobacteria, and the methods of handling them are predominately those applicable to the true bacteria; they will not be included in this consideration of the myxobacteria. The evidence and rationale for excluding the genera Cytophaga and Sporocytophaga, as well as that for deviating somewhat from the systematic pattern found in “Bergey’s Manual”, will not be detailed here. T h e brief summary of the major faxa presented is my concept of the order, based on 15 years of work with these organisms and, generally, agrees with views of others in this field. This summary is presented only as a framework for a discussion of the methods of isolation, cultivation and maintenance of the myxobacteria. Three characteristics are common to all members of the myxobacteria as recognized here(1) They produce copious amounts of a polysaccharide slime and, hence, a distinctive, slimy colony. (2) They all possess vegetative cells that glide on solid surfaces, although the mechanism of movement has not bPen elucidated as yet. (3) They produce resistant fruiting bodies, which are aggregates of many shortened cells bounded by a slime sheath. “Bergey’s Manual” recognizes 72 species in 11 genera in five families in the order. It is my opinion that less than 50 species in nine genera in four families should be recognized. The members of the family Myxococcaceae possess the proportionately long, flexuous, Gram-negative vegetative cells with pointed or tapered FIG.1. Vegetative cells of the long, flexuous, tapered type typical of the Myxococcaceae, Archangiaceae, and Polyangiaceae. FIG.2. Vegetative cells of the short rigid, blunt type typical of the Sorangiaceae. FIG.3. Myxococcusfulvus. Habit of fruiting bodies. FIG.4.Myxococcusfztlzvrs. Whole mount of a single fruiting body. FIG. 5 . Archangium primigenium. Habit of fruiting bodies. FIG.6. Archangium primigmium. Whole mount of a single fruiting body showing the tubules, which contain the shortened cells, embedded in slime. FIG.7. Sorangium cellulosum. Habit of a portion of a colony showing many fruiting bodies. The dark masses are mature fruiting bodies in the centre of the colony; the light masses are immature fruiting bodies on the periphery. FIG.8. Sorangirtm cellulosum. Whole mount of two fruiting bodies.

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ends, which are considered typical of the myxobacteria (Fig. 1). These cells shorten to become spherical microcysts in the resistant stage ; hence the family name. Great numbers of microcysts are produced in simple fruiting bodies, which take the form of cushions, globes and unbranched or sparsely branched columns. There are two genera in the family: Myxococcus in which the bounding slime of the fruiting body is deliquescent (Figs. 3 and 4), and Chondrococcus, in which the slime sheath bounding the fruiting body is extremely persistent and apparent. T h e members of the family Myxococcaceae are commonly found in soil, on various animal dungs and on the bark of living trees; they have been found less commonly in fresh waters, and one of them is parasitic on both freshwater and marine fishes. Some of them are strongly eubacteriolytic and, in addition to lytic enzymes, have been shown to produce antibiotic substances. None of them are cellulolytic. Numerous species and varieties have been described in the two genera, and it is impossible at present to state how many valid members the family contains. Five species appear to be valid, with possibly two or three others worthy of recognition. Two genera, Stelangium and Arclzangium, comprise the family Archangiaceae. The vegetative cells cannot be distinguished from those of the Myxococcaceae, but they do not shorten to become coccoid in the fruiting bodies; they do shorten appreciably, but they always retain the form of a rod. These shortened rods are encased in a simple cushion- or columnar-shaped fruiting body in the genus Stelangium and in irregular slime tubules, which are in turn embedded in slime, in the genus Archangium (Figs. 5 and 6). T h e members of the family are known from soil, from dungs, from plant debris and from the bark of living trees. None of them is parasitic, and none of them has been shown to be either cellulolytic or eubacteriolytic, although one might reasonably expect that they are both. There are probably a total of five valid species in the two genera. The family Sorangiaceae includes only one genus, Sorangium, with probably only three species,. although eight are recognized in “Bergey’s Manual”. The critical character of the genus is that the vegetative cells are quite unlike those found in any of the other myxobacteria. They are proportionately short, generally rigid cells with rounded ends (Fig 2). Once FIG.9. Polaangium vitellinum. Habit of a moist, nearly mature fruiting body. FIG. 10. Polyangium e~itellinum. Habit of a mature, desiccated fruiting body. FIG.1 1 . Polyangium vitellinum. Whole mount of a single fruiting body. FIG.12. Podangium gracilipes. Habit of several fruiting bodies on a moss plant. FIG. 13. Chondromyces crocatus. Habit of a fruiting body. FIG.14. Chondromyces crocatus. Whole mount of a single fruiting body. FIG. 15. Chondromyces apiculatus. Habit of fruiting bodies on a piece of bark. FIG.16. Chondromyces catenulatus. Habit of a single fruiting body on a piece of bark.

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one has been acquainted with the two types of myxobacterial vegetative cells, there is no possibility of ever failing to differentiate between them. A few to many of these cells aggregate into rounded cysts with slime walls, several of which are in turn bounded by a fruiting body wall of slime (Figs. 7 and 8). T h e encysted rods shorten very little, as compared to the vegetative rods. These organisms are predominately soil forms. Many isolates are strongly cellulolytic, although some do not possess this property. T h e family Polyangiaceae contains the largest number of recognizable species of the myxobacteria, and also those species that form the most complex, intricate fruiting bodies. Some members of the family possess vegetative cells that are flexuous with tapered ends similar to those in the Myxococcaceae and Archangiaceae ; they are proportionately long in relation to width, but they are somewhat shorter and thicker than those of the first two families. Other members of the family possess cells that resemble the Sorangium-type cell, although they are proportionately longer and more flexuous. I n the resistant state, the vegetative cells shorten proportionately less than do those of any of the other families; they are always distinct, relatively long rods. Four genera are readily recognizable on the basis of fruiting-body morphology. I n the first of these genera, Haploangium, the encysted rods are contained in a single cyst with a wall composed of slime. There appear to be four valid species of Haploangium. T h e members of the genus Polyangium produce fruiting bodies, which are multiples of cysts bound with a common slime envelope. These fruiting bodies are morphologically similar to those of the Sorangiaceae, although they are usually larger (Figs. 9, 10 and 11); of course, they contain quite a different type of encysted rod. “Bergey’s Manual” includes fifteen species and four varieties of Polyangium, but probably only three or four species should be recognized as valid. T h e third genus of the family Polyangiaceae, Podangium, is characterized by the production of fruiting bodies that are solitary cysts on a slime stalk (Fig. 12). Five species seem to merit recognition. T h e genus Chondromyces contains some 5-1 0 species of myxobacteria that produce intricate, tree-like fruiting bodies. The fruiting bodies are composed of central stalks of slime, which branch in varying degrees, with the cysts, in which the rods are contained, produced on the ends of the branches in various fashions. These fruiting bodies are truly architectural marvels, particularly when they have been produced by the collective ingenuity of several hundred individual cells! (Figs. 13-16). T h e members of the Polyangiaceae may be isolated with reasonable frequency from various soils, but they occur with amazing frequency on dungs, on decaying plant debris and, even more so, on the bark of living

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trees. None of them has been shown to be cellulolytic, although it would appear that some of them ought to possess this characteristic, at least adaptively. Some of them certainly are eubacteriolytic, although this character has not been studied in detail for any of them. Two species have been reported to be parasitic, one on lichens, the other on an alga; neither of them has really been studied critically by other than the original observer. With one or two exceptions, the members of the Polyangiaceae are almost entirely unstudied, even though they produce the largest, the most spectacular, and the most commonly occurring fruiting bodies of any of the myxobacteria. This is, indeed, unfortunate, since they would be excellent tools for morphogenetic and other studies. Perhaps we will soon learn to handle them more easily in culture. 11. T H E ISOLATION OF MYXOBACTERIA The myxobacteria are common organisms. They occur in large numbers in a wide variety of habitats. They are cosmopolitan, having been reported from the Arctic, Swedish Lapland, Russia and Japan in the north to near the equator on the south, and from the continents of America, Europe and Asia. They undoubtedly will be found south of the equator and in other areas of the northern hemisphere when workers search for them in these locales. Yet, the myxobacteria are not often observed, and most microbiologists have never seen a single representative of the order. There are three reasons, in particular, for this seeming rarity of occurrence. First, the myxobacteria are relatively slow growers. A minimum of 4 or 5 days usually elapses before 'myxobacterial colonies and fruiting bodies can be detected. With some forms,,2-3 weeks are necessary before the organisms become apparent. Second, even though several of the myxobacteria produce antibacterial and antifungal substances, they must be considered poor competitors in terms of the usual isolation techniques; hence, they are rapidly over-grown or, more often, never get started toward a detectable colony at all. Filamentous fungi are the primary offenders, although these competitors can often be suppressed satisfactorily by incorporating cycloheximide (Actidione; The Upjohn Company, Kalamazoo, Michigan) or other antifungal antibiotics in the isolation medium. T h e third reason for the infrequent appearance of myxobacteria is actually dependent on the first two; this is a matter of proper selection of media that will enhance the slow-growing myxobacteria while inhibiting as many of the competitors as possible. If due consideration is given to these three points, myxobacteria tan be successfully obtained in large numbers with minimal time and tffort.

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A. Soil as a source of myxobacteria Considerable numbers of Myxococcus spp. and Archangium spp., practically all of the Sorangiunz spp. and limited numbers of Polyangium spp. and Chondromyces spp. can be isolated from soil sources. T h e best technique for such isolations is the use of what may be referred to as K & K plates. Their use for myxobacteria was first reported by Krzemieniewska and Krzemieniewski (1926). T h e technique consists of filling a deep Petri dish one-third full of the soil from which one wishes to isolate, moistening the soil slightly if necessary, and placing eight or ten autoclaved rabbit-dung pellets on the soil. T h e dung pellets may be from domesticated rabbits, although those from wild rabbits are preferable; they should be autoclaved for 20-30 min at 121°C and cooled before use. They should be pushed about half-way down into the moist soil with a sterile forceps. Large supplies of pellets may be prepared in a suitable covered dish and used as needed over a period of several weeks. Myxobacteria have a strong affinity for rabbit-dung pellets. They tend to grow onto the dung from the soil, and to fruit on the pellets from where they can be picked off and transferred. Eubacteria and fungi may sometimes be a problem, but the pellets seem to be somewhat selective for myxobacteria over other groups of organisms. T h e pellets may be dipped in a solution containing 1.0mg of active cycloheximide in about 40 ml of distilled water before they are placed in the soil, but it is generally advantageous simply to discard those plates that are too overgrown. Plates are kept at room temperature and examined under a sterioscopic microscope daily from the 4th until the 7th day after preparation. Two further methods of isolation from soil use Stanier plates and Singh eubacterial-circle plates, which are described later. T h e Stanier plates are primarily selective for cellulolytic organisms, although some non-cellulolytic myxobacteria can also be isolated on them. T h e Singh eubacterialcircle plates are selective for eubacteriolytic organisms. Although a few non-eubacteriolytic forms may occur on Singh plates, these are so limited in numbers that it is profitable to use this method only if one is seeking eubacteriolytic myxobacteria.

B. The bark of living trees as a source I t is my opinion that bark of living trees is the single best source of myxobacterial species, and they occur there in large numbers. Nearly any species of myxobacterium may be found on this substrate, and this is certainly the source of choice for the more complex fruiting forms. MyxoCOCCUS spp. are quite common, but one can usually isolate more of these for a given amount of effort from soil. Chondrococcus spp., with the exception of the fish parasite, Chondrococcus columnaris, which is a special case, are very

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profuse; bark, in fact, can be considered the substrate of choice for these myxobacteria. Archangiurn spp. are very common on bark, often in conjunction with Podangium spp. Bark is'the substrate of choice for Podangium spp., some of which are known only from this substrate. T h e two species of Stelangiurn also are known only on bark. Members of the genus Sorangiurn occur very infrequently on bark and, when they do, they are invariably not cellulolytic forms. The genus Sorangium is the only major group of the myxobacteria for which bark is a poor source. All of the members of the four genera of the Polyangiaceae may be found on bark; it is the source of choice for all of them. Certainly, results will vary with the geographical location and with the type of bark selected, but it has been my experience that myxobacteria may be found on about one out of every four pieces of bark. Myxobacteria have been found in large numbers on bark of many tree species collected from a wide range of environments, ranging from isolated forests to large cities. The technique, first used by Gilbert and Martin (1933) for myxomycetes, is simple. Pieces of bark, which are in reasonably intimate contact with the tree are placed on filter paper or blotting paper in a suitable covered dish (deep Petri dishes are excellent), and soaked thoroughly with water. A piece about 1 x 2 in. in size is very satisfactory. After 24 h, excess water is poured off and the dishes are left undisturbed at room temperature in ordinary room light (i.e., diffuse daylight without artificial light). T h e bark may be examined with the stereoscopic microscope (about 15 x ) after 4 or 5 days, but one conserves time and does not disturb the developing myxobacterial fruiting bodies by waiting until 10-12 days to examine the bark. Newly formed fruiting bodies can readily be seen and easily picked-off with a needle for transfer onto a suitable medium for purification and culture.

C. Dung and plant-debris sources Many species of myxobacteria, sometimes in considerable profusion, may be developed on various animal dungs and on pieces of plant debris, preferably woody materials, placed in moist chambers. However, in terms of the numbers of myxobacteria obtained in relation to the numbers of individual isolations set up, this is an unrewarding method of obtaining them. If naturalist colleagues, who routinely examine quantities of dung and woody plant debris in the field are familiar with the appearance of myxobacteria on these materials considerable effort can be saved. There is another possibly rewarding facet to this source. The dung pellets of wild rabbits, when gathered and placed in moist chambers, frequently produce a considerable number of myxobacteria on them. I t is a simple matter to locate rabbit runs, to collect fair quantities of pellets, and to place them in moist chambers; they are often rich in myxobacteria.

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D. Sources of cellulolytic forms Soil is the only reasonable source if one wishes to isolate cellulolytic myxobacteria. The only cellulolytic forms are members of the genus Sorangium. “Bergey’s Manual” indicates that Polyangium cellulosum, as described by Imshenetski and Solntzeva (1937), and the four varieties of this species described by Mishustin (1938) are cellulolytic; I am convinced that these all really belong to the genus Sorangium and, moreover, to a single species of that genus. The type of soil selected for isolation does not appear to be important other than that cultivated soils will yield the greatest numbers of myxobacteria. All cultivated soils that have been examined seem to contain these myxobacteria in reasonable numbers. In addition, they may be found in many non-cultivated soils. They have been found in various soils in Russia and Poland (Krzemieniewska and Krzemienieski, 1926, 1927, 1937; Imshenetski and Solntzeva, 1937; Mishustin, 1938; Zhukova, 1959, 1960, 1962); I have found them in soils associated with cultivation in all four Scandinavian countries. I have found them in tremendous profusion in the Sonoran desert soils of Mexico and Arizona, both in those soils which are native and those that have been irrigated and placed under cultivation. I have found them in cotton field soils of southeastern Missouri, and my colleagues and I have found them in various soils in central United States from the Canadian border to Alabama, Louisiana and Texas. T h e most satisfactory method of isolating these organisms is with the use of the Stanier plate. This cellulolytic-selective medium was first reported by Winogradsky (1929), and later modified by Stanier (1942); hence the present designation. It is simply an agar medium containing routine salts, which is overlaid with a piece of sterile filter paper as the carbon source. T h e medium is as followsAgar medium KN03 KzHPO4 MgS04.7Hzo CaC12.2HzO FeC13.6HzO Agar Distilled water

1.og

0 - 2g 0.1g 0 - 1g 0.02 g 10.0 g 1 litre

T a p water has proved to be superior to distilled water in all geographic locations where we have used it; because of the variability of tap water, however, this may not always be so. Sterilize the agar at 121°C for 15-20 min, pour it fairly deep (because the incubation time for this isolation is long) and allow it to solidify. Aseptically place a piece of ordinary filter paper (we have generally used Whatman No. 1) that has been sterilized in the autoclave on the surface of the agar.

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The soil sample is mixed with sufficient water to make a thick, muddy paste. Volumes of soil the size of a small pea are placed on the surface of the filter paper with a small spatula; five such piles can readily be placed on each standard 9 cm dia. Stanier plate. When many dozens of such plates are to be made, it is a simple matter to construct either a bulb-operated pipette, or a syringe-type applicator, from large-bore glass tubing. We have used such instruments to plate piles of soil rapidly and in reasonably uniform volumes when the soil being sampled did not contain rocky inclusions of undue size. The plates are best incubated at 28"-30°C in a relatively humid atmosphere for a minimum of 3 weeks. The cellulolytic myxobacteria can readily be seen growing out of the piles of soil, usually with the unaided eye, although a stereoscopic microscope is useful in examining the plates. Of course, eubacteria, actinomycetes and fungi, immense numbers of nematodes, some algae, some insects and a scattering of small plants will all appear on the plates. Only the fungi present any problems, in that they sometimes cover the plates to the point where all is obliterated. Cycloheximide incorporated into the agar medium in the amount of 1 mg of active antibiotic/40 ml of medium will generally suppress the fungal growth satisfactorily;we have used it in concentrations up to 1 mg/25 ml of medium without any apparent effect on the myxobacteria growing out of the soil piles.

E. Sources of bacteriolytic forms Many myxobacteria have the ability to lyse eubacterial cells and to utilize the resultant products in their metabolism. In all probability, myxobacteria play an appreciable role in the control of eubacterial populations in soil and other such ecological systems. Soil, then, is again the most logical source from which to isolate eubacteriolytic myxobacteria, although they can also be recovered from decaying plant debris, from animal dungs and from the bark of living trees. The most commonly occurring species of Myxococcus certainly are eubacteriolytic ; Chondrococcus coralloides is, some of the Polyangium spp., such as P.fuscum, are and many of the species of Chondromyces are strongly so. Other myxobacteria may also be recovered if one is using eubacteriolytic-selective methods of isolation. The simple technique developed by Singh (1947) is the method of choice. Plates of 1.5% water agar are poured and permitted to solidify. A sizeable mass of eubacterial cells is scraped from a 48-72 h slant with a loop, and spread thickly on the surface of the water agar in a circle. Three such circles can conveniently be placed on a standard plate. T h e choice of eubacterium depends primarily on which one the isolater prefers to seek activity against; any common laboratory strain is satisfactory. Singh used Aerobactm

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aerogenes (hence, the technique has also been referred to as “Aerobuctev circle plates”, or “Singh’s Aerobacter circles”), and we have found this organism, as well as many others, to be quite satisfactory. After the circles have been spread on the agar surface, a small pile of the soil is placed in the middle of each circle. Those organisms, a sizeable proportion of which will be myxobacteria, that can utilize the eubacterial cells can be seen growing out of the soil in 2-3 days. T h e myxobacteria will begin to produce fruiting bodies in 5-7 days, which, of course, greatly facilitates the recognition and further isolation of these particular organisms. Because of the minimal nutritional nature of this particular isolation method, the over-growing of the plates by other organisms is seldom a problem. A modification of the Singh circles may be used, in that the eubacteria can be streaked heavily across the agar rather than placed in a circle. The pile of soil is placed on the centre of the streak, and the outgrowing organisms will tend to go down the streak in both directions, producing fruiting bodies as they go. We have not found that streaks are particularly advantageous over circles.

F. Parasitic forms T h e reports of Graf (1962) of myxobacteria in the human oral cavity are apparently concerned with a flexibacterial, or cytophaga-like, organism rather than a fruiting myxobacterium ; hence, the isolation of this organism will not be considered here. Geitler’s (1925) isolation of a myxobacterium that parasitizes algae is still unique, and reference to the original beautiful work should be made. Podangium lichenincolum, so called by Thaxter (1892), has been seen by us on several occasions on disintegrating lichens on the bark of living trees. Although there is no direct experimental evidence to establish this species as parasitic on the lichens, bark containing lichens is the most likely to yield Pod. lichenincolum by methods described earlier. Currently, the only bona fide parasitic rnyxobacterium appears to be Chondrococcus columnaris, which attacks both marine and freshwater fish. There is no question about the parasitic nature of this organism. Some question as to whether the organism is a genuine fruiting myxobacterium is developing, but, certainly for the present, it is desirable to continue to consider it as such. T h e early contributions on this organism and its parasitic and pathogenic nature were made by Davis (1922), Fish and Rucker (1943) and Ordal and Rucker (1944). Anyone wishing to isolate and handle C. columnaris is referred to the excellent works of Ordal and his colleagues over the past twenty years, although there is no reason to review this work here.

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The recommended agar for its isolation isAgar Tryptone Yeast extract Beef extract Sodium acetate Agar Distilled water

0.5 g 0.5 g 0.2 g

0.2 g 9.0 g 1 litre

The agar should never be used in a concentration greater than 1.1%, since the organism grows into the soft agar to form characteristic colonies. T a p water is often preferable to distilled water, but, again, this may not be true of all tap waters. Standard-grade agar as prepared by the Difco Laboratories, Detroit, Michigan, is preferable to the purified agar, which is somewhat toxic. The medium should be adjusted to a p H of 7-2-7.3. Isolations may be made from the edges of relatively small lesions, usually by making scrapings or by taking bits of tissue and planting them on the agar. After colonies have been located under the stereoscopic microscope, they may be put onto fresh plates of the same medium, except that it should contain only 0.4% agar, and be poured very deep. C. columnaris can also be isolated directly from water. It is difficult to do, however, and infected fish are much more economical sources for isolation.

G. Aquatic sources With the possible exception of C . columnaris, no truly aquatic forms of the fruiting myxobacteria are known. A few Cytophaga-like organisms have been included in studies of marine aquatic organisms; no fruiting myxobacteria, other than C. columnaris have been included. Only two studies have actually attempted to examine the myxobacteria of freshwater. One of these was done by Jeffers (1964) on a single body of water in Minnesota. T h e other was reported in an as-yet unpublished Master’s thesis from the University of Missouri by one of my students, Galen Renwick, and included examination of 24 bodies of fresh water in the midwestern United States. Fruiting myxobacteria can be isolated from fresh water. They are all regular soil and decaying piant-debris species, and they are present in such low numbers that they are probably there only in the resistant forms which were washed into the aquatic environment. Water sources certainly cannot be considered to be of importance, and they are not worthy of time and effort as an isolation source. If myxobacteria are to be isolated from water, some method of concentration of particulate matter in large volumes of water, such as a filtration technique, must be employed.

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cuImvmioN OF MYXOBACTERIA

A. Media After the initial isolation of a fruiting myxobacterium as described in the preceding Section, one must select an appropriate medium onto which the organism can be transferred, ultimately purified and grown. The procedures of purification and cultivation are, in general, relatively routine with members of the Family Myxococcaceae, but they become increasingly demanding as one deals with the more complex forms. Though sometimes difficult, one can usually succeed with suitable media and with persistence.

1. Dung-containing media Petri dishes of 1.5% water agar, into which three or four autoclaved rabbitdung pellets have been placed aseptically while the agar was molten, are useful in the early purification and cultivation process. T h e pellets may be from either domesticated or wild rabbits, providing the domestic rabbits have not been fed on antibiotic-containing, or similar, food; pellets from wild rabbits are always preferable, however, if they are available. The agar should never be so deep as to cover the top half of the pellets, and the pellets should never be tolled in the molten agar or otherwise coated with it. Each pellet should be seeded with the myxobacterium. T h e colony will usually grow onto the agar from the pellet where the colony edge can be isolated for further transfer. Dung-decoction agars are also useful in some early cultivation procedures, but they are never as useful as the pellet plates. They may be made from either rabbit dung or horse dung by steeping the dung (about 1000 g of dung/litre of water) for an hour or so, filtering through several layers of gauze and, finally, through filter paper, and then bringing back to volume before sterilizing and pouring into plates. Many of the myxobacteria will produce vegetative colonies on such decoction agars, but growth is usually not robust, and many of them will not fruit on the agar surface. Pellet plates are quite superior to docoction agars for the production of fruiting bodies.

2. Cellulose-containing media For those myxobacteria that degrade cellulose, namely certain Sorangium spp., the medium of choice for early purification and cultivation is that described earlier for the Stanier plates. We have distributed pulverized filter paper and other forms of cellulose in this same agar medium and succeeded in growing cellulolytic myxobacteria. We have added pure cotton into the same agar medium as the carbon source, and have laid it on the agar surface with some success in growing cellulolytic myxobacteria. We have

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incorporated pulverized filter paper into liquid cultures with some success. In no instance, however, have any of these modifications of a cellulosecontaining medium yielded as good a growth, fruiting, pigmentation or cellulolytic activity as is obtained on Stanier plates. Methyl cellulose, used in media as a soluble form of cellulose, has not been at all satisfactory. With some isolates, media containing cellobiose as the carbon source in lieu of cellulose has been satisfactory for both solid and liquid cultures; this will be discussed in a following paragraph.

3. Media containing other natural entities Water agars in which either autoclaved or washed pieces of bark, various plant materials and various cellulosic materials have been placed have been tried as media for myxobacterial growth. None of them have proved to be of any value. Bark extracts and soil extracts, made in various ways from several sources, have been incorporated into various media; in no instance have we found any indication of enhancement of growth.

4.Defined and nearly defined media A useful medium for cultivating a wide spectrum of myxobacterial species is that which McCurdy (1963) modified from an earlier suggestion of McDonald and Peterson (1962) as a medium primarily suitable for cultivation of species of the Family Archangiaceae. McCurdy’s medium containsMcCurdy’s medium (1963) 1.og

Raffinose Sucrose Galactose Soluble starch Casitone (Difco)

1.og

1.og 54g

2.5 g 0.5 g 0.25 g 15.0g 1 litre

MgS04.7HzO

KzHPOi Agar Distilled water Adjust the pH to 7 . 4 !

Occasionally, additions of a vitamin solution to the medium after autoclaving will improve growth, but this is usually unnecessary and it is not normally added for routine work. Some representatives of all four families of the fruiting myxobacteria have been grown on this medium, and in this medium when the agar has been omitted.

N o r h (1952) has used the following incdium to grow three species of the Family Myxococcaceae-

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Norin’s medium (1952)

2.5 g 2.5 g 2.0 g 1.2g 0.1 g 0.01 g 0.001 g 0.003 g 1 litre

Casein hydrolysate Asparagine I
I t has since been used for the succcssful cultivation of a few other species of the fruiting myxobacteria. T h e most defined medium reported for a myxobacterium is that of Dworkin (1963), which he developed for the vegetative growth of his strain xanthus. This medium is as followsof M~XOCOCCUS Dworkin’s medium (1963) Tyrosirie Asparagine Leucine Isoleucine Proline Arginine Histidine Glycine Lysine Methionine Phenylalanine Tryptophan Serine Threonine Valine Djenkolic acid Alanine Glycogen MgS04.7H20 Distilled water Make the medium 0.01 M in K2HP04-KI-IzP04

0.6g 0.5 g 1.0 g 0.5 g 0.5 g 0.1 g 0.05 g 0.05 g 0.25 g 0.05 g 0.15 g 0.05 g 0.1 g 0.1g 0.1 g 0.1 g

0.05 g 3.0 g 1.og 1 litre buffer (pH 7.6)

T h e phenylalanine, tryptophan and alanine were all used in concentrations of 1.0 g/litre for other strains of Myxococcus xanthus on occasion. Dworkin found that the omission of phenylalanine and tryptophan caused the cells to become microcysts, hence initiating the fruiting stage in the organism. This medium will support the growth of other myxobacteria, but the great efficiency it has shown for Myxococcus xanthus is not necessarily seen with other species.

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20 1

We have used a medium which is essentially that used in Stanier plates 'for the cultivation of cellulolytic Sorurixium spp. This medium isMedium for cellulolytic Sorangium spp. KN03 1.og KzHP04 1.og MgSO4.7Hz0 0.2g CaCI2.2tI20 0.1 g FeCls. 61 I n 0 0.02 g Cellobiose 2.5 g Agar 10 fi Distilled water 1 litre

The cellobiose may be autoclaved as part of the medium, but we have found that we get better growth if this is not done. To circumvent this, we have mixed the remainder of the ingredients and autoclaved them, dissolved the cellobiose in 25 ml of water (with warming) and filtered the cellobiose solution through a bacteriological filter. T h e two sterile components were mixed aseptically. We have also used this medium without the agar in shake cultures. Still simpler carbon sources, such as glucose and xylose, have been used in lieu of the cellobiose in various concentrations, but growth has always been markedly inferior. In addition, glucose is clearly toxic in concentrations only slightly higher than that indicated above for cellobiose. Other partially defined media have been used for various myxobacteria from time to time. In general, they are modifications of the above, or they are for a special organism as, for example, that described earlier for Chondrococcus cohmnczris. T h e effects of vitamins, trace elements and other growth factors have been investigated for some particular myxobacterial isolates. In some instances, the addition of such substances appears to enhance growth but, in general, the myxobacteria, as a group, do not appear to possess requirements for a particular factor or group of factors, nor do they appear to have any particularly exotic requirements. It should be possible to work out a completely defined medium for any myxobacterial isolate, as Dworkin has done for his strains. A final comment on the selection of media for myxobacterial cultivation is that one may have trouble growing a myxobacterial isolate on any of the reasonably defined media if the isolate has not been essentially purified first. In spite of the antibiotic and lytic substances produced by many of the myxobacteria, they do not compete well, and they are slower growing than are the common eubacteria and fungi. Hence, they never appear on rich, defined media if sizeable numbers of contaminants are present in the seeding material. Therefore, purification of the isolates is essential before one can hope for success in cultivating the desired organism under definitive conditions.

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B. Purification T h e copious polysaccharide slime produced by the myxobacteria is responsible for two major problems encountered in purification procedures: (1) the cells tend to stick together in clumps; and (2) associated and contaminating organisms may be entrapped and, hence, protected in the slime. For these reasons, the usual purification techniques of dilution and singlecell isolation are not generally satisfactory for most myxobacteria. We have been able to obtain single, sometimes pure, colonies on occasion when harvesting lyophilized cultures with procedures that essentially amount to dilution plating of the lyophilized material ; usually, we have been unsuccessful. T h e use of antibiotics and other inhibitors in purification is likewise of limited value. Antifungal inhibitors can be utilized readily, but contaminating eubacteria, not fungi, present the most persistent problems. A sizeable proportion of these are Gram-negative, as are all of the myxobacteria. We have found little difference between the two groups in sensitivity to various of the antibiotics we have tested. Since the myxobacteria produce highly resistant structures, the fruiting bodies, a reasonable approach to purification is to wash these structures in various cleansing and disinfecting agents. This can be done with limited success. We have sometimes been able to produce successively cleaner cultures by immersing fruiting bodies in solutions of sodium hypochlorite, mercuric chloride, various detergents, ordinary soaps and other such agents for varying periods of time, followed by growing them on a suitable medium, and then repeating the process. We have also had many failures. The primary difficulty, of course, is that contaminating organisms are often embedded in the slime of the fruiting body along with the myxobacterial cells. Cleansing the fruiting body surface will not necessarily remove those within the mass, and a procedure that will destroy or inhibit the contaminant in the slime is apt to do the same with the myxobacterial cells. McCurdy (1963) reviews the purification procedures applied to the myxobacteria, and presents a method he has found to be successful. This paper is well worth consulting before embarking on pure culture isolations. McCurdy’s method is essentially a dilution procedure preceded by growth on media that tends to favour myxobacterial growth over that of the eubacterial contaminants. Cellular material from these media is homogenized, dispersed in serial dilutions, and seeded in pour plates. Discrete colonies can be picked after 5-10 days of incubation at 27°C. With several of the myxobacteria, we have had considerable success in making rapid transfers from the advancing edge of the vegetative colony. This can be done on the rabbit-dung pellet plates described earlier with representatives of the Myxococcaceae, Archangiaceae and Polyangia-

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ceae, since the swarm will usually grow across the agar from the pellet. With the cellulolytic members of the Sorangiaceae, we use a similar plate, which we call “filter-paper-strip plates”. These are exactly the same as the previously described Stanier plates, except that two 1 x 14 in. strips of filter paper, instead of the usual complete disc, are placed on the agar surfaces. The myxobacterium is seeded on the paper strip and, in most instances, will grow onto the agar from the paper where transfers can be made from the cleanest sections of the advancing edges.

C. Culture conditions Little need be said about the conditions of temperature, p H and light required for cultivation of the myxobacteria. Certainly, the optimum conditions for a given isolate may differ slightly from those for another isolate, but, in general, the myxobacteria are not fastidious in their requirements. For practical purposes, most of them grow equally well at temperatures from 25°C to about 32°C with 27”-28°C the optimum for the majority. It is usually difficult to detect differences in growth over a p H range from about 5 to 8, with most isolates continuing to grow appreciably down to levels below p H 4 and up to those slightly above p H 9. We have isolated soil forms in equal profusion from soils ranging in p H from 3.0 to 9-5; we have been unable to detect appreciable preferences in pH for growth over this range for these same isolates in the laboratory. Until one has established otherwise for a particular isolate, a p H of 7.0-7.4 may well be considered optimal for general isolation and cultural work. Although the myxobacteria are pigmented, neither the quantity nor the quality of light, within reasonable limits, is of particular importance. We have not examined all possible isolates, of course, but we have grown a great many over some 15 years under varying conditions of ordinary room daylight, incandescent light, fluorescent light and total darkness with no appreciable differences. As with pH, until one has determined specific light requirements for a particular isolate, one may perform routine isolation and cultivation of the myxobacteria under those light conditions most readily available.

D. Liquid culture In general, liquid culture is not a particularly useful rnethad of cultivation for the myxobacteria. The principal deterrent, again, is the slime. Myxobacteria do not grow in a dispersed fashion. Nor do they grow in the liquid medium itself. Rather, they produce clumps on the walls of the culture container along the splash line, or they grow in clumps and irregular pellets on the bottom of the container. Usually, the liquid medium itself remains entirely clear, rather than becoming turbid as one might expect, throughout

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the course of growth. For some purposes, this is an advantage; it is a simple matter to separate the supernatant from the cellular components, and to harvest either fraction. On the other hand, it is quite impossible to apply photometric methods. Dworkin and his colleagues, at the University of Minnesota, have contributed extensively to the biochemistry and morphogenesis of the myxobacteria over the past few years. One of the reasons they have been so productive is that they have used a strain of Myxococcus xanthus, which will grow dispersed in liquid culture. For their purposes, this isolate has served beautifully, although it has, quite properly, often been referred to as a “laboratory strain”. In all probability, strains of other myxobacterial species that will grow in a dispersed fashion in liquid media can be selected or produced. T h e same media described earlier, minus the agar, of course, have been used for liquid culture of the myxobacteria. These media may be dispensed in flasks and placed on shakers, or they may be established as cultures aerated by bubbling. In our experience, none of the myxobacteria grow particularly well in stationary liquid cultures, although some growth will result.

E. Problems encountered during cultivation Although none of these problems is unique to the myxobacteria, several of them are particularly prominent when one is working with this group of organisms. Consequently, there is merit in discussing some of the common difficulties the myxobacteriologist encounters rather frequently. There are two distinct aspects to the problem of the eubacteria associated with the myxobacteria. As has been discussed above, it is relatively difficult to purify many of the myxobacteria because of the copious slime. The first aspect, then, is simply the problem of getting rid of the associant. The second aspect may be an even more difficult problem, and possibly one of considerably broader biological import. It is entirely possible that some of the myxobacteria may exist in something of a symbiotic relationship with certain eubacteria. Certainly, this has not been definitely established. Zmshenetsky (1945, 1946) has suggested that such an association between Sorangium spp. and Nitrosomonas spp. may result in a synergistic or symbiotic increase in nitrifying potential ; he did not document this suggestion. I t has been our impression that many isolates of cellulolytic species of Sorangium are intimately associated with another organism that may play a role in controlling the level of the glucose pool in the environment. We have no real evidence to support this impression; we do know that some isolates of Sorangium are glucose-sensitive, that another organism is often present and that we are unable to grow the Sorangium spp. when the associant is

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ultimately removed. It is pertinent now only to point out that the myxobacterial-eubacterial association may be a problem area, which the worker with these organisms should bc advised to keep in mind. A second problem area encountered in working with the myxobacteria concerns the loss of fruiting ability and the change in fruiting body morphology under continued cultivation. Little need be said about the loss of fruiting ability; isolates that produced fruiting bodies in abundance when they were isolated sometimes only grow vegetatively after repeated transfers. An agar surface is clearly not conducive to the production of the complex fruiting bodies, and fruiting can often be enhanced by transferring the isolate onto plates containing pieces of dung, bark, paper or other such materials embedded in, but projecting above, the agar surface. Other isolates permanently lose all ability to fruit after continued cultivation; that is, no known nutritional or physical substrate changes can stimulate them to again produce the resistant structures even though they continue to grow very well vegetatively. Closely associated with the loss of fruiting, in fact sometimes a progressive step toward total loss, is a change in fruiting-body morphology. T h e intricate Chondromyces spp. fruiting bodies often become unrecognizable as to species, although they usually remain recognizable as to genus if they fruit at ali. Archangium, Sorangium and Polyangium spp. routinely produce masses of cells and slime that look not unlike the normal fruiting bodies in gross appearance, but in which the internal morphology fails to delimit as expected. Distinctly stipitate forms of Myxococcus, which can readily be identified as Myxococcus stipitatus when isolated, invariably fail to produce the stalk after a few transfers and, then, must be identified as Myxococcus fulvus or Myxococcus xanthus. On numerous occasions, we have observed a curious, and most interesting, occurrence in Archangium primigenium cultures; suddenly tiny solitary cysts on stalks, which are perfectly typical of Podangium gracilipes as found in nature, have appeared in profusion. T h e cultures in which this has occurred have not been single-cell-derived cultures, but they have been perfectly pure, typical Arch. primigenium cultures for several months. Such observations have caused us to reflect on the validity of current myxobacterial systematics. I t is not clear whether such variations in myxobacterial fruiting-body morphology under laboratory-culture conditions are genetically inherent, nutritionally controlled, stimulated by the culture environment or a combination of these possibilities. It is clear that they are common occurrences. Yet another aspect of fruiting-body modification in culture can best be described as germination of the cysts, a short, local swarming of the cells, followed by a re-encystment. In nature, one would expect the so-called

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typical fruiting bodies to be formed as the environment became less moist, and for the fruiting bodies to dessicate considerably; this is usually the condition in which one finds them in nature. In the culture dish, the typical fruiting bodies may be formed in 4-7 days, but, under the continuing conditions of high humidity in the dish, they fail to dry-out and become truly mature resistant bodies. Instead, the cells swarm after another 3-4 days, produce a small, malformed secondary cyst on top of the first one, swarm again, re-encyst, etc. This may happen three or four times. T h e process results in a completely abnormal fruiting body of partially empty cyst walls and partially encysted cell masses, often roughly in the form of a chain of decreasingly smaller masses. One can circumvent this phenomenon somewhat by opening the culture dish, thus drying the environment, just as the fruiting bodies are forming. A final problem associated with cultivation of some of the myxobacteria is that of loss, or change, of pigmentation. The problem is most apparent with isolates of the cellulose-decomposing Sorangizim spp., but it occurs with other myxobacteria also. T h e pigmentation generally is quite stable once the myxobacterium has been purified and cultivated for a short time, although some isolates tend to continue to produce colour “sectors” for long periods of time. We have strains of Myxococcus spp. in which this has occurred after several years of culture and lyophilization; we see it quite routinely in some isolates of the cellulolytic Sorangium spp. T h e most pronounced and, hence, most troublesome changes occur between the original isolation, the purification stages, and the establishment of uniform clones. T h e worker with myxobacteria should be aware of such possible changes in pigmentation.

IV. T H E MAINTENANCE OF MYXOBACTERIAL CULTURES Although one may encounter considerable difficulty in purifying and cultivating many of the myxobacteria, once this has been accomplished and an isolate established in culture, the maintenance of such cultures is not particularly difficult. They survive well when subjected to routine microbial maintenance methods, and, in addition, they can be maintained in fashions generally not applicable to eubacteria. The methods of maintenance can be summarized briefly and readily.

A. Maintenance as living cultures As with most bacteria, the majority of the myxobacteria can be maintained as living cultures at either room or refrigeration temperatures if they are transferred periodically. It has been our experience that transfer of roomtemperature-maintained cultures every 2 weeks is appropriate. As a general rule, none of them is lost when they are transferred only every 4-6 weeks.

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McCurdy (1963) has found that a wide range of species may be maintained if transferred only every month. Since myxobacteria survive well for 4-6 weeks at room temperatures, there is little advantage in keeping living cultures at refrigerator temperatures. They must still be transferred every 4-6 weeks if one wishes to assure viability, although this period between transfers may be lengthened slightly with relatively little loss. at 15"-20°C with no loss of fruiting competence

B. Maintenance by freeze drying Most of the myxobacteria lend themselves to freeze-drying procedures, and maintenance in this state, very readily. No study of a broad spectrum of species under varying conditions of lyophilization has been reported in the literature, although various species have been lyophilized in various laboratories. We have lyophilized several isolates of Myxococcus spp., in both the vegetative and resistant states, with positive results in all cases. We routinely lyophilize dozens of isolates of cellulolytic members of the genus Sorangium, nearly always with excellent results. We have lyophiIized successfully both fruiting and non-fruiting cultures of several species of Chondromyces, and we have done the same with Polyangium fuscum. We have been unsuccessful with most of the Archangiurn spp. we have attempted to lyophilize, although we have had occasional successes. We have compared re-hydrated skim milk, re-hydrated beef serum, and physiological saline as the suspending medium in which to freeze-dry various myxobacteria. All myxobacteria tested survived in all three of the media, although the number of colonies recovered from the saline was much reduced. We have compared storage of the lyophilized product at room temperature and at - 20°C and, as expected, found no difference. We have harvested Iyophilized myxobacteria annually over a 7 year period with comparable recovery each year; we are convinced that their recovery could be achieved after several more years. The procedure for freeze-drying the myxobacteria is simple, and there is nothing unusual about it. The organism may be grown either on agar or in liquid medium. Surprisingly? it is as easy to strip or scrape most myxobacterial colonies from agar surfaces as it is to harvest the cells from liquid. In either event, the aggregates df cells and slime are homogenized gently in order to separate them into small units. The homogenate is centrifuged to a soft pellet so that the supernatant may be decanted, and sterile skim milk is added directly to the centrifuge tube, which is then shaken. We have never been particularly concerned about the volume of cell material to skim milk; our standard procedure is to use a pellet the size of a small pea in 20-25 ml of milk. Since practically all of the myxobacteria are pigmented, the resultant suspension is obviously coloured; coloured milk has been our

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yardstick for suitable proportions of cells to milk. Ten drops of cell-milk suspension are placed in a standard glass lyophil tube, quick-frozen at - 70°C, dried overnight, and sealed under vacuum. T h e tubes are harvested by scoring and cracking the tube, pipetting a routine liquid medium (McCurdy's or Stanier's) into the tube, mixing by drawing and expelling into the pipette several times, and spreading the re-suspended material on the surface of a suitable agar medium with the pipette. We prefer harvesting onto agar initially so that we may examine and count the colonies resulting, which we cannot do in liquid. Freeze-drying maintenance of most myxobacteria, certainly those that are commonly isolated and studied, is simple and efficient. I n general, it is clearly the maintenance method of choice for preserving numbers of isolates for periods of time.

C. Maintenance by freezing We have maintained some isolates of Myxococcz~s,Sorangium and Chondromyces for several months by simply growing them on slants, freezing them at -2O"C, and storing them at this temperature. T h e numbers of species that survive such storage, and the lengths of time for which they survive, both appear to be less than under conditions of continuous transfer of living cultures or of lyophilization. However, this is a crude observation, since no actual study has been made of maintenance at -20°C. A few myxobacteria have been subjected to freezing and storage at liquidnitrogen temperatures. It appears that survival under these conditions is good, but this, again, is a statement based on only a few attempts and not on a methodical study. In general, the other methods of maintenance of myxobacteria are sufficiently effective, so that the space-demanding lowtemperature methods are inappropriate.

D. Maintenance of desiccated resistamt structures Since the myxobacteria produce highly resistant structures, the fruiting bodies, which are not found in other bacteria, it is obvious that they may be maintained in this state under certain circumstances. I t is appropriate to mention two such aspects in the present consideration, the dried fruiting bodies per se, and preservation in soil in which, presumably, the resistant state has been formed. All of the myxobacteria produce their characteristic fruiting bodies on bits of dung, decaying plant debris, bark, etc. I t is then a simple matter to place these bits of detritis in small boxes and to store them either at room temperature or at refrigerator temperature. T h e myxobacteria have a remarkable tenacity for survival in this resistant state. For example, I have been successful in germinating bits of fruiting material which had been stored under

IX. MXYOBACTERIA

209

ordinary herbarium conditions for over 60 years. On the other hand, it is depr that a considerable percentage of such “naked” fruiting bodies rapidly hetheir viability. In summation of this point, desiccated fruiting bodies of the myxobacteria on bits of organic debris can be maintained for considerable periods of time, but this cannot be considered a reliable method of etenance. Most of the myxobacteria, possibly all of them, inhabit the soil under K)me natural conditions. When the soil environment becomes dry, they presumably produce their respective fruiting bodies, and survive the dverse conditions in this resistant form for many months. We have recovered them from dry desert soils that were so hot that they could not be held in the hand ( > 70°C); we have recovered them from soils that were so h e n that we had to chip the sample out of the soil mass ( < 0°C). Growing myxobacteria in pure culture in moist, sterile soil, and permitting the soil to desiccate at a normal rate should be a satisfactory method of maintenance. For some of them, at least, this is true. We have prepared autoclaved dishes of soil in which we have incorporated pulverized filter paper as a cellulose source before seeding with cellulolytic Sarungium spp. These dishes have been stored at -20°C) 4”C, 25°C and 40’C; others have been stored under humidities of 0%) 33%, 66% and 95%; some of them have been periodically moistened, others have been periodically thawed and re-frozen. From all of them, with one exception, we have been recovering the organisms in profusion after 7 years. The one exception has been the dish stored in 95% relative humidity; here the organism was lost after only a few weeks. The same soil storage method of maintenance has been successful with two Myxococc~(sspp. In these instances, a eubacterium was mixed with the mil instead of cellulose before the myxobacterium was seeded into the plate, dnce Myxococcus spp. all appear to be eubacteriolytic; certainly, none of them are cellulolytic. Although no systematic study of various species has been made, it seems quite possible that such soil preparations can be a leasonable method of maintaining the myxobacteria. We have obtained species of Myxococcus, Archangium, Polyangium, Sorangium and Chondrouyces from soil samples which had been collected over a year earlier. REFERENCES Breed, R. S., Murray, E. G . D., and Smith, N. R. (1957). In “Bergey’s Manual of Determinative Bacteriology”, 7th Ed. Williams & Wilkins, Baltimore, Md. Davis, H.S. (1922). U S . Bur. Fish. Bull., 38, 261-280. Dworkin, M. (1963). J. Buct., 86,67-72. Fish, F. F., and Rucker, R. R. (1943). Trans. A m . Fish. SOC.73, 32-36. Geitler, L. (1925). Arch. Protistenk., 50, 67-88. 10

210

J. E. PETERSON

Gilbert, H. D., and Martin, G . W. (1933). Stud. nut. Hist. Iowa Univ. Griif, W. (1962). Arch. Hyg. Bakt., 146, 481491. Imshenetsky, A. A. (1945). Microbiology, 14, 177-190. Imshenetsky, A. A. (1946). Nature, Lond., 157, 877. Imshenetsky, A. A., and Solntzeva, L. (1937). Microbiology, 6, 1-15. Jeffers, E. E. (1964). Int. Bull. bact. Nomencl. Tuxon., 14, 115-136. Krzemieniewska, H., and Krzemieniewski, S. (1926). Acta SOC.Bot. Pol., 4, 1-54. Krzemieniewska, H., and Krzemieniewski, S. (1927). Acta SOC.Bot. Pol., 51-20, Krzemieniewska, H., and Krzemieniewski, S. (1937). Bull. Acud. pol. Sci. Lett., Ser. B , Sci. nut., 1, 1-59. McCurdy, H. D. (1963). Can. J, Microbiol., 9, 282-285. McDonald, J. C., and Peterson, J. E. (1962). Mycologiu, 54, 368-373. Mishustin, E. N. (1938). Microbiology, 7, 427-444. NorCn, B. (1952). Svensk bot. Tidskr., 46, 324-365. Ordal, E. J., and Rucker, R. R. (1944). Proc. SOC.exp. Biol. Med., 56, 15-18. Singh, B. N. (1947). J. gen. Microbiol., 1, 1-10. Stanier, R. Y. (1942). Bact. Rev., 6, 143-196. Thaxter, R. (1892). Bot. Guz., 17, 389-406. Winogradsky, S. (1929). Annls Inst. Pusteur, Paris, 43, 549-633. Zhukova, R. A. (1959). Microbiology, 28, 69-74, 904-910. Zhukova, R. A. (1960). Microbiology, 29, 220-228. Zhukova, R. A. (1962). Microbiology, 31, 10541060.

CHAPTER X

Isolation, Cultivation and Maintenance of Mycoplasmas R. J. FALLON IZuchill Hospital, Glasgow, Scotland AND

1'.

WHITTLESTONE

School of Veterinary Medicine, University of Cambridge, England I.

Introduction . . A. Nomenclature B. Characteristics . C. Sources and diseases D. Species .

.

. .

.

. .

11. Methods of Isolation of Mycoplasmas . A. Specimens and transport . B. Isolation media . C . Conditions of culture for primary isolation 111. Purificationof Cultures . A. Cloningmethods B. Differentiation from bacteria and artefacts C. Confirmation of identity as a mycoplasma

.

IV.

V.

VI.

VII.

Growth of Mycoplasmas . A. Characteristics of growth B. Measurement of gTowth .

. . .

.

. .

.

.

.

.

.

Transportation of Cultures

References

.

222 222 225 237 238 238 239 245 245 245 247

.

Handling of Mycoplasmas . A. Subculture . B. Cultivation and concentration for serological studies C. Concentration for biochemical studies . D. Hazards and precautions E. Microscopy . Maintenance of Mycoplasma Cultures A. Serialpassage . B. Low-temperature storage C. Freeze-drying .

212 212 212 216 221

249 249 249 251 252 252

.

258 258 259 260

.

.

260 .

261

212

R . J. FAL1,ON AND P. WHITTLESTONE

1. INTRODUCTION Mycoplasmas occur in nature as normal flora or pathogens in humans, animals and birds. Until recently those recognized were principally of veterinary importance and, indeed, the first mycoplasma to be isolated, Mycoplasma mycoides, was cultured from cattle suffering from contagious bovine pleuropneumonia (Nocard et al., 1898). The great epizootics of this disease and much of the early history of mycoplasmas are reviewed by Klieneberger-Nobel(l962) and Turner (1959). It was not until 25 years after the isolation of M . mycoides that BridrC and Donatien (1923) isolated the second mycoplasma species-this from infectious agalactia of sheep and goats. Because of the close resemblance between this organism and the bovine pleuropneumonia organism, the name pleuropneumonia-like organism (or PPLO) was adopted, and until recently members of this group have usually been referred to as PPLO.

A. Nomenelatwe l’he first binsminal name given to the organism of contagious bovine pleuropneumonia was Asterococcus mycoides (Borrel et al., 1910). However, the generic name Asterococcus is illegitimate because it is a later homonym of the algal genus Asterococcus (Scherffel, 1908; Editorial Board, 1955). T h e Editorial Board of the International Bulletin of Bacterial Nomenclature and Taxonomy (1955) suggested two alternatives-Mycoplasma (Nowak, 1929) and Borrelomyces (Turner, 1935). Edward and Freundt (1956) proposed the use of Mycoplasma as the generic name on the grounds of priority and general usage. Their recommendations have been accepted by most workers. The name Mycoplasma is derived from “fungus-form” which describes the pleomorphic forms seen on microscopy of the organism. Recently it has been proposed (Edward and Freundt, 1967; Edward et al., 1967) because of the fundamental differences between mycoplasrnas and the Class Schizomycetes (see page 213) that a new Class Mollicutales (from mollis, soft, pliable, and cutis, skin) should be established for the Order Mycoplasmatales, parallel to, but distinct from the Class Schizomycetes. T h e differentiation of mycoplasmas into species is considered later (page 221).

B. Characteristics Mycoplasmas are the smallest known free-living forms of life. They multiply in cell-free culture media, although most species require complex substances such as protein and sterols to support growth. Mycoplasmas differ from viruses and rickettsiae in that they do not require living cells for growth and from bacteria in that they have no rigid cell wall

X. MYCOPLASMAS

213

nor cell-wall mucopeptide or its precursors. This lack of a rigid cell wall explains the pleomorphic forms seen when these organisms are examined by microscopy. The smallest cells, known as minimal viable units or elementary bodies, may be as small as 125 nm in diameter (which is well within the size range for viruses). The interpretation of the details of the life cycle varies with differentworkers using different mycoplasmas and methods. Some workers report that the elementary bodies enlarge into forms up to 2 pm or more in diameter and have a potential for growing into branching filaments (Razin et al., 1967). These forms themselves give rise to further elementary bodies (Edward, 1967). Other workers report binary fission of the elementary bodies (Furness et al., 1968b; Morowitz and Maniloff, 1966). Because of the small size of the elementary bodies and also, in the case of the larger forms, their plasticity, mycoplasmas can pass through filters which retain bacteria. This has led, on many occasions, to mycoplasmas being confused with viruses. Motility, characterized by the gliding of rods and the spinning of spherical forms, has been observed with some mycopfasmas (Andrewes and Welch, 1946; Nelson and Lyons, 1965). Mycoplasmas, with the exception of Mycoplama laidlam'i, and Mycoplasma granularum (Tully and Razin, 1968) are dependent on cholesterol for growth, this sterol being an important component of their cell membrane. M,laidlmii, and probably M, granularum, differ from other mycoplasma species so far recognized in that they can synthesize carotenoids. It has been suggested that carotenoids may have the same function as sterols, in the non-sterol requiring strains (Smith, 1963); this would explain the ability of these strains to grow in cholesterol-free medium. Because of the absence of cell wall mucopeptide, mycoplasmas are usually resistant to antibiotics such as penicillin which act on bacterial cell walls, although Mycoplasma nezrrolyticum is temporarily inhibited by even low concentrations of penicillin (Hottle and Wright, 1966). Cephaloridine, however, which is also thought to interfere with the synthesis of cell walls inhibits the growth of a number of mycoplasma species (Taylor-Robinson, 1967; Fallon and Hutchinson, 1967). Antibiotics which act on other metabolic pathways (such as protein metabolism) inhibit mycoplasmas; thus the tetracyclines inhibit a wide range of mycoplasmas. The action of tetracycline and other antibiotics can be used to aid in the differentiation of viral and mycoplasmal cytopathic agents in tissue cuItures. The effect of a wide range of antibiotics and drugs on mycoplasmas has been considered by Newnham and Chu (1965). Of the mycoplasmas of human origin, only Mycoplasma pneumoniae and T-strain mycoplasmas are inhibited by low concentrations of erythromycin (Shcpard et al., 1966). The growth of mycoplasmas, either in fluid or on solid mcdium, can bc

214

R . J . FALLON A N D P. WHITTLESTONE

inhibited by specific anti-sera; this feature will be discussed later (p. 221). On solid media, most species of mycoplasmas form colonies wicli a central nipple growing down into the medium. This gives the colonies a “poached egg” appearance when viewed by transmitted light. Mycoplasma colonies vary in size from those of the ‘I’-strains which are only 10-20 prn in diameter up to 600 p m in diameter for a few species. ‘1’-strain colonies may not show the typical “poached egg” appearance and the colonies of M . pneumoniae, which are smaller than those of most other mycoplasmas also have an atypical reticulated appearance. Mycoplasma suipneumoniae colonies have no central nipple. Figs. 1 , 2 , 3 and 4 illustrate some of the colony types. Because of their

FIG. 1. Colonies of Mycoplasma gallinarum, strain B2, 6 days incubation. Unstained x 40.

FIG.2. Colonies of Mycoplasma B3, 6 days incubation. Unstained x 40.

X. MYCOPLASMAS

215

small size, niycoplasma colonies can usually oiily be seen with ease by using low power (c.g., x 25) magnification. 'l'-strain colonies may only be seen using a higher magnification ( x 100).

FIG.3. Very large mycoplasma colonies, strain B3. Unstained x 24.

FIG. 4. Colonies of Mycoplasrna szripneumoniae, 7 days incubation. Unstained x

40.

Mycoplasma colonies may be mistaken for very small bacterial colonies (seep. 241) and they have many features in common with colonies of bacterial L-forms which may lead to the two being confused (see p. 239). For further

216

R. J. FALLON AND P. WHITTLESTONE

general information on the mycoplasmas and L-forms the reader is referred to a multi-author book entitled “The Mycoplasmatales and the L-phase of Bacteria” (Hayflick, 1969).

C. Sources and diseases Mycoplasmas have so far been isolated only from members of the animal kingdom except for the original isolation of M. laidlawii from sewage. This mycoplasma, however, has also been isolated from man, poultry, cattle, pigs and horses (see Taylor-Robinson and Dinter, 1968), and this could presumably account for its presence in sewage. The possibility that mycoplasmas may also occur in plants has been raised by Japanese workers (Annotation, 1968). The names or designations of the more important mycoplasmas or those of particular interest are listed in Table I ; this table also contains the sites of origin of the mycoplasmas, the diseases which these organisms are associated with or cause, and key references to their isolation, pathogenicity and serological identification. Many mycoplasma isolates have not been adequately characterized and are not included in the table. In human medicine, only one mycoplasma is of proven pathogenic significance, but in veterinary medicine, mycoplasmas are the causal agents of a number of economically very important diseases. The most important diseases are as follows. 1. Respiratory diseases Primary atypical pneumonia in man is caused by M. pneumoniae (Eaton agent). This disease is particularly a problem when young persons live together in groups under conditions of fairly close contact, e.g., military recruits (see Chanock et al., 1967). Contagious bovine pleuropneumonia, caused by M. mycoides var. mycoides, has been one of the major disease scourges of the world, but has been eradicated from America, Britain and many European countries. It still exists in Asia, Africa and Australia (see Klieneberger-Nobel, 1962; Turner, 1959; Piercy, 1960). A similar severe disease in goats, contagious caprine pleuropneumonia, is not well defined aetiologically, but the causation by mycoplasmas is agreed by most workers. The disease is of major economic importance in the Middle East, West and East Africa, India and Pakistan (see Longley, 1940, 1951; Hudson et al., 1967). Enzootic pneumonia of pigs, caused by M. suipneumoniae ( N . hyopneumoniae), is of major economic importance on a world-wide basis, because of its chronic depressant effect on the efficiency of food production (see Betts et al., 1955; Goodwin, 1963).

TABLE I

Mycoplasmas of medical or veterinary importance or interest Key references

,

I

Host

&me or designation

Site of origin

Man

J I .fermentans

G-U tract ’Bone marrow G-U tract Pharynx .Blood Eye ,Oral cavity

__

___-_-

-

’

Luns

M. orale type 1 (syn. M. pharyngis)

pharynx .Bone marrow Pharynx

M. orale type 3

Pharynx Respiratory tract

M.orole type 2 -1.3.pneumoniae

-

1

M . hominis

M . laidlawii M . lipopfriliae

Pathogenicity -

?

-

?

-

+

+

+

M.saZivarium

Pharynx

-

T-strains

G-U tract

?

“Navel” Several species mates One species

Pri-

Umbilicus Oropharynx Vagina

-

Ruiter & Wentholt (1955)

3

serological identification Huijsmans-Evers& Ruys

Nid tk Edward (1953);

Stokes (1955,1959); Tully et af. (1965) C N m ~ t aconjunctivitis l Jones & Tobin (1968) Razin et (11. (1964) CPneumonia Del Giudice & Carski (1968) Herdenchei?et a1 (1963) Taylor-Robinson et al. (1965) Purcell & Chanock (1967) Eaton et al. (1944) Primaryatypical pwumoma Chanock et ul. (1962) Febrile respiratoryPelvicSepsis

+

Pharynx

Disease

isolation and/or pathogenicity

(1956) Del Giudice & Carski (1968) Clyde (1964) Taylor-Robinson et a1 (1964) Ta$lor-Robinson et al. (1965) Purcell & Chanock (1967) Chanock etal. (1962)

tractdisease

Rifkind et al. (1962) Myringitis Erytheme multiforme Ludlam et al. (1964) ; LyeU et al. (1967) Nicol & Edward (1953) CNon specific urethritis

Nicol&Edward(1953); Edward & Freundt (1956) Purcell etaZ. (1966a)

Shepard (1954,1956); Fordetal.(1%2); Csonka etul. (1966) Taylor-Robinson & Purcell (1966) Ruiter & Wentholt (1955) Lemcke (1964)

Davidson & Thomas (1968) Davidson &Thomas (1968)

TABLE I-continued Key references Host

Name or designation

Ox

M . mycoides var.

i

mycoides (Bovine group 1) M . bovigenitalium (Bovine group 2) M.laidlawii (Bovine group 3) M . boeirhinis (Bovine group 4)} .M.agalactiae var. bovis (syn. M. booimastitidis) (Bovine group 5 )

Genital tract Respiratoryand G-C tracts Respiratory tract 8Mammary gland

“Sauire” and others I “Squire” (Bovine group 6) “N29” and others (Bovine group 7) “D 12” (Bovine group 8) T-strains

Respiratory tract Mammary gland Joints hTasalcayit!

}

Goat

Site of origin Respiratory tract Mammary gland

Mammary gland

+

Disease

+ ? ? ?

Strains related to M . mycoides var. mycoides Strain P Respiratory tract Joints Peritoneum

seroloeical - identification

Nocard et al. (1898); Turner (1959) ;

Mastitis

Stuart et al. (1963);

Leach (1967)

Afshar& etHaig al. (1966) Hoare (1964)

Leach (1967)

Leach (1967)

Klieneberger-Kobel(1962)

? ?

isolation and/or pathoeenicitv -

Contagious bovine plei iropneumonia

CPneumoniaand shipping fever Mastitis Salpingitis CPneumo-enteritis CMastitis CArthritis

>

G-u tract

1

Strain 0

+

+

M . agalactiae (strain H of Cottew & Lloyd Mammar). gland 1965) &respiratory tract (type A of Arisoy et al. 1967) .M.Capri Respiratory tract Mammary gland, joints & other sites

Strain Y

Pathogenicity

Langer & Carmichael(l963); Harbourne et ai. (1965); Leach (1967) Dawson et al. (1966) Hale et al. (1962); Leach (1967); Carmichaeletal. (1963); Jainetal. (1967) Jain et al. (1967) Hartman et al. (1964) Langer Sr. Carmichael(l963) Leach (1967) Connoleetal. (1966); Simmons & Johnston (1963) Leach Hudson& Etheridge (1963) Leach (1967) Taylor-Robinson et al. (1967)

+

+

-BridrC & Donatien (1923); Cottew& Lloyd(1965); Arisoy etal. (1967); Watson etal. (1968) Contagious caprine ’Longley (1940,1951) ; pleuropneumonia, Debonera (1937); contagious agalactia Cordyetal. (1955); and other syndromes Hanko& Otterlin(1955); .Dhandaeta/. (1959)

Contagious agalactia and pneumonia

Contagious caprine pleuropneumonia Polyarthritis

Hudson et ol. Laws(1956)

Fibrinous peritonitis or chrovic polyarthrit!is

Hudson etal. (1967); Nasri (1967a); Cottew et al. (1968) Hudson etal. (1967); Nasri (1967a)

Hudson etal. (1967) Hudson et al. (1967)

TABLE I-~otltimred r

Host Name or designation Dog

Site of origin

Astmococcuscanis I Asterococcus canis 11 ather isolates

Respiratory tract Respiratory tract Respiratory tract

M. felis M. gateae Mouse M. neurolyticumb Cat

Eye Saliva Brain Respiratory tract Respiratorytract

& rat

M .pulmonisb

M.arthritidis

1

(syn. M. horninis type 2 or L 4) M.laidlaw*i(PGS)

Pathogenicity

Disease

1-}

Shoetensack(1934; 1936a

?

Conjunctivitis

+ + + ++

Rolling disease Conjunctivitis Infectiouscatarrh Otitis media Pneumonia

+

Arthritis

-

Joints Lw? Respiratory tract

-

Poultry M.m t i s M. galliseptuum (Avian serotypeA)

Respiratory tract Respiratorytract

? "Duck sinusitis Fowlcoryza Turkey sinusitis

Respiratory tract Pericardial sac & respiratory tract

--

-

I

M.gallinarum (Avian serotypeB) M.i w s (Avian serotype G) M.meleagridis (Avian serotype H) M.synmae (Avian semtype S ) 14 other serotypes from poultry

+ +

+

Respiratory tract +

Joints Mainly respiratory tract

Klieneberger(1938)

-&?

Rabbit M-pulmonis

}

isolation andlor pathogenicity

Key references * 7 serological identification

?

-

Air sacculitisof turkeys Infectioussynovitis of poultry

Cole etal. (1967) Coleet at. (1967) Cole etal. (1967) Cole et al. (1967) Findlayetal. (1938; Nelson (1950);Tully(1965)) Lemcke (1964) f Klienebereer & Steabben Lemcke (1964) . . { (1937); EdGard(l947b); [Lutsky & Organick (1966) Lemcke (1964) ; Klieneberger(1938) Edward & Freundt (1965) Tully (1965) ;Tully & seeTully(1965) Razin(1968) Deeb & Kenny (1967) Deeb & Kenny (1967) Roberts (1964a) Roberts (19%) Nelson (1936); Markham & Wong(1952); Adler & Yamamoto (1957) Yamamoto & Adler Chu(1954); (1958a&b); Fabricant (1960) Kleckner (1960); Yamamoto & Adler (1958a); . Fabricant (1960); Edward & Kanarek (1960) Yamamoto & Adler (1958a); Yoder & Hofstad(1964); Roberts (1964b); Yamamoto (1967) Dierks etal. (1967) Chalquest & Fabiicant (1960) J

a = Occasional isolationsmade from that site but it is not the usual habitat. b = Speciesusually latent. c = Mycoplasma isolated from that Ppthological material, but not established as the cause of disease. + = Pathogenic. j , = Occasionallypathogenic. ? = Uncertain or unronfirmed pathogenicity. = Nonpathogenic.

-

TABLE I-continued Key references * Pathogenlcity

Host

Name or designation

Site of origin

Dog

Asterococcus canis I Asterococcus canis I1

Respiratory tract Respiratory tract

Other isolates

Respiratory tract

-&?

Eye Saliva Brain Respiratory tract Respiratory tract

?

Conjunctivitis

+ + + + +

Rolling disease Conjunctivitis Infectious catarrh Otitismedia Pneumonia

M . jelis M .gnteae Mouse M . neurolyticumb Cat

& rat

M. pulmonisb M . arthritidis (syn. M . horninis type 2 or L 4) M . laidZawii(PG5) Rabbit M . pulmonis

Disease

r} -

(Avian serotype G)

hi.vieleagridis

}

f Arthritis Lung

i

(Avian serotype H) M . synoviae (Avian serotwe S) 14other s e r o t y p i s from poultry

Respiratory tract Respiratory tract

respiratory tract Respiratov 'IXct Joints Mainly respiratory tract

?

+ +

eDuck sinusitis Fowlcoryza Turkey sinusitis

+

+

Y

serological identification

Shoetensack (1934; 1936a Klieneberger (1938) b) Greig (1954); Switzer (1967b); Armstrong et (1968) Armstrong et al. (1968) Cole e t a l . (1967) Cole et al. (1967) Cole et al. (1967) Cole et al.(1967) Findlay et aZ. (1938 ; Nelson(1950);Tully(1965)}Lr"'cke(1964) f Klieneberger & Steabben Lerncke (1964) { (1937); EdGard(1947b); lLutsky&Organick(1966) Lemcke (1964); Klieneberger (1938) Edward & Freundt (1965) TuIly(1965); Tully& see Tully (1965) Razin (1968) Deeb & Kenny(1967) Deeb & Kenny(1967)

{

Respiratory tract

Poultry M. onotis M . gallisepticurn (Avian serotype A)

isolation and/or pathogenicity

Air sacculitis of turkeys Infectious synovitis of poultry

{

Roberts (1964a) Nelson (1936); Markham & Wong(1952); Adler& Yarnarnoto (1957) Chu(1954); Fabricant (1960) Yarnarnoto & Adler (1958a); Edward & Kanarek (1960) Yamamoto & Adler (1958a); Yarnarnoto (1967) Chalquest & Fabricant (1960)

oberts (1964a)

-

Yamarnoto & Adier (1958a& b); Kleckner (1960); Fabricant (1960); Yoder & Hofstad (1964); Roberts (1964b); Dierks e t a l . (1967)

?-

= Occasional isolations made from that site but it is not the usual habitat. b = Species usually latent. c = Mycoplasma isolated from that pathological material,but not established as the cause of disease. + = Pathogenic. -C = Occasionally pathogenic. ? = Uncertain o r un-ntinned pathogenicity.

-

= Nonpathogenic.

X. MYCOPLASMAS

221

The chronic respiratory disease of poultry, caused by Mycoplasma gallisepticum similarly causes considerable economic loss (Yoder, 1965), due mainly to retarded growth and lowered egg production. .Infectious catarrh, otitis media and pneumonia of rodents caused by Mycoplasma pulmonis, are similar common chronic diseases, and of particular importance in laboratory animals; much effort has been devoted to establishing such stock free from mycoplasma infection (Innes et al., 1957).

2. Joint involvement Mycoplasmas are associated with arthritis and tendo-synovitis in several species of animals. Arthritis is an important manifestation of contagious agalactia of sheep and goats, and several other caprine mycoplasma isolates produce arthritis in goats experimentally. Synovitis occurs in cattle inoculated with M . mycoides var. mycoides, and has been associated with bovine group 7 and a number of unclassified bovine isoIates. At least two species of mycoplasmas, Mycoplasma hyorhinis and M . granularum, are associated with arthritis in the pig. In rats, Mycoplasma arthritidis has a tendency to localize in joints and in poultry arthritis and tendo-synovitis occurs with Mycoplasma synoviae infection.

3. Mastitis Contagious agalactia of sheep and goats is a generalized disease with mastitis as one of the main clinical findings. At least three species of mycoplasmas, Mycoplasma bovigenitalium and bovine groups 5 and 7 have been associated with naturally-occurring mastitis in cattle; in addition, bovine group 4 induces mastitis experimentally.

D. Species Broad groups of mycoplasmas can be defined by their metabolic activities, e.g., there is a group that produces acid in media containing glucose, and T-strain mycoplasmas are recognized as a biotype by their small size and their unique ability, amongst mycoplasmas, for decomposing urea. It is, however, much more difficult than with bacteria, to demonstrate and define the biochemical reactions of mycoplasmas precisely. Because of this, mycoplasmas are most easily classified into species by their serological reactions. Many serological techniques have been used for the identification of mycoplasma species but the most satisfactory ones are those based either on the complement-fixation test (Lemcke, 1964) or on the inhibition of growth or metabolism of mycoplasmas, these tests being performed with specific hyperimmune antisera. The simplest test is the disc growth-

222

R. J. FALLON AND P. WHITTLESTONE

inhibition test described by Wuijsmans-Evers and Ruys (1956) as modified by Clyde (1964). In this test a plate of solid medium is flooded with a culture of the mycoplasma being examined, and after allowing the medium surface to dry, filter-paper discs impregnated with antiserum are placed on the surface of the medium in a manner analogous to that used for determining the sensitivity of bacteria to antibiotics. Specific hyperimmune antiserum will inhibit the growth of the mycoplasma for several millimeters around the disc. In some instances the incculum for the test must be diluted to obtain a zone of inhibition. It is not always possible to prepare satisfactory growth-inhibiting sera (Huijsmans-Evers and Ruys, 1956), and with some mycoplasmas, e.g., M . hyorhinis, the disc-growth inhibition test may not work (Dinter et al., 1965). In tests for the inhibition of metabolism by specific antiserum, use is made of the fact that many mycoplasmas will metabolize a substrate in liquid medium to produce a recognizable change. For instance, with the glucose-fermenting species the pH shift is from alkaline to acid (Taylor-Robinson et al., 1966a), whereas with mycoplasmas that decompose urea or arginine the change is from acid to alkaline (Purcell et al., 1966a, 1966b). In the case of M . pneumoniae, the reduction of triphenyltetrazolium chloride to a red formozan may similarly be used (Jensen, 1964; Taylor-Robinson et al., 1966b). Inhibition of metabolism by antiserum results in failure of the medium to change colour. This test is more sensitive than the growth-inhibition test and can be used in the study of natural sera. I t is quantitative and can be used not only to classify mycoplasmas into species but also can aid in the differentiation between members of a species (Fallon and Jackson, 1967; Purcell et aE., 1967). For the routine identification of species, the disc growth-inhibition test is the method of choice, but ideally, for full characterization of a newly isolated mycoplasma, other serological tests should be used. Complement fixation and metabolic inhibition tests are particularly useful ; additional information may be obtained from precipitation in agar gel and biological tests such as fermentation reactions and animal pathogenicity studies. It should be stressed that the species of mycoplasmas listed in Table I, have usually not been adequately compared serologically one with another; thus, the possibility exists that they are not all distinct species.

11. METHODS OF ISOLATION O F MYCOPLASMAS

A. Specimens and transport 1. Human sources Specimens may come from patients with respiratory tract disease (throat swabs or sputum), patients with genito-urinary tract sepsis (purulent material or swabs) or from various other tissues or situations (blood, syno-

223

X. MYCOPLASMAS

vial fluid, blisters of patients with erythema multiforme, products of conception). Ideally, infected tissue or pus should be used for isolation studies but where this is not possible specimens may be taken on cotton wool swabs. There is no firm evidence to date as to whether specimens should be submitted on specially treated swabs (such as those impregnated with charcoal or serum) in preference to plain cotton wool swabs although unpublished preliminary observations suggest that serum-impregnated swabs may be more satisfactory. However, there is no doubt that, as with any laboratory specimen, there should be the minimum of delay between obtaining the specimen and culturing it, and in some situations such as the isolation of mycoplasmas from bone marrow it may well be that any delay is critical (Murphy et ul., 1967a). Where delay is unavoidable the material should be refrigerated at 4°C (Andrews et ul., 1968) or may be forwarded in transport medium such as the one described by Stuart (1959); this medium is prepared as followsTransport Medium (Stuart, 1959) Anaerobic Salt Solution Thioglycollic acid (Difco) 2 ml Distilled water (passed through an ion exchange resin column to ensure that the water is free from 900 ml chlorine) 1 N NaOH (sufficient to bring the pH to 7.2) 12-1 5 ml 100 ml Sodium glycerophosphate (20% w/v aqueous) 20 ml Calcium chloride (1 % w/v aqueous) Agar solution Agar (Bacto) Distilled water (chlorine free) Dissolve by steaming Melt the agar solution and add the anaerobic salt

6 g 1 litre

Preparation of cnmplete medium Melt the agar solution and add the anaerobic salt solution. Adjust the pH to 7.3-7.4. Add 4 ml methylene blue solution (1 g/litre) mix well and distribute in 5 ml screw capped bijou bottles filling nearly to capacity. Autoclave at 121°C for 15 min and immediately tighten caps, When cool the medium should be colourless.

Mycoplasma culture medium is also suitable as a transport medium; either semi-solid agar medium (Stewart and Chowdray, 1968) or preferably two phase medium may be used. When semi-solid medium is subcultured onto solid medium there may be a problem in distinguishing agar debris

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R. J. FALLON AND P. WHITTLESTONE

from mycoplasma colonies ; with similar subcultures from the fluid of two-phase medium, the identification of mycoplasma colonies is easier. For T-strain mycoplasmas, Shepard (1967) has recommended the use of calcium alginate wool swabs transported in tubes containing T-strain liquid medium known as TDB-16. This medium is the same as medium A2 (p. 228) except that the basal broth is a tryptic digest of beef heart muscle prepared as described by Shepard (1956) and modified by Shepard (1967). The pH of this transport medium is adjusted to 7.6. 2. Animal sources Specimens for the isolation of mycoplasmas from the natural diseases of animals derive mainly from the respiratory tract or pleural cavity, the genitourinary tract, the mammary gland and the synovial cavities ; mycoplasmas may also be isolated from the brain of laboratory rodents. (For further details see page 229 and Table I.) Lesions in laboratory animals injected with material for the isolation of possible pathogens should also be cultured for mycoplasmas. There is a special problem in the interpretation of such cultural results in laboratory rodents, as mycoplasmas are frequently present as latent pathogens unless the animals derive from deliberately established mycoplasma-free stock. In animals, the group is usually more important than the individual; thus with the smaller domestic mammals, poultry and laboratory rodents it is often practicable to kill a few animals to harvest suitable material for mycoplasma isolations. Thus, material can often be collected aseptically at the laboratory under ideal conditions and cultured immediately or stored at - 60°C prior to culture. In the latter case, freshly made touch preparations or smears may be stained and examined microscopically for mycoplasmas, to enable a selection to be made of the most suitable material for cultural examination. M . mycoides var. mycoides and M . suipreurnoniae as seen in the natural diseases are illustrated in Figs. 5 and 6. If tissue or exudate is collected in the field the same general principles, as described under human sources, apply. As veterinary mycoplasmas may be collected some distance from the laboratory, it may be more practicable, especially in hot climates, to inoculate suitable culture medium directly, rather than to transport the swab back to the laboratory. Swabs are often used for isolating mycoplasmas from the nasal cavity. The following method was used by Hudson and Etheridge (1963) for isolating M. mycoides var. mycoides from the bovine nasal cavity: cotton wool swabs are twisted on the end of 12 in. lengths of stout wire and sterilized in 2 in. test tubes. A swab, freshly moistened with 3 ml BVF-OS broth (see page 229) is inserted 1 in. into one nostril, twisted, withdrawn and replaced in the test tube. On arrival in the laboratory the liquid is

225

X. MYCOPLASMAS

squeezed out of the swabs against the side of the tube and used to inoculate liquid medium and plates. Dry swabs may also be used to collect nasal discharges (Harbourne et al., 1965), which are then used to inoculate semisolid medium within 12-1 8 hours.

FIG. 5. Touch preparation of pneumonic lesion showing delicate mycoplasma organisms-mainly rings and bipolars. Giemsa stain, x 960.

(a)

(b)

FIG. 6. Line drawing superimposed on photographs of Giemsa-stained touch preparations of pneumonic lesions. (a) Contagious bovine pleuropneumonia lesion showing ring and filamentous forms of Mycoplasma mycoides var. mycoides x 750; (b) enzootic pneumonia of pigs showing ring and bipolar forms of Mycoplasmu suipneumoniae x 750.

B. Isolation media Very many different media have been used by different workers for the primary isolation of mycoplasmas, and generally it is not known to what extent the differences between the different media used are significant. Only

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R. J. FALLON AND P. WHITTLESTONE

a selection of the media used can be described. Many other media may be traced from the references in Table I. To obtain the optimum results in culturing mycoplasmas, only glassware with a high standard of chemical cleanliness, or non-toxic plastics should be used. Water used for preparing media should also be of high standard ; either glass-distilled water or very pure de-ionized water may be used. It may also be an advantage to sterilize media by means of Millipore filters rather than asbestos pads, and to use highly purified agars for preparing solid media. If possible, the same standards as are adopted for growing tissue cultures should be maintained. Nearly all the mycoplasma strains, can, after some adaptation, be propagated using Hayflick‘s media(a) Horse serum broth (modified from Chanock et al. 1962)

Medium No. 45 (see Lapage et al., this Series, Vol. 3A) Bacto PPLO broth (without crystal violet) 70 ml 20 ml Horse serum 10 ml Yeast extract 2.5 ml Thallous acetate 0.2 ml Penicillin Bacto PPLO broth (without crystal violet) (Difco code 0554) Suspend 21 g of powder in 1 litre of de-ionized water. Distribute in 70 ml amounts in 4 oz medical flats (or 350 ml amounts in 20 oz bottles). Autoclave at 121°C for 1 5 min. Store at room temperature. (Bacto PPLO broth with crystal violet is not recommended.) Horse serum “Wellcome” Brand, horse serum No. 3 is satisfactory; do not inactivate. Store at -30°C. Yeast extract Suspend 250 g of baker’s yeast in 1 litre of de-ionized water. Heat at 100°C for 30 min, cool rapidly and clarify by centrifugation (M.S.E. Major, 2000 rev/rnin for 30 min)--8a) g. Discard the sediment and re-centrifuge the supernatant if necessary. Dispense in 10 ml amounts, autoclave at 115°C for 10 min and store at - 30°C.

Thallous acetate Make a 1% w/v solution in de-ionized water. Seitz-filter and store in 2.5 ml amounts at room temperature.

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227

Penicillin Use benzylpenicillin sodium B.P., 100,000 units/ml in sterile de-ionized water. Store at 4°C (discard after 2-3 weeks).

+

(b) Horse serum agar (modified from Chanock et al. 1962) Medium No. 44 (see Lapage et al., this Series, Vol. 3A) Make as horse serum broth (medium No. 4 9 , except 70 ml of Bacto PPLO agar should be used instead of 70 ml of Bacto PPLO enrichment broth. Plates should be prepared on the day of inoculation. Melt the agar and cool to 50°C, mix with the other ingredients (warmed to 50°C) and pour plates-8 ml of medium per 5 cm diameterplastic Petri dish.

These media are commonly supplemented by adding 20 pg/ml of the sodium salt of calf thymus deoxyribonucleic acid (British Drug Houses Ltd., Poole, England). DNA appears to be essential for the primary isolation of a number of mycoplasmas, particularly some of the cattle strains (Edward and Fitzgerald, 1952). There is an antagonistic relationship between RNA and DNA (Crowther and Knight, 1956; Razin and Knight, 1960) the DNA being necessary to overcome an excess of RNA in the complex medium. When either RNA or DNA is present in excess, growth of mycoplasmas is inhibited. T h e basic Difco PPLO broth may be replaced by the basal broth used in Edward's medium (Edward 1947a); this broth is an ox heart infusion broth with 1% added peptone, adjusted to p H 8.0. Some workers prefer to prepare solid media using I y' Ionagar (Oxoid) plus the horse-serum broth medium No. 45 instead of the Difco PPLO agar. 1. Human strains The basic media are as described above. T h e optimal pH for the isolation and growth of M . pneumoniae is 7.8. T h e other human mycoplasmas appear to grow best between pH 5.5 and 6.5 (Morton and Roberts, 1967), although Mycoplasma hominis will grow quite well at pH 7.8. Unlike other human mycoplasmas, M . pneumoniae is resistant to methylene blue (Kraybill and Crawford, 1965); Smith et al. (1967) utilized this property of M . pneumoniae together with its ability to ferment glucose in the formulation of a selective diphasic medium. This medium consists of horse serum agar No. 4 4 overlaid with horse serum broth No. 45 to which has been added methylene blue (0.01 g/litre), dextrose (10 g/litre) and phenol red (0.02 g/litre). T h e most convenient and economical container for the medium is either a bijou bottle or a similar small screw-cap bottle of 4-5 ml capacity, the volume of broth used being about 2 ml. Throat swabs may be used to inoculate the medium which should then be incubated for 3 weeks or more if necessary.

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R. J. FALLON AND P. WHITTLESTONE

M . pneumoniae ferments the glucose in the medium and also reduces the methylene blue so that the medium changes colour from violet to green. Subcultures should then be made onto mycoplasma agar and any mycoplasma colonies identified serologically (e.g., by the disc growth inhibition test-see page 221). Sometimes, this colour change in the selective medium may be produced by yeasts or bacteria which can grow in the presence of the various inhibitors (penicillin, thallium acetate and methylene blue) present in the medium. T o check for the presence of these organisms, subcultures should be made onto suitable media, such as Sabouraud's medium (see Beech and Davenport, this Series, Vol. 4) and horse-blood agar respectively. T-strain mycoplasmas, with the exception of the slow growing Boston T-strains (Kundsin et al., 1967), will grow on horse serum agar medium No. 44 provided the pH is 6.0. However, the medium must be modified because thallium acetate is inhibitory for T-strains (Shepard and Lunceford, 1967); the concentration may be reduced or, more conveniently, the penicillin and thallium acetate may be replaced by ampicillin at a strength of 1 mg/ml of medium. A medium specifically designed for the isolation of T-strain mycoplasmas has been described by Shepard (1967). The medium, designated A2, has the following compositionA2 Medium (Shepard, 1967) Trypticase soy broth powder (Baltimore BiologicalLaboratories) 30 g 11.33 g) (Oxoid Ionagar No. 2-for solid medium Distilled water 1 litre N HCl to PH 6 Bottle in 76 ml amounts. Autoclave at 121°C for 15 min. For use addUnheated horse serum (brought to pH 6 with^ HCl) 20 mi Yeast extract 5 ml (for Agar medium 3 ml) 1000 unitslml. Benzyl penicillin Yeast extract is made by boiling a 25 % suspension of dried (active) yeast in distilled water for 2 min, clarifying by filtration, bringing to pH 6 and autoclaving at 121°C forb15 min. If Trypticase soy broth powder is not available, medium TDB-16 (see p. 224) may be used.

Presumably ampicillin may be substituted for benzyl penicillin and the wider antibacterial spectrum of this antibiotic should increase the efficiency of the medium for primary isolation purposes.

X. MYCOPLASMAS

229

The addition of urea (1-10 g/litre) to the solid medium results in the formationof slightly larger colonies which have a yellow to brown pigmentation. Purcell et al. (1966a) have described a useful indicator medium which may be used for the detection of T-strain mycoplasmas or for measuring antibody using the metabolic inhibition technique. This medium is horseserum broth medium No. 45 (without the thallium acetate) adjusted to pH 7.2 with added crystalline urea (1 g/litre) and phenol red (0.02 g/litre). Yellow to red colour change indicates the presence of urease. A serum-free medium (designatedU-6) for T-strains consists of tryptic digest broth (pH 6-0) plus pure urea (0.5 g/litre), the sodium salt of phenol red (0.01 g/litre) and benzyl penicillin (1000 units/ml) (Shepard and Lunceford, 1967). These authors found that a yellow to red colour change in this medium after overnight incubation at 36°C correlated well (90%) with methods for the actual isolation of T-strains. They also noted that even the occurrence of Proteus species in clinical material from the genito-urinary tract has not resulted in false positives. Subculture to solid medium should be made as soon as a colour change is noted because T-strain mycoplasmas die rapidly at pH 8.0 (Shepard and Lunceford, 1967). Complex media for the isolation of mycoplasmas from human bone marrow are described by Barile et al. (1966) and Murphy et al. (1967b). For other mycoplasma media see Lemcke (1965) and Haflick (1967).

2. Bovine strains M , mycoides var. mycoides can be cultured in various types of medium; a good medium for the isolation of this organism is the buffered viande-foie (meat-liver) ox-serum medium (BVF-0s medium) described by Turner et al. (1935). This medium, which is preferred by the Australian workers is prepared as follows: the basic V F broth is a peptic digest made by digesting 100 g of finely-minced lean ox muscle and 100 g of finely-minced normal ox liver (free from blood vessels) with pepsin (120 g of washed pig stomach in 1 litre of water to which 10 ml of concentrated hydrochloric acid has been added). These constituents are held at 50°C for 24 hours; the action of the pepsin is then terminated by heating to 80°C. The resultant digest is filtered through filter paper and the pH of this VF broth restored to 7.5-7-6 by adding 10% sodium hydroxide solution (ca. 40-50 ml/litre). After holding at 80°C for 15 min, finely ground powdered buffer salts (379.0 g anhydrous disodium phosphate 90.8 g potassium dihydrogen phosphate) are added at the rate of 10 g/litre to buffer the pH to 7.4. This buffered broth (known as BVF broth) is kept at 18"-19"C for at least 4 hours and then filtered through filter paper. 10% ox serum is finally added

+

230

R . J. FALLON AND P. WHITTLESTONE

to complete the BVF-OS broth which is sterilized by filtering through a Seitz EK pad. This medium is still widely used in Australia but with various modifications. For example, Hudson and Etheridge (1963) increased the serum concentration to 30% and added 100 units of penicillin per ml plus 0.167 g/litre of thallium acetate. Their solid medium was prepared from the BVF broth using 2% agar, the inactivated ox serum being added after cooling the medium to 56°C. Penicillin and thallium acetate is added in the same concentrations together with 0.071 g/litre of actidione. Cultures are incubated aerobically at 37"C, the solid-medium plates being kept in an air-tight tin containing moistened cotton wool. I n Africa, Newing and MacLeod (1956) showed that the fresh meat extract was not essential for M . mycoides var. mycoides, snd they preferred a medium based on Bacto-tryptose broth, which gave very reliable results. This medium was modified by Gourlay (1964), the composition then being as followsModified M. mycoides Broth (Gourlay, 1964) Bacto tryptose (Difco) Dextrose NaCl NazHP04 Glycerol Yeast extract (Difco) Distilled water to 1 litre Pig serum (inactivated at 56°C for 30 min) Penicillin

20 g 5 g 5g

2.5 g 5g

Ig 100 ml 100 units/rnl

T h e complete medium is sterilized by Seitz filtration. The pig serum is preferred because it contains no natural antibodies to M. mycoides var. mycoides. This medium is used for all routine isolations, titrations and vaccine production, but where p H control is necessary the NaCl is replaced by KHzPO4 1.3 g/litre and the NaZHP04 is increased to 8.7 g/litre (Gourlay and MacLeod, 1966). Compared with the Australian medium these media have the advantage of cheapness and ease of preparation. I n the Sudan, the isolation of M. mycoides var. mycoides directly onto solid medium was studied by Dafaalla (1961), who found that the medium described by Turner et al. (1935), when solidified by agar, did not produce goad colony growth. DafaaIla's best medium was prepared as follows: ox heart and ox liver were minced separately, extracted separately in double amounts of distilled water in the refrigerator for 24 h and boiled for 30 min. Each was strained through muslin and to each filtrate was added peptone (20 g/litre) and NaCl (5 g/litre), the broths being boiled until the ingredients dissolved. After cooling, the pH was adjusted to 7.6, equal

X. MYCOPLASMAS

23 1

volumes of the two broths mixed, 2.5% agar added and the mixture steamed for 30 min. After cooling to 55°C the following supplements were added: 30% ox serum, DNA (0-6 g/litre) and penicillin (30 unitslml). Dafaalla’s medium was improved by Nasri (1967b); prior to mincing, the liver was cut and washed in running tap water for 30 min to remove bile. The final broth was buffered with phosphate, the pH adjusted to 8.0 and the medium solidified with 2% agar. The mycoplasma strains, other than M , mycoides var. mycoides, isolated from cattle can usually be cultivated using media Nos. 44 and 45. A study of the nutritional requirements of mycoplasma strains, including a number of cattle isolates, has been reported by Ern0 el al. (1967). The bovine T-strain mycoplasmas have been isolated by Taylor-Robinson et al. (1967) using a medium consisting of Difco PPLO broth supplemented with 2.5% yeast extract, 20% horse serum, urea (1 g/litre) and phenol red (0.02 g/litre). Thallium acetate (0-5 g/litre) and penicillin (1000 units/ ml) is also added and the pH of the medium is adjusted to 6.0. Cultures are incubated at 35°C. Those showing an alkaline pH shift are subcultured either into further liquid medium or onto solid medium. The solid medium contains the constituents of the liquid medium together with 1% Ionagar. It is incubated in an atmosphere of 95% N2 and 5% COz. 3. Caprine strains The literature on the mycoplasmas isolated from goats is particularly confusing; many workers have studied a large number of caprine isolates, but apart from Mycoplasma agalactiae, the cause of contagious agalactia of sheep and goats, there is no general agreement about the classification of most of the isolates. The tentative classification given in Table I, follows that suggested by Hudson et al. (1967) with the addition of the strains identified by Cottew et al. (1968). The position is likely to remain unsatisfactory until type cultures of all the important isolates are compared serologically one with another. As an example of the present confusion that exists with the causal agent of contagious caprine pleuropneumonia, the original infectious mgterial worked with by Longley (1940) appears to be no longer available for comparison ; the relationships of the Cambridge culture isolated by Chu and Beveridge and named Mycoplasma mycoides var. Capri (PG3) to M. mycoides var. mycoides and to the species listed in the table, are either controversial or unknown. Many of the goat strains isolated grow well using accepted mycoplasma media, e.g., BVF-OS medium (see p. 229) or Difco PPLO medium with 20% pig or horse serum together with 0.5 ”/, yeast autolysate (Cottew and Lloyd, 1965). The Pillai (P) strains were isolated on Dafaalla’s solid medium

232

H. J. FALLON AND P. WHITTLESTONE

(see p. 230). T h e many isolates of Arisoy et al. (1967) were made by one of two methods. Firstly, conjunctival or nasal swabs were inoculated into a fluid medium prepared from an infusion of fresh ox heart to which was added 0.5% yeast extract (Difco), 20% sterile horse serum, 500 unitslml penicillin (sodium salt) apd thallous acetate (0.125 g/litre). T h e inoculated liquid medium was usually held at 4°C overnight before being plated onto agar medium prepared by the incorporation of 2% agar into the liquid medium. Secondly, milk samples were retained at 4°C overnight and then plated directly onto solid medium.

4. Ovine strains Several media have been used for the isolation of ovine mycoplasma strains. Grieg (1955) used a beef heart infusion broth supplemented with a boiled yeast extract and 1 or 2% Bacto-PPLO serum fraction. Thallium acetate (0.5 g/litre) and penicillin (500 units/ml) were added as inhibitors if necessary, Solid media incorporated 1% Bacto agar. Boidin et al. (1958) obtained abundant growth using Difco PPLO agar or broth enriched with 10% serum and 1% yeast autolysate; thallium acetate (0.25 g/litre) was added and the p H adjusted to 7.9. Harndy et al. (1959) inoculated nostril swabs or portions of lung into liquid media and then, after 2-7 days incubation, a loopful of the liquid culture was streaked onto Difco agar plates containing 2% serum fraction. Mackay et al. (1963) and Mackay and Nisbet (1966) initially used tissue cultures for primary isolation but later used a cell-free medium consisting of Hanks’ balanced salt solution, lactalbumin hydrolysate (5 g/litre), Difco yeast extract (1 g/litre), 10% filtered horse serum, penicillin G (20 units/ml) and streptomycin sulphate (50 pglml). Solid media were prepared by incorporating 1.5% Difco agar, and plates were incubated in an atmosphere of 5% C02 in air. To prepare agglutinating antigen, these authors used the tryptose phosphate broth-yeast extract medium described by Newing and MacLeod (1956) (see p. 230). T h e ovine mycoplasma isolates of Arisoy et al. (1967) were made from mixed populations of sheep and goats using the methods of these authors described under “caprine strains”.

5. Porcine strains M . suipneumoniae ( M . hyopneumoniae) may be isolated using the liquid medium described by Goodwin and Whittlestone (1964) and improved as described by Goodwin et al. (1965 and 1967). T h e recipe is as follows-

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233

Medium (Goodwin and Whittlestone, 1964; Goodwin et al., 1965, 1967)

Either lung broth or Hartley’s broth (autoclaved) 300 ml 50 g/litre lactalbumin hydrolysate in Hanks’ 100 ml balanced salt solution (autoclaved) Pig serum from enzootic-pneumonia-free pigs? 200 ml (inactivated at 56°C for a half hour) Hanks’ balanced salt solution (autoclaved at 400 ml 120°Cfor 10 min) 5 ml Yeast extract+ Penicillin 200 unitslml Thallium acetate? 0.125 g/litre ?Sterilized by Millipore filtration

The lung broth is made by mincing enzootic-pneumonia-free pig lung with twice its weight of water, leaving overnight at 4°C and filtering through double gauze. T h e filtrate is simmered for 15 min, filtered again through gauze and centrifuged (900 g for 20 min) and the supernatant autoclaved (112°C for 30 min) and tested for sterility. The Hartley’s broth is made as described by Cruickshank (1965)’ by digesting ox hearts with trypsin prepared from fresh pig pancreas. T h e enzootic-pneumonia-free pigs are obtained from herds checked in an enzootic pneumonia control scheme (Goodwin and Whittlestone, 1967). T h e Hanks’ salt solution is made as described by Cruickshank (1965) except that the sodium bicarbonate solution is only added if necessary to adjust the pH to 7.0-7.2. T h e yeast extract is prepared by extracting baker’s yeast in its own weight of permutit water brought to p H 4.5 by adding 38% HCI, for 20-30 min at 80°C as described by Herderscheb (1963). T h e indicator is the phenol red present in the Hanks’ salt soIution. T h e final p H of the medium is about 7-6. T h e complete medium is stored at - 25°C. From each batch of medium, samples are tested for sterility and for their efficacy in supporting the growth of M . suipneumoniae by titrating deep-frozen, stock ampoules of the organism and by checking that the organism’s growth is satisfactory during three passages. The liquid medium used by Mark and Switzer (1965) for cultivating this organism was the same as the one described by Goodwin and Whittlestone (1964) but incorporated Difco yeast extract. Mark and Switzer (1966) improved their medium by replacing the Hanks’ salt solution with phosphate buffered saline (Dulbecco and Vogt, 1954) and by including beef heart infusion, and 2004 turkey serum instead of the pig serum. Although Mark and Switzer’s liquid media supported good growth of the organism, the solid media derived from them by the incorporation of agar gave rather poor colony growth in the absence of nurse colonies of a micrococcus.

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R. J. FALLON AND P. WHITTLESTONE

The only satisfactory solid media so far reported for the serial cultivation of colonies of M . suipneumoniae are those described by Goodwin et al. (1965 and 1967); these media have the same recipe as the liquid media previously described (see p. 233) but 1% Oxoid Ionagar No. 2 is autoclaved with the Hanks' solution and then cooled to 56°C before adding the other constituents warmed to the same temperature. M . hyorhinis and M. granularum will grow well in the media described above but will also grow in or on more readily available media. Satisfactory growth of M . hyorhinis occurs in an ox heart infusion medium with 20% chicken serum as described by Switzer (1955). T o prepare this medium, 1 part of fat-free fresh ox heart is minced and infused with 2 parts of distilled water for 12 h at 5°C. T h e infusion is heated in a boiling water bath for 30 min and then filtered through coarse filter paper. T h e pH of the filtrate is adjusted to 7.8 (using 0-1N sodium hydroxide solution). Sodium chloride (5 g/litre) and Bacto haemoglobin (2 g/litre) are added, the infusion is again heated in a boiling water bath for 30 min, and the medium then clarified by filtration. Finally, 20% heat inactivated chicken serum is added and the complete medium sterilized by filtration. If necessary, thallium acetate (0.24 g/litre) and penicillin (2500 units/ml) are added. M. granularum grows in this medium, but growth is considerably improved by adding 0.5% swine gastric m u c h (Ross and Switzer, 1963; Switzer, 1947a). T h e isolation of a number of other mycoplasmas from swine has been reported. Three of the four apparently new swine serotypes reported by Dinter et aE. (1965) have recently been serologically identified by TaylorRobinson and Dinter (1968) as M . laidlawii, Mycoplasma gallinarum and Mycoplasma iners. T h e latter two isolates are usually considered to be avian mycoplasmas. T h e fourth serotype (B3) may well be a new porcine serotype (Taylor-Robinson and Dinter, 1968) but Tully and Razin (1968) have found that its electrophoretic pattern resembles that of M . mycoides var. Capri. T h e mycoplasma isolates reported by Moore et al. (1965, 1966a, 1964b) and Pillai et al. (1967) are not adequately characterized and will not be discussed further.

6. Canine strains T h e isolations of mycoplasmas by Shoetensack (l934,1936a, b) from cases of the virus disease, canine distemper, were made on a meat infusion or meat digest agar with 30% defibrinated blood added. Shoetensack's observation that these strains were more exacting in their growth requirements than M . mycoides var. mycoides was confirmed by Klieneberger (1938, 1962) who used a boiled-blood horse-serum agar.

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X. MYCOPLASMAS

The mycoplasmas cultured by Edward and Fitzgerald (1951a) from the genito-urinary tracts of dogs suffering from infertility, and from the throats of dogs, were isolated using Edward’s selective medium (Edward, 1947a). Certain isolates did not grow in rabbit serum broth, but good growth was obtained when cholesterol (0.1 mg/ml) was added to the medium as a finely dispersed suspension (Edward and Fitzgerald, 195lb).

7 . Feline strains The original isolate of Mycoplasma felis was made using Columbia agar base (BBL) supplemented with 5% horse blood (Cole et al., 1967). T h e other isolations of this species and those of Mycoplasma gateae were made by these authors on 10% horse serum PPLO agar (unspecified) with 1000 units/ml of penicillin G added.

8. Rodent strains The rodent strains of mycoplasmas may be successfully isolated using the liquid and solid media Nos. 44 and 45 (see pp. 226,227).

9. Avian strains Many of the avian strains may be isolated using the horse serum broth medium No. 45 or similar media. The medium used by Dierks et al. (1967) for cultivating all the 19 avian serotypes except Mycoplasma meleagridis (serotype H) and M. synoviae (serotype S ) has the following recipeMedium (Dierks et al., 1967) Difco brain heart infusion broth Yeast autolysate (Albimi Laboratories, New York) Tris Turkey serum 2, 3, 5-triphenyl-2H-tetrazolium chloride (as growth indicator) Distilled water

37 6 5g

0.25 g 100 ml 0.05 g 1 litre

Penicillin and thallium acetate are added as bacterial inhibitors and the pH is adjusted to 8.0 with sodium hydroxide. T h e medium is sterilized by Seitz filtration. T h e solid medium is prepared by adding enough of the broth medium to an autoclaved agar solution (100 g/litre) to give a final agar concentration of 10-12 g/litre. These same authors cultivated M . meleagridis using the VF medium described for the isolation of M . mycoides var. mycoides (see p. 229) except that swine serum was substituted for ox serum and penicillin and thallium acetate were added as bacterial inhibitors, M . synozliae, which

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li. J . I:AI.I.ON AND P. WWI'I"I'LEST0NE

requires nicotinamide adenine dinucleotide (NAD), was cultivated using the medium described by Chalquest and Fabricant (1960) as modified by Chalquest (1962) and Olson et al. (1963). This is a two-phase medium, the agar having the following compositionMedium (Chalquest and Fabricant, 1960;Chalquest, 1962; Olson et al., 1963) 30.6g PPLO agar (Difco) dehydrated Soluble starch 5.0 g Trypticase (Difco) 5.0 g Thallium acetate 0.25 g Phenol red 0.025 g Nicotinamide adenine 0.1 g dinucleotide (NAD) 100 ml Swine serum (inactivated) 1,000,000units Penicillin 1litre Distilled water

The agar is dissolved by heating, and the trypticase, soluble starch, thallium acetate and phenol red added; this mixture is autoclaved (121°C for 15 min), allowed to cool to 56°C before adding the other constituentsswine serum, penicillin and the NAD (beta-diphosphopyridine nucleotide (DPN) powder from Sigma Chemical Co., St. Louis, Missouri, U.S.A.), which is made up in 10 ml quantities, Seitz filtered and kept frozen until used. The liquid overlay, which is similarly prepared (the cysteine hydrochloride is included with the autoclaved constituents), containsLiquid Overlay PPLO broth (Difco) dehydrated, without crystal violet Cysteine HCl Phenol red Thallium acetate NAD Swine serum (inactivated) Penicillin Distilled water

18.9g

0.1 g 0.025 g 0.25 g 0.1 g 100ml 1,OOO,OOO units 1litre

Kerr et al. (1964) found that this medium was also very suitable for the propagation of M . pneumoniae.

10. Other sources Mycoplasmas are frequently found as Contaminants in tissue cultures, especially those of continuous cell lines where repeated subculture exposes the cultures to the hazard of contamination from man, animals or possibly even constituents of the culture medium. The presence of mycoplasmas

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may be recognized with relative ease if they give rise to a cytopathic effect but they may exist in cultures without giving rise to any observable effect until and unless the conditions in the tissue cultures are altered (e.g., by the addition of material for virus isolation studies) so that the equilibrium between mycoplasma and tissue culture is upset. Methods such as staining (Fogh and Fogh, 1964) or detecting activity of L-arginine iminohydrolase (arginine deiminase) (Barile and Schimke, 1963) may be used to detect the presence of mycoplasmas in tissue cultures but for confirmation, isolation on solid medium is necessary. Mycoplasmas from tissue cultures will usually grow well on subculture onto artificial medium. If negative results are obtained, however, liquid, two-phase or semi-solid media should be tried.

C. Conditions of culture for primary isolation Although most mycoplasmas may be isolated under aerobic conditions, a number of species will only grow on primary culture on solid medium under conditions where the concentration of oxygen is reduced and that of carbon dioxide increased. Many workers incubate parallel plates in different atmospheres, e.g., aerobically, aerobically+ 5-10% C02 and in 95% Nz +5% C02. Incubation under anaerobic conditions is particularly important for mycoplasmas of human origin (Mufson et al., 1967). On subculture, nearly all mycoplasmas grow in aerobic conditions. With solid medium the surface of the agar should be moist; plates should not be dried before use and should be incubated in a humid atmosphere in a closed container. The need for special atmospheric conditions is obviated by using twophase, liquid or semi-solid medium in screw cap bottles. Stewart and Chowdray (1968) report favourably on semi-solid medium for isolating mycoplasmas from the respiratory tract. Although it is not so easy to isolate mycoplasmas directly on solid medium, this method has the obvious advantage over fluid or semi-solid medium that overgrowth by bacteria and yeasts may be avoided more easily. Also, on solid medium, mixed mycoplasma cultures may (a) be purified by single colony subculture (see p. 238) or (b) be examined directly by the fluorescent antibody technique so that different species within one culture may be identified (Del Giudice et al., 1967; Stewart, 1967). Cultures should be incubated for at least three weeks before being discarded, especially if the presence of M . pneumniae is suspected. Failure to propagate mycoplasmas which form colonies on primary isoiation is not uncommon (Hayflick and Stanbridge, 1967). Presumably in these instances the inoculated material (e.g., secretions, or infected tissue or an isolate already growing in tissue cultures or liquid medium) provides essential nutrients that are not present in the solid medium in adequate amount. In the case of M . pneumoniue (Chanock eta]., 1962; Hayflick, 1965)

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and M . suipneumoniae (Goodwin et al., 1965) this difficulty was overcome empirically by eventually finding a medium which provided all the nutrients necessary for the serial propagation of these fastidious colonies. This situation, which may well apply to other, as yet unrecognized species, is still a substantial problem because, until serial propagation of a new isolate has been achieved on solid medium, it cannot be asserted that the “colonies” seen on solid medium are truly those of mycoplasmas. There is much need both for the improvement of solid media and also for the discovery of selective inhibitors for different mycoplasmas, as often more than one species may occur in a particular site.

111. PURIFICATION O F CULTURES

A. Cloning methods More than one species of mycoplasma may be present in tissues and secretions submitted for culture. For example, at least 5 species may occur in the oropharynx of man and 4 species in the respiratory tract of the ox. T h e purification of cultures is clearly essential before preparing antigens for making critical antisera and before cultures can be identified. Not surprisingly, it has been found that more than one species of mycoplasma can be present in what appears morphologically to be a single colony (Crawford and Kraybill, 1967; Del Giudice et al., 1967). Purification is usually effected either by plunging a wire into the centre of a single colony and then subculturing or by punching out the colony on its piece of agar with a hypodermic needle or Pasteur pipette while examining the plate under the dissecting microscope; the block of agar with its colony may then be transferred to and streaked on a fresh plate or preferably the colony may be emulsified in broth and serial dilutions up to made before inoculating piates (Ern0 1968). Colonies must be serially cloned at least twice more, to be reasonably certain of eventually obtaining a pure culture. I n studies of the different mycoplasma species occurring in health and disease, the different types of colonies occurring on one plate should be individually selected for subculture along the lines described by Crawford and Kraybill(1967). In this type of work several different media and incubation methods are used and the various mycoplasmas isolated are selected as single colonies and serially cloned. Most workers, however, are interested in the isolation, purification and identification of particular mycoplasmas of known pathogenic significance. T h e problem is much easier if isolations can be made directly onto solid media. For a few mycoplasmas, selective media or cultural methods are known, e.g., for M . pneumoniae (see pp. 227,228) and the human and bovine T-strains (see pp. 228, 231) and thus the problem of purification after

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isolation is minimized. More usually, a pure culture of a particular mycoplasma is obtained from a mixed culture by sub-culturing a colony with the expected morphological appearance. This type of selection is only really practicable when the colonial morphology of the particular mycoplasma is known under the exact cultural conditions being used. When the wanted mycoplasma is more fastidious than the other mycoplasmas that usually accompany it, and/or when it will not grow adequately on solid medium on primary isolation, as occurs with M . suipneumoniae (Goodwin et al., 1968), a pure culture is usually obtained fortuitously. One possible method of overcoming the problem where the likely identity of the unwanted mycoplasma species is known, is either to incorporate its antiserum in the solid medium or, even more simply, to place a disc impregnated with the antiserum on to the medium as for the disc growth-inhibition technique (see p. 221). Growth of the unwanted mycoplasma will usually be inhibited so that the wanted mycoplasma may be cloned from the zone adjoining the antiserum-impregnated disc. In the case of M . hyorhinis, however, this method may not work; for with some strains, colonies of the organism may grow close to the antiserum-impregnated disc. It has been suggested by Hayflick and Stanbridge (1967) that these could be antigenic variants of the species. Some isolates of organisms thought to be mycoplasmas have been made in liquid medium but cannot be grown on solid medium, while others are very difficult to propagate from single colonies and much easier to handle in liquid medium. Thus serial limiting titrations can be used for purification instead of single-colony serial cloning. T h e property of many mycoplasmas to metabolize medium components with a resulting pH change in the medium (see p. 222) can be used to determine the infectivity end point in titrations.

B. Differentiation from bacteria and artefacts 1. L-jorms of bacteria Bacterial L-forms are described elsewhere (see Maxted, this Series). Colonies of L-forms of bacteria however (which closely resemble mycoplasma colonies morphologically) are described here because they may arise from bacteria by the action of various inducing agents, particularly penicillin, which is a common component of mycoplasma media. The main differences between bacterial L-forms and mycoplashas are summarized in Table 11. L-form colonies tend to have a darker centre than mycoplasma colonies, and their peripheral portion is lighter with a coarse lace-like structure. They may reach a size of 0.5 mm after a few days incubation. Under crowded conditions, the minor morphological differences between

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TABLE 11 Differences between Mycoplasmas and bacterial L-forms Mycoplasmas

1. Occur in nature. 2. Require sterols for growth (except M. laidlam? and M. granularurn). 3. Do not revert to bacterial forms if grown in antibiotic-free media. 4. Limited demonstrable metabolic activity.

5. Not related to bacteria genetically. 6. Guanine-cytosine % in DNA bases may be lower than for any known bacteria. 7. Sensitive to lysis by digitonin.

L-forms

1. Usually laboratory artefacts. 2. No absolute requirements for sterols,

3. Unstable L-forms will revert to bacterial form on removal of agent (e.g., antibiotic) used to produce L-form. 4. L-forms will often have similar metabolic activity to parent organism (e.g., enzymic processes such as catalase, coagulase, urease, carbohy. drate fermentation). 5. Genetically indistinguishable from parent organism. 6. Guanine-cytosine % in DNA bases as for parent bacteria and often higher than mycoplasmas. 7. Largely resistant to lysis by digitonin.

the two types of colony may not be so evident; subjective morphological criteria can be very misleading particularly in view of the wide range of colonial morphology seen even with the recognized mycoplasma species. Some experienced workers, however, assert that they can reliably differentiate between mycoplasma and L-form colonies. Suspected mycoplasma colonies should be subcultured without delay onto antibiotic-free medium (as well as serum-free medium), to give an opportunity for the reversion of any L-form colonies to the normal bacterial form, since the risk of producing stable bacterial L-forms increases with passage in the presence of the inducing agent. Morphological differences between rnycoplasmas and L-forms are not yet clearly defined either by light microscopy (Dienes, 1960; Kagan, 1967) or by electron microscopy (Dienes and Bullivant, 1967; Cole, 1967). A study of the cell membrane by Razin (1967) showed that mycoplasma membranes seem to be more stable and elastic than membranes of bacterial protoplasts. This may be related to the cholesterol content of the mycoplasma membrane. It has also been shown (Smith and Rothblat, 1960, 1962; Razin and Argaman, 1963) that mycoplasmas are sensitive to Iysis by digitonin, whereas L-forms, spheroplasts and protoplasts are resistant to such lysis. The sensitivity of mycoplasmas to digitonin probably indicates that cholesterol forms an integral part of the cell membrane (Razin et al.,

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1963). Electrophoretic analysis of cell proteins (see p. 251) also promises to be of value in distinguishing between mycoplasmas and L-forms (Razin and Rottem, 1967). ‘l’he percentages of guanine + cytosine (G + C) in the DNA bases of some strains of mycoplasmas are among the lowest reported for any micro-organisms and lower than any known for eubacteria (Neimark and Pkne, 1965). McGee et al. (1967) who studied the G C base ratios of mycoplasmas and bacteria! L-forms, came to the following conclusion : “The possibility that some of the mycoplasma are the L-forms of a limited number of bacteria cannot be excluded, but there seems to be little evidence to support such a hypothesis. The view that all of the mycoplasma are the L-forms of bacteria seems untenable on the basis of presently available evidence.”

+

2, Bacteria On solid media, very small bacterial colonies may simulate mycoplasma colonies. Opaque colonies without a central nipple usually prove to be bacterial. T h e centres of mycoplasma colonies are usually embedded in the agar surface, whereas nearly all bacterial colonies may be rubbed off the agar surface. This is best seen by examining the plate at low magnification ( x 25) while stroking the colony with a bacteriological loop. Touch preparations of bacterial colonies fixed in methyl alcohol for 2-5 min and stained with Giemsa stain (1 in 20 for 3 h) show deeply-stained organisms with a distinct morphology related to the rigid bacterial cell wall ; mycoplasma colonies, by this method, are composed of lightly-stained, very pleomorphic elements with a range of sizes down to 0.2 ,um (see p. 253 and Fig. 11). By Gram’s staining method, mycoplasma cells are extremely difficult to see, being Gram-negative and very delicate. The staining method of Dienes (1939) is useful in differentiating between bacterial and mycoplasma colonies. T h e stain, as detailed by Hayflick (1965), is prepared by dissolving 2.5 g methylene blue, 1.2 g azure 11, 10 g maltose and 0.25 g of sodium carbonate in 100 ml distilled water. The stain may either be dried on a cover-slip which is then applied to the surface of a block of agar bearing colonies, or the stain may be applied with a cotton wool swab directly onto the surface of an agar culture. Mycoplasma colonies stain distinctly with dense blue centres and do not decolourize the stain, whereas a Iiving bacterial colony will decolourize th: stain after about 30 min (Hayflick, 1965). It may be necessary to differentiate mycoplasmas from bacteria in liquid media particularly if the mycoplasma species in question cannot be cultivated on solid medium (see p. 256 for a description of the technique). Such centrifuged deposits mzy, in special cases, be sectioned and examined by I1

electron microscopy iooking espccially for thc characteristic triple-layered cell membrane (see p. 258 and Fig. 15).

3. Artefacts T h e following artefacts are comnonly confused with mycoplasma colonies by less experienced woriiers. (a) Air bubbles. I n the preparation of some batches of medium, minute air bubbles are trapped in the agar, and may appear to have nipple-like centres and may reflect the light rather like mycoplasma colonies (see Fig. 7). They can be differentiated morphologically from mycoplasma colonies

FIG.7. Air bubbles trapped beneath agar surface and simulating mycoplasma colonies (compare with Fig. 2). Unstained x 24.

fairly readily with a little experience; they are beneath the agar surface, they are also often present away from the inoculated area or on uninoculated plates from the same batch, they do not enlarge on incubation and cannot be subcultured, they do not take Dienes’ stain, and do not show mycoplasma-type elements in Giemsa-stained touch preparations. Although bubble formation can be prevented by flaming the agar surface as the plates are prepared and before the agar has solidified, care must be taken not to dry the agar surface, as this is detrimental to mycoplasma growth. Some workers prefer not to flame the surface because of this risk. (b) Tissue cells. Agar plates inoculated with tissue suspensions or tissueculture material can be quite difficult to interpret. A single cell with its central nucleus can closely resemble a very small mycoplasma colony with its central nipple. Groups of tissue cells may closely resemble mycoplasma colonies. They may be differentiated from rnycoplasma colonies because they do not enlarge on incubation and do not stain as do mycoplasmas; although they do not subculture they may be mechanically transferred to further solid medium. When solid media have been initially inoculated with

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material likely to cause confusion, it is good practice to examine the plates before incubation, to assess the situation. Aggregates of platelets are particularly liable to be confused with mycoplasma colonies (Ryschenkow et al., 1967); after inoculation of platelet-rich plasma onto agar, the platelets progressively coalesce over a period of 4-24 h. This aggregation occurs more rapidly at 37°C than 25”C, and the resultant structures closely resemble mycoplasma colonies morphologically. They differ from mycoplasma colonies in that they do not show mycoplasmatype elements in stained preparations and they cannot be subcultured. (c) Fluid droplets. During the normal incubation of plates under humid conditions, minute droplets of fluid sometimes form on the agar surface. They are usually not confined to the inoculated area of the plate, they do not contain mycoplasma-type elements in stained preparations and cannot be subcultured. (d) Pseudocolonies. To new workers in the mycoplasma field, pseudocolonies present the greatest problem of all artefacts. For they often have a darker central area, rather like the centre of a mycoplasma colony, they “grow” slowly on incubation reaching maximum size in about a week or more, and they may be subcultured in series on agar. Various types of pseudocolony are illustrated in Figs 8, 9 and 10; for illustrations of other types of

FIG.8. Pseudocolonies (arrowed) on agar plate, with colonies of Mycoplasma plreumoniue (marked C). Unstained x 100.

pseudocolony see Brown et al. (1940) and Hayflick (1965). Pseudocolonies form on agar media having a high serum content, and they consist of calcium and magnesium soaps which crystallize in localized areas on the agar surface. ‘I‘hcy usually rcach a diameter of about 150 pm. When subcultured they

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break up and the fragments presumably re-initiate crystal growth in the inoculated areas. Their formation can be induced by streaking the surface of sterile agar with a sterile block of agar, loop or pipette.

FIG.9. Pseudocolonies on agar plate. Unstained x 160.

FIG.10. Six pseudocolonies on agar plate. Unstained x 64.

Pseudocolonies can be made to appear on media containing high enough concentrations of merthiolate or formalin to prevent growth of living organisms (Brown et al., 1940). They will also appear on inoculated media that have been exposed to a dose of ultraviolet radiation sufficient to prevent the formation of mycoplasma colonies (Furness el al., 1968a). With experience, pseudocolonies can be differentiated from mycoplasma colonies morphologically, and stained preparations of the pseudocolonies do not contain mycoplasma-type elements.

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C. Confirmation of identity as a mycoplasma At the present time the mycoplasmas are not identifiable as such on the basis of any common fundamental property. A distinctive biochemical characteristic shared by the vast majority of the mycoplasmas is a requirement for sterols but this characteristic cannot be used for routine identification of isolates as mycoplasmas. In practice an isolate is confirmed as a mycoplasma on the basis of some or all of the following characteristics(1) T h e formation of minute characteristic colonies on enriched agar medium, with the centre of the colony burrowing into the medium (see p. 214). (2) T h e resistance of cultures to penicillin and (with the possible exception of T-strains) to thallium acetate (0.125 gllitre). (3) T h e sensitivity of cultures to tetracyclines. (4) T h e failure of all but saprophytic and T-strain mycoplasmas to grow in serum-free medium. (5) T h e failure to revert to a bacterial organism when passed in the absence of penicillin. (6) T h e ability of colonies on solid medium to take up and retain Dienes’ stain (see p. 241). (7) T h e inhibition of growth, both in fluid and on solid medium by specific hyperimmune antibody (see pp. 221,222). (8) T h e presence of delicate, pleomorphic organisms with elementary bodies about 125-200 nm in diameter. (9) By electron microscopy, the presence of a triple-layered cell membrane, the middle layer being less dense than the inner and outer layers (seep. 258 and Fig. 15). IV. GROWTH OF MYCOPLASMAS

A. Characteristics of growth 1. Turbidity In liquid media, mycoplasmas usually produce either no turbidity or a faint turbidity when viewed by transmitted light. High-titre cultures,particularly in large volumes, may be quite turbid when examined by side light against a dark background. Kelton (1960) observed turbidity with several strains of mycoplasmas at the peak titres only, when the viable counts were in excess of lo7 or 108 units per ml. Some of the well-adapted strains of mycoplasmas readily produce turbidity in liquid medium, e.g., M . mycoides var. mycoides and the FH strain of M. pneumoniae which also produces macroscopically visible spherical colonies (Kenny and Grayston, 1965).

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Turbidity may sometimes be due to changes occurring in the medium during incubation.

2. Titres in liquid medium Maximum titres of viable units in liquid medium vary with different inycoplasmas and media from about 106 or 107 colony forming unitslml for the T-strain mycoplasmas (Taylor-Robinson and Purcell, 1966; Shepard, 1967) to l o 9 colony forming units/ml for adapted strains of M. pneumoniae (Somerson et al., 1967) and M . mycoides var. mycoides (Nasri, 1967b). Laboratory-adapted strains of mycoplasmas usually reach maximum titre after 1-4 days incubation but freshly isolated strains may take considerably longer; T-strain mycoplasmas take only 16-18 h. These titres are maintained for a day or so or only for a few hours (e.g., with T-strain mycoplasmas) due to the elaboration of toxic substances, e.g., acid or alkali, from the substrate by the organisms. As with bacteria, the total organism count increases after the peak of the viable titre is reached. Sometimes better yields are obtained by shaking or rotating cultures. 3. Growth curves All the strains studied by Kelton (1960) showed growth curves with the characteristics of bacterial growth curves. In the logarithmic phase the organisms increased in numbers in a geometrical progression with a common ratio of two. Other workers (e.g., Low and Eaton, 1965; Taylor-Robinson and Purcell, 1966; Purcell et al., 1966a) have published growth curves showing a log phase of growth. Kelton interpreted his data as favouring binary fission as the mode of reproduction of mycoplasmas; there is, however, no general agreement on this matter (see p. 213). T h e morphological characteristics of mycoplasmas in liquid and on solid medium are described on pp. 256 and 214.

4. Growth on glass and plastic surfaces Several species or strains of mycoplasmas, when grown in liquid medium, adhere to glass or plastic surfaces and form colonies or sheets of cells, e.g., M . pnmmoniae (Somerson et al., 1967), M . suipneumoniae (Goodwin and Whittlestone, 1966), M. gallisepticum, M . pulmonis, M . hominis, Mycoplasma orale types 1 and 2 and Mycoplusma salivarium (Taylor-Robinson and Manchee, 1967). T h e sheets of cells may be seen more clearly after staining. Taylor-Robinson and Manchee (1967) also noted that mycoplasmas would adhere to plastic Petri dishes. (See also pp. 250 and 254.)

5. Growth on solid medium On primary isolation, mycoplasma colonies may take many days to appear

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on solid niecfium. ’l’his property varies from species to species and also with the origin of the inoculum, e.g., subcultures from tissue cultures or embryo-

nated hens’ eggs will usually grow more rapidly than those from other sources; on the other hand, sometimes a mycoplasma adapted to another culture system, e.g., liquid medium or experimental animal, may grow with great difficulty on solid medium. After several passages on solid medium, colonies of most species appear within 2-4 days of inoculation. As with bacteria, increase in colony size continues for many days under good conditions, but once colony growth ceases the viability of the organism falls off rapidly. Agar plates must be incubated under suitable atmospheric conditions (see p. 237).

B. Measurement of growth T h e techniques that are commonly employed for quantitative estimations of bacterial growth (e.g., turbidity, packed-cell volume and dry weight) are unsuitable for mycoplasmas because of the small size and the low yields of these organisms. There are four main methods that are suitable for general laboratory use for the quantitative measurement of mycoplasma growth. They are(1) the measurement of tubidities of concentrated cultures; (2) the measurement of colony diameters; (3) the measurement of viable cells either by colony counting or by determination of the number of metabolic units; and (4) the counting of sedimented cells. 1. Turbidities T h e method employed by Smith (1956) is as followsBroth cultures are centrifuged under sterile conditions at 13,000 rpm in the Servall angle centrifuge (ca. 15,000 gfor 5 min); this results in recovery of about 90y0 of the mycoplasma cells but this yield can be improved to about 990/b by increasing the speed to 20,000 rpm (ca. 40,000 g). T h e supernatant is decanted, and the sedimented mycoplasma cells resuspended in physiological saline to 1/40 of the original culture volume. T h e turbidity of the suspension is determined in a photoelectric colorimetcr with a 420 nm filter. Great care must be taken not to disturb the sediment while decanting the supernatant, otherwise loss of sedimented mycoplasma cells may occur. , T h e relationship between turbidity and numbers of viable cells is determined by making high dilutions of suspensions of different turbidities, plating out and making colony counts after incubation (see below).

2. Colony diameters Smith (1956) showed that, provided the dilution of inoculum is sufficient to prevent crowding of colonies, colony size at diflerent lengths of incubation can be used as a quantitative measure of the nutritive value of different media. With mycoplasma cultures that normally yieId fairly uniform colonies, the mean diameters of 10 colonies will give a meaningful result; if a culture normally produces a big range of colony sizes, the number of colonies measured must be increased. Colonies are conveniently measured using an ocular micrometer scale (the conversion factor in micrometres being calculated for the microscope being used).

3, Viable cell count (a) solid media colony counts. 'The method used by Smith (1956) is as fo~~owsExactly 0.01 rnl aliquots are plated onto solid medium, in duplicate, without spreading, so that the area inoculated is completely visible in one microscope field. (This eliminates the errors occurring, (i) if organisms adhere to a spreader, and (ii) when it is necessary to count more than one field.) After appropriate incubation, the plates are flooded with dilute Dienes' stain (see p. 241) and counts performed. To ensure that the 0.01 ml plate inoculum results in a countable number of colonies, several different dilutions (e.g., 10-4, 10-5 and 10-6) may have to be used. This technique is similar to that of Miles and Misra (1938) (using Pasteur pipettes calibrated to deliver 50 drops per ml), which also may be applied to the enumeration of viable mycoplasmas in a culture. It is convenient to record cultures photographically using an automatic camera as described on p. 253. The negatives can then be printed at any desired magnification for counting colonies. T h e same method is also useful for measurement of colony diameters. (b) Titrations of metabolic units. One group of mycoplasmas, including, e.g., M . pneumoniae, M . suipneumoniae and M. gallisepticum, metabolizes carbohydrates, with acid formation ;many of the non-acid producing species release ammonia from arginine or urea (see p. 222). T h e resultant change in pH of the liquid medium is detected with suitable indicators. Although these two properties have not been successfully used to quantitate mycoplasma growth directly, they are used widely to measure indirectly the titre of viable units in cultures. T h e method can be applied either to the fluid phase of broth cultures or to sheets of mycoplasma cells, e.g., M . pneummiae harvested from a glass surface (Somerson et al., 1967).Titrations are made in duplicate or triplicate in the appropriate medium and then incubated. T h e highest dilution

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which changes colour is presumed to contain one viable unit; thus the number of viable units in the original culture can be calculated.

4. Total cell count Clark (1965) has described a method of enumerating the total cells present in a culture by the technique of sedimentation counting. Here the cells are centrifuged onto a cover-slip which is then drained and fixed in 0 ~ 0 4 The cells are stained with crystal violet and counted under the oil-immersion lens.

V. HANDLING OF MYCOPLASMAS A. Subculture 1. From liquid medium The methods of subculture of mycoplasmas in liquid media and from liquid to solid media are the same as those used for bacteria and the hazards and precautions (see p. 252) are essentially the same.

2. From solid medium Although, like bacteria, mycoplasmas may be subcultured from solid medium by scraping a wire loop across the surface of the growth and inoculating a fresh plate, this method is not reliable. T h e usual method of subculture is to cut out with a flamed scalpel, a 1 cm2 block of agar bearing colonies. The area may be marked off previously with a marking pencil, or the scalpel can be used while examining the plate under the dissecting microscope. T h e scalpel is slipped under the block of agar which is transferred face downwards to the surface of a fresh plate of culture medium. This block is then pushed over the surface of the medium with the scalpel, and left near the edge of the plate. After several days, colonies will appear on the surface of the fresh plate. Fluid medium may be inoculated either by transferring a block of agar bearing colonies as above into the broth or by plunging a wire into a colony (while examining the plate under the dissecting microscope) and then inoculating the broth with the wire.

B. Cultivation and concentration for serological studies Mycoplasmas required for serological studies are usually most conveniently grown in liquid medium. If the organisms are to be used in techniques where the presence of residual medium constituents might cause nonspecific reactions (e.g., in precipitation in agar gel or in immunofluorescent studies), the mycoplasma should preferably be grown in a medium that is

.

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non-antigenic for the laboratory animal being used for the production of antiserum. Mycoplasmas grown in the usual media, even after repeated washings, still have associated antigenic medium constituents. Thus, if antisera are to be prepared in rabbits, the mycoplasma should idealiy be adapted to a medium consisting of a rabbit meat infusion broth (TaylorRobinson et al., 1963) or the dialysable fraction of some other mycoplasma medium, with added rabbit serum together with any other enrichments that are non-antigenic for the rabbit. As rabbit serum is a poor source of sterols, 0.02% cholesterol should be added to the medium. If it is not practicable to grow the mycoplasma satisfactorily in such a medium, or if residual medium constituents do not interfere with the serological test being used (e.g., growth inhibition on solid medium or metabolic inhibition in liquid medium) the mycoplasma is grown in any convenient medium. The eventual resulting antisera may have to be absorbed appropriately with the medium’s antigenic constituents before use. For maximum yield of organism, cultures are usually shaken and incubation continued beyond the peak of viable titre (see p. 246). Larger volumes of culture may be obtained using continuous culture as described by Brighton et al. (1967), who obtained a daily yield of 1.5 to 2.0 litres of culture containing 105 colony forming units/ml. Cultures are centrifuged at 27,000 g for 30 min (i.e., particles as small as 100 nm will usually be deposited) and the resultant deposit is washed several times by resuspending in buffered saline and recentrifuging. T h e final suspension usually represents a 20-100 fold concentration of the original culture. A simplified method for preparing M. pneumoniae antigen has been described by Somerson et al. (1967); the method is based on the fact that M . pneumoniae attaches to glass surfaces and this eliminates the necessity of centrifuging large volumes of culture acd has the additional advantage that for complement fixation, the antigen is free from anticomplementary effects without additional treatment. T h e method is as follows-2 litre Povitsky bottles (similar to Thompson’s culture flasks obtainable from Camlab, Cambridge) containing 500 ml of broth medium are inoculated with 10’0 organisms. After 5 days’ incubation, the layer of mycoplasmas attached to the glass is washed three times with phosphate-buffered saline and then removed from the glass, either with a rubber policeman or by trypsin treatment, and resuspended in 20 ml of saline. For use as antigens in gel diffusion or complement fixation tests, suspensions are usually disrupted by sonication or by freezing and thawing for 10 cycles. Kenny and Grayston (1965) have described the preparation of a complement fixing lipid antigen from M . pneumoniae, which is low in anticomplementary activity and is of reproducible high quality.

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C. Concentration for biochemical studies . ’I’he concentration procedures described above may be suitable starting points for some biochemical studies but, in view of the wide variety of techniques available and the many facets of the biochemistry of these organisms, the original refercnces shodd be consulted. (See the review by Smith, 1964, and Hayflick, 1967, for recent relevant papers.) One aspect of the biochemistry of the mycoplasmas which is of interest and which may become of increasing importance in the classification of these organisms is the patterns produced by their proteins when mycoplasma extracts are subjected to electrophoresis. T h e applications of this technique to the analysis of cell disintegrates is described by Sargent and by Cooksey (this Series, Vol. 5), but the specific applications to mycoplasmas are outlined here. Fowler et 01. (1963) studied the patterns produced by the proteins of ultrasonically disintegrated mycoplasmas and L-forms of bacteria when these extracts were subjected to electrophoresis in starch gel. Different patterns were obtained with different species but the protein separation was not always clearly discrete. Further work was carried out using microimmunoelectrophoresis (Fowler et al., 1967) but the results were difficult to interpret and therefore polyacrylamide column “disc” electrophoresis was tried with much better results. As would be expected, protein bands were much more discrete and well defined than with starch gel. Rottem and Razin (1967) used polyacrylamide gel to study the patterns obtained when mycoplasma membrane proteins were subjected to electrophoresis. As separation of cell membranes is tedious, involving cell fractionation, Razin and Rottem (1967) and Haas et al. (1968) examined the electrophoretic patterns of whole cells with very encouraging results. T h e mycoplasmas were grown in 100-200 ml of Hayflick’s mycoplasma broth for 24-48 h at 37°C and were harvested by centrifugation at 13,000 g for 10 or 20 min. Cells were washed and suspended in 0.25 M saline, dissolved in a mixture of phenol, acetic acid and water (2 : 1 : 0.5 w/v/v) centrifuged at 30,000g for 15 min to remove insoluble material and were then applied to a polyacrylamide gel in 6 x 100 mm glass tubes. Electrophoresis was carried out at 5 mA per tube for 2 h. Gels were stained with naphthalene black, and were electrolytically destained. Clear patterns were observed which did not appear to be affected by different growth conditions or by thc age of the cells. l’he electrophoretic patterns were species specific and thc identification of strains correlatcd well with the growth-inhibition test. Hazin and Iiottem (1967) suggest that the simplicity of the method may havc wide appeal and may be cheaper than techniques depending on expensive antisera. This development is clearly one of considerable importance not only for identifying and

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classifying mycoplasmas but also for differentiating them from bacterial L-forms.

D. Hazards and precautions It is easy to cross-contaminate mycoplasma cultures and the literature contains many instances of this (see Edward and Freundt, 1965). Not only may established cultures be contaminated but, as already noted, clinical specimens may contain several strains of mycoplasmas. For these reasons it is important to ensure that a culture in fact contains only the organism which it is thought to contain. Before making antigen for antiserum production, cultures must be cloned (see p. 238) and subcultures should be stored (see pp. 259-260) for reference purposes. Serial passage should be avoided as far as possible and cultures should be checked against specific antisera before any critical work is carried out. These precautions are even more essential with mycoplasmas than with bacteria because of the fact that colonial appearance is of little help in establishing that a culture is contaminated and also by the fact that one colony may contain more than one species of mycoplasma. T h e literature on rnycoplasmas is often unsatisfactory because many workers have not purified or characterized their isolates adequately. However, a sub-committee on the nomenclature of the Mycoplasmatales has recently been formed (Edward et al., 1967) and has issued a plea for adequate serological characterization, including comparisons with previously recognized species, before any isolate is claimed to be a new species; such new mycoplasma species should be deposited with a National Type Culture Collection. A Mycoplasma Reference Laboratory has been established in Britain at the Central Public Health Laboratory, Colindale, London. International WHO Mycoplasma Reference Centres have also recently been established at the National Institutes of Health, Bethesda, U.S.A. and at the University of Aarhus, Denmark. As a result of these measures, type cultures for reference should become increasingly available.

E, Microscopy 1. Mycoplama colonies (a) Morphology of colonies on solid medium. For best results the medium should be transparent and free from particulate matter visible at low magnifications, and the Petri dishes should be free from scratches; disposable non-toxic plastic Petri dishes are thus ideal. Conventionally, colonies are examined at low magnification (i.e., x 25 to x 100) using a stereoscopic microscope with an external light source reflected substage by an adjustable plano-convex mirror. With the light directed obliquely, colonies can be

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seen extremely well, but it is often difficult to co-ordinate the rather different images in the two eyepieces. This difficulty can be overcome by using a modified Leitz Ortholux microscope which has additional advantages. The substage condenser is removed and a Laborlux plano-convex mirror is slotted into the base of the Ortholux and reflects light from a 6 V, 2.5 A external light source controlled through the usual transformer. Piano objectives ( x 1, x 2.5, x 4 and x 10) are used and an Orthomat automatic camera is fixed to the photo tube. The system gives an adjustable light angle and brightness, a mechanical stage for systematically searching plates, a flat field, a big enough range of magnifications for all types of colonies, the opportunity for spectacle wearers to use high point eyepieces and the automatic camera for objective recording of results. (For photographs of colonies and pseudocolonies taken with this equipment see Figs. 1, 2, 3, 4, 9 and 10.)The appearance of mycoplasma colonies is described on p. 214. It is possible, but very difficult, to observe elementary bodies germinating on the agar surface using an oil immersion objective (Freundt, 1958). A much more revealing method was devised by Brskov (1927, 1942) and used by Freundt (1958,1960) to study the elements present in solid-medium colonies of M . mycoides var. mycoides. This technique is as follows: select young cultures with colonies of a uniformly small type and cut out a square of agar. On the surface of this block put a few cotton-wool fibres and a drop of broth and cover with a cover-slip. Incubate for 15-30 min and then examine with the phase-contrast oil-immersion lens. Fairly long filaments can be seen emerging from the edge of the colonies, particularly in the areas where a cotton fibre crosses a colony. Sometimes the filaments consist of chains of elementary bodies (i.e., very like those observed in liquid-medium cultures of M . mycoides var. mycoides). The elements composing colonies can be examined in stained preparations relatively easily as follows: cut out a square of agar on which a few colonies are growing and rest it face uppermost on a piece of filter paper. With a glass microscope slide make a very light touch on to the agar surface, without smudging the colonies. Allow the preparation to air dry, fix in methyl alcohol for 2 min, stain for 3 h with Gurr’s improved Giemsa stain diluted 1 : 10 in phosphate-citrate buffer pH 7.2 (a mixture of 9.1 ml of 0.1 M citric acid solution, 40.9 ml of 0.2 M dibasic sodium phosphate and 50 ml Elga water). Wash in 2 changes of similar buffer, dry, wash in acetone for 10 sec and dry. T h e wide variety of morphological structures from elementary bodies ca. 0.2 /cm diameter to much larger structures, is shown in Fig. 11, For other photographs of the structures in rnycoplasma colonies stained with Ciemsa, or examined by phasc-contrast or dark-field microscopy see Klieneberger-Nobel(1962). Whole colonies may also be fixed to the surface of glass slides using the tcchnique of Clark et al. (1961): a block of agar

bearing colonies is cut from the platc and is placed on a clean glass slide SO that the colonies are in contact with the glass. l'he slide is inverted and placed at an angle of about 45" into distilled water at 80°C. The water temperature is raised rapidly until the agar melts and flows and falls off the slide. The slide is removed, rinsed gently and air dried. Such preparations may be stained either with Giemsa or by the fluorescent-antibody technique for specific identification (Stewart, 1967).

FIG.11. Touch preparation of an avian mycoplasma colony showing a wide variety of morphological structures including elementary bodies. Giemsa x 960.

(b) Morphology of organisms in colonies in liquid medium. A number of mycoplasmas will adhere to a glass surface under liquid medium (see p. 246). The morphology of M . suipneumoniae in colonies forming under such conditions has been described by Goodwin and Whittlestone (1966) who used the following technique-the mycoplasma is grown in 2 ml of liquid medium in test tubes containing a cover-slip. T h e tubes are incubated in a tissue-culture roller drum until the pH falls to 6.9-7.0. Cover-slips are then removed, washed in buffered saline, fixed and stained in Giemsa as described on p. 253 for organisms from solid-medium colonies, dried, passed quickly through two changes of a 501.50 mixture of acetone-xylol and two changes of xylol and then finally mounted on microscope slides with DePeX (George T. Gurr Ltd., London). The larger colonies can be seen with the low power of the microscope, but in these larger colonies the organisms are very crowded and few details can be seen. Small colonies, when examined under oil immersion are seen to be composed of a wide variety of morphological forms-rings and bipolars are common in some, wht:reas in others the predominant forms are cocci (about 0.3 p m diameter) which are strung on fine branching filaments. Larger globular bodies are also seen. One of these colonies is illustrated in Fig. 12.

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FIG.12. Detail of a mycoplasma colony growing on glass under liquid medium, showing coccal forms strung on branching filaments and one large globular body. May-Grunwald Giemsa x 2400. (From B Y . J .exp. Path, 47,520 (1966).)

2. Mycoplasma cells in liquid medium (a) Dark-field microscopy. T h e morphology and life cycles of M . mycoides var. mycoides were studied by dark-ground illumination by Turner (1935). In this paper he produced a remarkable series of photographs by minute attention to detail. In summary, Turner’s technique is as followsT h e organism is cultured in the medium described on p. 229. Scrupulous attention is paid to optical accuracy and cleanliness of slides and coverslips. A very small drop of culture is placed in the centre of the slide, which is then gently lowered on to a cover-slip on a pad of filter paper. T h e preparation is pressed firmly on the filter paper to remove excess fluid and the edges are sealed with paraffin wax. For photography the preparation must be so thin that the organisms do not move with Brownian movement; a very small drop of culture is used and the preparation rolled with a glass tube to squeeze out all excess fluid. Turner was able to show the great pleomorphism of the mycoplasmas ; the very delicate long branching filaments and the elementary particles could be seen particularly well by this method.

(b) Phase-contrast microscopy. T h e examination of unfixed mycoplasma cells in liquid medium by phase-contrast microscopy, as used by Razin and Cosenza (1966) and Razin et al. (1967) to study the morphology of various mycoplasma strains, minimizes artefact formation. Small drops of culture were covered with a cover-slip and the organisms photographed as soon as they settled onto the glass slide. Mycoplasmas photographed in this way showed an appearance corresponding closely with that of the organism moving freely in liquid medium. I3y this technique, these authors were able

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to confirm the earlier suggestion of Freundt (1958) that the ability to grow in filaments is a general property of mycoplasmas. Filament formation, however, only became manifest at an early phase of growth and in a growth medium with an adequate and balanced supply of cholesterol and longchain fatty acids, notably oleic, linoleic, linolenic and arachidonic acids. (c) Stained preparations. A simple technique for detecting mycoplasmas in liquid media is as follows. Mark the centre of a very clean glass slide with a small ring (2-3 rnm diameter) using a diamond pencil. Put a small drop of mycoplasma culture within this ring and allow it to air dry. Fix in methyl alcohol for 2 min and stain with Giemsa as described on p. 253. Examine unmounted, under the oil immersion, using the diamond ring for orientation and focussing. If cultures are not of high enough titre for the organisms to be detectable by this direct method, a 2 ml aliquot is centrifuged at 25,000 g for 25 min, the supernatant removed and a smear made of the deposit, which may not be visible. A variety of morphological forms may be detected depending on the strain of mycoplasma and its method of culture. By the above method, M . suipneumoniae is seen mainly as ring and bipolar organisms (see Fig. 13); it has a similar morphology in touch preparations taken from the natural disease (see Fig. 5) but looks very different when grown attached to a glass surface (see Fig. 12).

FIG.13. Smear of centrifuged deposit from a broth culture of mycoplasma x 960.

T h e morphology of mycoplasmas in liquid medium may also be studied by the method used for sedimentation counting described by Clark (1965) (see p. 249). T h e methods he describes are suitable both for light and electron microscopy.

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(d) Electron microscopy. Although it is beyond the scope of this chapter to present a comprehensive summary of the literature on the electron microscopy of mycoplasmas, this aspect is mentioned briefly because it relates to the identification of an isolate as a mycoplasma, and its differentiation from other structures. Not surprisingly, in the various electron-microscopy studies, an even wider variety of s t ~ l c t u r e shas been identified than by light microscopy. Even within the same species there are wide differences, probably due to the diffcrent methods of preparation and study or to the use of different strains of the organism. For example, with 111. gallisepticum, Chu and Horne (1967) showed the presence of surface projections thought probably to be the haemagglutinin, whereas Morowitz and Maniloff (1966) and Maniloff and Morowitz (1967) did not detect such projections but described the detailed structure of the “tear-drop-shaped” cell and its relationship to the binary fission of the organism. The organisms may be sedimented as described by Clark (1965), Anderson and Barile (1965) or Kim et al. (4966) and then fixed with a suitable agent (e.g., 0~04, glutaraldehyde, phosphotungstate or formalin vapour) and examined either as shadowed or negatively-stained whole organisms or as ultra-thin sections. Although as noted above, different techniques may give different results it is interesting to note the differences

Flu. 14. Section of centrifuged pellet of Mycoplasma siiipnezrmoniae showing a wide variety of morphological forms. (Whittlcstone et al., 1969) x 3600.

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in appearance of different strains of mycoplasmas when prepared in an identical fashion by one group of workers (e.g., Clyde and Kim, 1967). I n ultra-thin sections, mycoplasmas show a characteristic triple-layered cell membrane, about 70 A thick, the middle layer being less electron dense than the inner and outer layers (see Anderson and Barile, 1965). The structure of M . suipneumoniae as seen by this method is illustrated in Figs. 14and 15.

FIG.15. Section of centrifuged pellet of Mycoplasm suipneumoniae. The small particles are 100-150 nm diameter. Note the triple-layered membrane of the mycoplasma cells and the surface projections. (Whittlestone et al., 1969) x 48,000.

VI. MAINTENANCE OF MYCOPLASMA CULTURES A. Serial pass@ge Mycoplasmas may, like bacteria, be propagated by serial passage either in fluid or on solid medium as described on p. 249. With most strains this is readily accomplished once the critical step from primary to secondary culture has been made. However, T-strain mycoplasmas tend to die rapidly so that they must be subcultured at frequent intervals-no longer than 48 h. Most other species survive for up to a week in fluid medium at 37"C, but M. pneumoniae will survive for even longer than this, and one strain

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of M. orale type 1 (N-1) was found to survive in broth at 37°C for 90 days (Hayflick and Koprowski, 1965). Again, most species will survive for at least a week on solid medium at 37°C and even longer in two-phase medium. Unless there are special reasons (e.g., no means of storage of cultures or a constant requirement for actively-growing cultures) mycoplasma strains should not be maintained for any length of time by serial passage because of the very real danger of contamination of the culture with other mycoplasmas (or other organisms).

B. Low-temperature storage Mycoplasmas, either in fluid medium or as blocks of agar bearingcolonies, may be kept at - 60°C to - 75°C for well over 12 months, sometimes for several years. At higher temperatures, viability is lost more rapidly, but even at -25°C cultures will survive for several months. Fluid cultures should be grown to near the end of the log phase of growth or until a pH change of half a unit has occurred and then frozen to - 60°C to - 75°C. Cultures may be conveniently stored in 1 ml amounts in ;t dram (2 ml) screw cap vials (Flow laboratories, Irvine, Ayrshire, Scotland), which take up a minimum of storage space, In this way cultures of tested titre are instantly available, e.g., for metabolic-inhibition tests or for testing the efficacy of new media. The medium for storage may be either normal growth medium or one designed specially for the purpose. Kelton (1964) investigated the storage of 26 strains of mycoplasmas (19 avian, 2 human, 3 canine and one saprophytic strain) in the following mediumMedium (Kelton, 1960) Heart infusion broth (Difco) Yeast extract (Difco) Proteose peptone No. 3 (Difco) pH 7.9

1 Iitre 10 €? 10 6

The medium was autoclaved at 121°C for 15 min, and before use 100 ml horse serum was added. 48-72 h broth cultures were stored in 0.5 ml amounts at various temperatures. Cultures stored at - 65°C showed little decrease in viable numbers after 12 months storage (compared with the one-month sample); at -26°C all cultures survived for 10 months but one strain was non viable at 11 and 12 months. At 5°C all cultures survived for 3 weeks and some lived much longer. For T-strain mycoplasmas, Shepard (1967) recommended the following storage broth-

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Storage broth (Shepard, 1967) Tryptic digest broth (see Shepard 1956) (pH 7.6) 78 ml 20 ml Sterile horse serum (pH 7 . 6 ) 1 ml Yeast supplement “C” (Difco) 1ml Potassium penicillin G 100,000units/ml

T h e strain is cloned and then incubated in growth medium, e.g., A2 medium (see p. 228) for 16 h. T h e broth is then centrifuged at 27,000 g for 30 min. T h e pellet is resuspended in one tenth of the original volume of broth using the above storage medium at pH 7.6 and is stored at -60°C to --85”C. Alternatively, the concentrated suspension can be well preserved by lyophilization.

C. Freeze-drying Long term storage of mycoplasma strains is best effected by freezedrying the organisms either in their normal growth medium, or in Shepard’s storage broth described above, or in the media described by Lapage (this Series, Vol. 3A). Kelton (1964) lyophilized 48-72 h cultures in the medium described on p. 259, mixed with equal parts of sterile skim milk (10 g of powdered milk added to 100 ml of distilled water and autoclaved). Freezedried cultures were stored at - 65°C or - 26°C. All cultures remained viable for 3 or 4 years.

VII. TRANSPORTATION OF CULTURES Mycopfasma cultures may be transported in two main ways, either as freeze-dried cultures or as fresh cultures. Little need be said about the first method except that, as always, adequate precautions should be taken to avoid damage to the ampoule in order to prevent escape of the organisms with the possible production of a dangerous aerosol. It is important also to ascertain that there are no import regulations affecting the importation of cultures, as is the case in many countries with respect to veterinary mycoplasmas. When fresh cultures are to be transported they should be grown either in broth or on agar and then transferred to a screw-cap container of two-phase medium or semi-solid agar. This medium in turn may be incubated overnight and is then sent by air-mail (or express internal mail) to the recipient laboratory. In this latter way the authors have sent cultures of mycoplasmas to many parts of the world. T-strain mycoplasmas will withstand an overnight journey by post and this may well represent all the time that is needed by air-mail (or by special air freight) for distances of up to 6,000 miles. Care must be taken to ensure that the cultures to be transported are not allowed to grow to the point where the essential nutrients are exhausted or toxic substances (e.g., acid) are produced from the substrate in the medium

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since this would rapidly reduce the viability of the organisms. It is always wise to take a subculture from the culture just before dispatch so that should the culture fail to grow in the recipient laboratory it will be known whether or not the culture was viable at the time of dispatch. This information may save time-consuming checks on media formulation and control. Also it is helpful to send a small quantity of uninoculated growth medium with the culture so that an immediate subculture on to medium to which the organism is adapted may be made on arrival. After incubation, several aliquots of this subculture may be deep frozen to provide a basic working stock. REFERENCES Adler, H. E., andyamamoto, R. (1957). Am.J. wet. Res., 18,655-656. Afshar, A., Stuart, P., and Ruck:R. A. (1966). Vet. Rec., 78, 512-519. Anderson, D. R., and Barile, M. F. (1965). J. Bact., 90, 180-192. Andrewes, C. H., and Welch, F . V. (1946).J. Path. Bact., 58,578-580. Andrews, B. E., Davies, J., and Inglis, J. M. (1969). Znt. Symp. Mycoplasma diseases of Man. In press. Annotation (1968). Nature, Lond., 219, 438. Arisoy, F., Erdag, O., Cottew, G. S., and Watson, W. A. (1967). Turk vet. Hekim. Dern. Derg., 37, 11-17. Armstrong, D., Yu, B., Tully, J., Morton, V., and Friedman, M. (1968). Bact. Proc., M 81,79. Barile, M. F., Bodey, G. P., Snyder, J., Riggs, D. B., and Grabowski, M. W. (1966). J. nutn. Cancer Znst., 36, 155-159. Barile, M. F., and Schimke, R. T. (1963). Proc. SOC.exp. BioZ. Med., 114,676-679. Betts, A. O., Whittlestone, P., Beveridge, W. I. B., Taylor, J. H., and Campbell, R. C. (1955). Vet. Rec., 67, 661-665. Boidin, A. G., Cordy, D. R., and Adler, H. E. (1958). CorneZZ Vet., 48, 410-430. Borrel, Dujardin-Beaumetz, Jeantet, and Jouan (1 910). Annls Inst. Pasteur, Paris, 24,168-179. BridrC, J., and Donatien, A. (1923). C. r. hebd. Skanc. Acad. Sci., Paris, 177, 841843. Brighton, W. D., Windsor, G. D., Andrews, B. E., and Williams, R. E. 0. (1967). Mon. Bull. Minist. Hith, 26, 154-158. Brown, T. M., Swift, H. F., and Watson, R. F. (1940).J. Bact., 40,857-867. Carmichael, L. E., Guthrie, R. S., Fincher, M. G., Field, L. E., Johnson, S. D., and Linquist, W. E. (1963). Proc. U S . live Stk sanit. Ass., 67, 220-235. Chalquest, R. R. (1962). Avian Dis., 6, 36-43. Chalquest, R. R., and Fabricant, J. (1960). Avian Dis., 4, 515-539. Chanock, R. M., Fox, H. H., James, W. D., Gutekunst, R. R., White, R. J., and Senterfit, L. B. (1967). Ann. N . Y. Acad. Sci., 143, Art. 1,484-496. Chanock, R. M., Hayflick, L., and Barile, M. F. (1962). Proc. natn. Acad. Sci., U.S.A.,48,41-49. Chu, H. P. (1954). Quoted by Edward, D. G. ff. (1954).J.gen. Microbiol., 10,27-64. Chu, H. P., and Horne, R. W. (1967). Ann. N.Y. Acad. Sci., 143, Art. 1, 190-203. Clark, H. W. (1965). J. Bact., 90,1373-1386. Clark, H. W., Fowler, R. C., and Brown, T. McP. (1961).J. Bact., 81,500-502. Clyde, W. A. Jr. (1964). J. Immun., 92, 958-965.

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CHAPTER X J

Algae M. R. DROOP Scottish Marine Biological Association, Oban, Scotland I. Introduction . 11. Synopsis of Algal Nutrition 111. Isolation Methods . A. Enrichment techniques B. Manipulative techniques C. Antibiotic techniques IV. Maintenance . V. Culture Media VI. Some Special Methods . A. Continuous systems B. Synchronous cultures References .

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269 270 272 273 273 277 279 286 303 303 307 308

I. INTRODUCTION Algae are a morphologically and biochemically diverse group of plants, ranging from forms barely distinguishable from streptococcoid bacteria at the lower end of the scale to the giant kelps of the Californian coast at the other. Naturally it is only the smaller, mostly unicellular, members of the assemblage that can be thought of as micro-organisms and lend themselves to microbial methods of study. Yet even they differ so much in their way of life, their size range and the time scale of their activities from the majority of micro-organisms dealt with in this Scries that they warrant a chapter to themselves. The uses of algal cultures are many and varied : in phycology (morphology, cytology, taxonomy), in genetics, in physiology and biochemistry, and in ecology. Other applications include their use as sources of food for invertebrates in culture, of protein and other fine chemicals in industry, and more recently as components of self-supporting closed systems in space research. The treatment adopted here is elementary and based for the most part on the author’s personal experience. T h e temptation to detail the hundreds of, mostly trivial, culture medium formulations has been resisted ; they are available in the litcrature and many arc referred to.

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‘I’he readcr should consult the classic work of Fritsch (1935, 1945) for the most comprehmsive account of the algae, and Lewin (1962) for the most recent compendium of algal physiology and biochemistry. The latter also contains a taxonomic synopsis mentioning nearly all genera that have found their way into laboratories.

11. SYNOPSIS OF ALGAL N U T R I T I O N An outline of the cardinal fcatures of algal nutrition is included here as essential background to successful cultivation.

A. Carbon As a source of carbon most phototrophic species of algae can utilize carbon dioxide, which is taken up either as the free gas or as bicarbonate. In either case the availability of the source is to some extent dependent on the p H of the medium; moreover the very low carbon dioxide tension in a normal atmosphere and the limited carbonate alkalinity of culture media render it necessary to aerate the cultures or at least to plug the vessels with stoppers permeable to gas, otherwise growth may be severely limited. Cyclic, or bacterial, photosynthesis, in which an organic acid, usually acetic acid, or occasionally a sugar, takes the place of carbon dioxide, has been demonstrated in an increasing number of Chlorophyta and Eugleninae. Some of these algae, notably strains of Pyrobotrys and Chlamydomonas pulsatilla, are virtually unable to utilize carbon dioxide and have an obligate requirement in photosynthesis for acetic acid. Many, especially “laboratory”, algae can assimilate organic carbon oxidatively, in addition to being able to photosynthesize. Yet others, lacking chromatophores (e.g., Astasia, Polytoma, Prototheca) necessarily depend on oxidative assimilation. Virtually all species of chemotrophic algae will utilize acetic acid, while acetate is the only source of organic carbon available to many. On the other hand, hexose sugars often give better growth if they can be assimilated. Most examples of sugar-utilizing algae are found among Chlorococcales, Bacillariophyceae and Xanthophyceae, whereas acetate algae are typically Volvocales or Eugleninae. Although pH of the culture medium controls the passive entry of undissociated organic acids, acetate organisms are able to take up the anion. Uptake of the anion is accompanied by a rise in medium pN.

. Nitrogen Apart from certain nitrogen-fixing Cyanophyta, which can dispense with combined nitrogen, nitrogen is assimilated as nitrate or ammonium ions or occasionally in the form of simple amino compounds such as amino-

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acids or purines. Ammonium salts have the advantage over nitrates in not requiring reducing power for their assimilation. Indeed some algae are mable to reduce sources of oxidized nitrogen at all, whilc others are able to do so during phototrophic but not during chemotrophic growth. When both are present together ammonium ions arc assirnilatcci in preference to nitrates, although this depends to some degree on the medium pH. 'The general usefulness of ammonium salts is, however, limited to algae with a fair degree of acid tolerance. Not only are they extremely toxic even in mildly alkaline media, but assimilation of the cation quickly lowers medium pH beyond the tolerance limits of many algae. T h e ability of algae to utilize organic nitrogen depends on their being able either to take up the compound whole, when pH will have some influence, or to de-aminate it and assimilate the ammonium formed. There seems to be very little uniformity in the compounds utilized by algae, although urea, uric acid, guanine, arginine, tryptophan and lysine are favourites with Chlorophyta. For some marine algae unable to reduce nitrates and at the same time limited to alkaline media the only practical form of nitrogen is amino nitrogen. However, except in the richest environments, the normal diet of algae is nitrate nitrogen, since this form is not so available to fast-growing heterotrophic bacteria.

C. Phosphorus, sulphur, silicon and iron All algae utilize inorganic phosphate and sulphate. Sulphate poses no problem, but the insolubility of iron and calcium phosphates under alkaline conditions at best introduces an unknown element into artificial culture media and at worst can cause deficiencies. A possible answer is to employ weaker organic phosphoric acids such as glycerophosphoric because their calcium salts at least are more soluble than simple calcium phosphate. Diatoms, and presumably those Chrysophyta with silicious skeletons, need silicon. Silicic acid is insoluble and weakly ionized so that silicate ions are only found in the presence of alkali. T h e absolute requirement of diatoms is small, and it will be found that for many purposes with neutral or slightly alkaline media, sufficient dissotves from the walls of the glass vessels. Pringsheim recommended the use of soft glass for cultivating diatoms, but of course quantitative work on silicon nutrition should ideally be carried out in plastic vessels. Silicious precipitates, like iron and phosphate precipitates, adsorb trace nutrients and deficiencies can occur if they are removed by filtration. Iron presents special problems because of its ehtreme insolubility in aerobic, neutral or slightly alkaline conditions. At pFI 7, for instance, thc while rhe solubility equilibrium ratio of Fe3+to Fezf is of the ordcr of product of Fe3 + and OH- is of:the order of 10 3:). 'l'hc velocitics of the equilibrating reactions arc slow, so that thc rate of rephishtnent of fcrrous ions

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needed by the cells is seriously limited. In the presence of organic matter, however, complex formation occurs so that not only are the solubility relations improved but the equilibrating reactions are speeded up and replenishment of the iron species utilized can take place. Amino and aliphatic acids, peptides, humates, etc., are all more or less effective. Simple coniplexing compounds, such as ethylenediamine tetraacetic acid (EDTA) or citric acid are generally employed in defined media, whereas humates (as soil extract) are the most useful of the undefined organic supplements, as they contain little easily metabolizable organic carbon.

D. Trace metals, etc. It is beyond the scope of this article to discuss trace-metal requirements of algae, which ,cannot be demonstrated without recourse to special methods. However, the use of chelating agents for iron may lead to deficiencies in some of the heavy metals, and Mn, Cu, Co, Zn, Mo and Bo are usually included in chelated media. There is a fairly high requirement for Mo in nitrogen fixation. Ca, Mg, K and probably Na and V are required by algae.

E. Growth factors Vitamin requirements are common among unicellular algae, but the loss of synthesis appears to be generally limited to vitamin B12, thiamine and biotin, occurring in about 70,lO and 2% of known strains respectively.

111. ISOLATION METHODS For many purposes adequate control and reproducibility of cultures of autotrophic species can be achieved notwithstanding the continued presence of other classes of micro-organism. The so-called “agnotobiotic” cultures, though anathema to the purist, have much to recommend them, especially in taxonomic and invertebrate nutritional studies. In the first place, some species, mainly Chlorophyta and pelagic diatoms, tend to assume more naturalistic morphology when accompanied by other organisms. Secondly, less rigorous microbiological techniques are required for their maintenance. Thirdly, it has not been possible to cultivate a great many species in any other way. On the other hand, absolutely pure, or “axenic” cultures are obligatory for most physiological and biochemical work, while “monoxenic” cultures with one other organism, are used extensively for phagotrophs needing a living diet. Isolation methods for algae fall into three groups: (i) enrichment methods, in which nutritional or other conditions favour the wanted organism; (ii) manipulative methods, by which the wanted organism is separated mechanically from the others; and (iii) antibiotic methods, in which poisons

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are used to kill off the unwanted flora. Very often a combination of several methods can be used to advantage. Whatever the method used the starting material should be in a state of active growth, when it is more likely to be healthy and vigorous with plenty of young cells free of attached microorganisms.

A. Enrichment techniques Populations of algae may be obtained simply by bringing in pond water, seawater, etc., into the laboratory, adding a little soil extract or nitrate and leaving to stand by a window away from the sun. This is an enrichment technique and is often very effective, especially for getting vigorous material for further isolation. More sophisticated enrichment techniques are designed to establish excIusive conditions and are mainly used to screen for organisms with particular nutritional abilities or physical tolerances, e.g., acetate or glucose utilizing algae (Pringsheim, 1936, 1946, 1951a; Lewin and Lewin, 1967), thermophils (Sorokin and Myers, 1953; Allen, 19-59), acidophils (Pringsheim, 1946; Allen, 1959; Fott and McCarthy, 1964), species able to withstand desiccation (Droop, 1953), and sexual algae (Lewin, 1951). Although it is sometimes possible to obtain agnotobiotic but unialgal cultures by enrichment alone the technique is usually followed by others. The principle underlying enrichment is ecological; so also is the main objection to it when it is used as an instrument of primary selection, for there is no guarantee that the resulting populations are ecologically representative, adapted to the desired conditions though they may be. Indeed, the opposite is often the case and one tends to end up with “laboratory weeds” having selected, apart from anything else, for tolerance of laboratory conditions. B. Manipulative techniques Mechanical means have been used extensively to separate algal cells from other organisms. Their success depends on the fact that young vigorously grqwing cells are normally free from attached micro-organisms. They are less open, than are antibiotic methods, to the objection that the cells may have been altered by the procedure without the operator knowing. The so-called washing technique, in which single cells are taken through several baths of sterile medium and used to start clonal cultures, has advantages over all other methods, namely that the actual cell to be isolated can be chosen, and it is possible to make an isolation even when very few cells of the wanted species are represented in the sample. However, single cells of many delicate species, especially pelagic forms, do not survive in bacteriafree culture media, although very often they will withstand isolation if bacteria are allowed to remain. The usual strategy for such forms is to

274

M. R. DROOP

establish clonal bacterized cultures and to purify these with antibiotics. The other widely used manipulative method is the conventional plating technique, which however is useful only with the limited range of algae able to grow on solid media (some Chlorophyta, Cyanophyta, Eugleninae, pennate diatoms), but has the great advantage of simplicity and ease.

1. Centrifugation As a preliminary to plating or washing, algal cells may be concentrated against bacteria by repeated differential centrifugation, a few minutes at 300 g being suitable. Gentle centrifuging is also useful for concentrating sparse plankton samples prior to isolation, although many of the larger dinoflagellates will not tolerate it. T o overcome this difficulty an ingenious and simple method of slow filtration has been devised by Dodson and Thomas (1964). 2. P+etting or washing Single cells are transferred under a dissecting microscope through successive baths of sterile medium with a micro-pipette. The method is very effective since a very small amount of liquid may be transferred with the cells and an effective dilution factor of over a million can be achieved at each stage. Cells as small as 30 pm in diameter are easily handled, but smaller cells require patience. The absolute limit of the method (Droop, 1954) is about 5pm; smaller cells would require a micromanipulator. For the baths, Lwoff (1929) employed single drops in cavity slides, while Pringsheim (1946) preferred three or four drops in watch glasses with correspondingly fewer stages because of the greater dilution factor. The advantage of single drops over larger volumes is the ease in locating the cells, which however is offset by the greater interference from surface effects when the pipette is introduced. A large surface is also deleterious to delicate cells sensitive to physical changes in their environment; indeed, there is much to be said for increasing the volume of the bath to 3 or 4 ml (Droop, 1954), when five or six baths are more than sufficient. Pringsheim described his technique very fully in his book (Pringsheim, 1946), but the following outline is based on the writer’s practice. The dissecting microscope should be an old-fashioned one without built-in illumination or heavy multiple nosepiece, as it is essential to be able both to vary the angle of the transmitted illumination and to have the focussing finger-light. The lower surface of the glass stage should be finely ground-a ground-glass filter disc cemented to the under side of the stage serves very well. A cardboard shield should be fitted to the microscope to protect the open Petri dishes from the operator’s breath. Elaborate aseptic

XI. ALGAE

275

chambers and inoculating cupboards are usually both unnecessary and undesirable. The Petri dish/watch glass combinations need not be absolutely sterile, but should have been oven sterilized and stored dry away from dust. The watch glasses are of the normal biological type with a flat central area to prevent rocking. The micro-pipettes are made from ordinary Pasteur pipettes plugged at the top end with cotton-wool, sealed at the point and stored sterile. The thin part of the pipette should be 8-10 cm in length and 1 mm in diameter. Just before use this is drawn out further to a very fine diameter in a small flame. The actual diameter required depends on the size of the organism being isolated, but in any case should not be less than 30 ,urn because of the strength of the capillary forces. The liquid is drawn into the pipette by capillarity-no suction is necessary-but the same force obliges one to use a 5 cm length of fairly stout rubber tubing stopped by a glass bead, in place of the usual eye-dropper cap, to eject the liquid. The fine part of the pipette should be terminated by a clean break to a length of 4 cm; a sharp longitudinal pull with flamed forceps effects this. Sterility of the micro-pipettes is assured if they are fashioned afresh between each step in the manipulation. The isolating medium should not differ too sharply in its gross physical characteristics from the medium the organism has been used to, but it should not have too low a surface tension, for persistent bubbles can be a nuisance, Usually it will be a mineral medium with the addition of a little soil extract. With brackish and marine organisms special attention has to be given to salinity. This, however, can be estimated to sufficient accuracy by preparing a series of seawater dilutions with distilled water and introducing carefully into the centre of each a small volume of the sample and observing whether it rises or sinks. It is an advantage to allow delicate cells to acclimatize for an hour in a variety of media before starting the manipulation and to carry on with whichever seems best. The actual manipulation requires patience but, like riding a bicycle, is not difficult once the skill is acquired. Once the cell is located it is picked up by momentarily lowering the point of the pipette to it. Several cells may be transferred together, except at the final stage since the aim is clonal cultures. The point of the pipette should not be allowed to remain submersed for longer than is absolutely necessary, and certainly never while searching, since it draws in water all the time it is beneath the surface. Larger cells can simply be ejected randomly into the fresh bath, as described by Pringsheim, but smaller cells are best watched under the microscope while being ejected. This is not easy but is well worth while in terms of time saved in searching. The trick is to rest the pipette for one second below the surface before picking up the cell in order to form a buffer of water between it and

276

M. R. DROOP

the air in the pipette. This allows time after ejecting the cells to withdraw the point from the vicinity before the inevitable train of bubbles carries them away. Motile flagellates should be placed on the bottom of the dish as they will swim up, but non-motile cells should be allowed to fall through the medium. I n general, the manipulations are carried out under the lowest practical power of the microscope. Once an organism has been identified under higher power it can usually be distinguished with much less magnification. Flagellates are particularly easy because their movement is often distinctive. T h e fact that motile cells tend to migrate to the meniscus renders searching less arduous than it might otherwise be. Moreover, some flagellates show marked, usually negative, phototaxis in isolation media, which can be made use of. In favourable instances when the material is sufficiently abundant the whole procedure, except the very last step of placing single individuals in the culture tubes, can be carried out without the aid of a microscope. Phototaxis permits organisms as small as 5 /cm to be manipulated with great ease (Droop, 1954).

3 . Agar methods Plating techniques were first applied to algae by Beijerinck (1890); they only differ from conventional bacteriological methods in small details. T h e sample for isolation will usually require a considerable degree of dilution before plating. A series of ten-fold dilutions is convenient from which the plates can be prepared by any of the conventional methods for streaking or pouring. Selective, purely mineral, agars are often used, but selectivity may have more drawbacks than advantages. A little soil extract greatly increases the chances of algal growth, although of course it also increases the variety and to a lesser extent the quantity of bacterial growth. Organic enrichments, which favour the growth of heterotrophs, are not suitable. Media for algae are usually prepared with 0.9-1.0% agar instead of the usual bacteriological 1.5%. The actual concentration of agar used varies with the grade. I n general the lowest usable concentration should be employed. This necessitates greater care in streaking the plates so as not to break the surface of the agar, and it is essential to incubate the cultures agar surface downwards and illuminate them from above to avoid drops of condensation forming on the agar. Excess evaporation from the agar is preferably prevented by part filling the Petri dish lid with a layer of water than by sealing lid and dish with tape. Algal plates will usually reqtlire two to three weeks’ incubation before isolations can be made from them. Plates may be either streaked or poured according to whether it is desired to isolate from surface or deep-growing colonics. Many algae, e.g., some

XI. ALGAE

277

Cyanophyta, fail to grow on the surface but will grow in the body of the agar (Allen, 1952), while on the other hand the temperature at which plates are poured is not tolerated by delicate species even for the short time of the operation. Colonies are more difficult to pick when they are below the surface but they are less prone to contamination from fast-growing motile bacteria. The agar surface may be washed and quickly sterilized with dilute Lugol’s iodine solution before beginning the isolations. Isolations are made from the plates as soon as recognizable algal colonies appear. It is not difficult to distinguish clean from contaminated colonies under the dissecting microscope, but clonal colonies should always be selected. Most bacteriologists recommend a wire loop or needle for the transfers to culture tubes. However, anyone who has once used Pringsheim’s micro-pipettes for this purpose is unlikely to return to the wire needle. One can isolate directly from colonies beneath the surface with a pipette without having first to cut them out and expose them with a sterile scalpel. Transfers from the isolation plates are invariably made to liquid or semisolid media. Most algae grow poorly on agar and it is best to let them become established in liquid culture before adapting them to the more rigorous conditions of an agar slant.

C. Antibiotic techniques Many algae, other than Cyanophyta, are more resistant than bacteria to the common antibiotics and can readily be purified by this means. The simplicity, reliability and wide application of antibiotic techniques have given them pride of place among algal methods. However, in common with other repressive techniques, one is never quite sure that their use has not damaged or otherwise altered the algal cells. For this reason one should in preference resort to antibiotics only when other methods have failed or obviously hold no promise. Since antibiotics mainly repress only the bacteria the starting material should be clonal. Antibiotics are most useful for gelatinous species, for minute species that fail on agar and especially for the many fastidious species, including the large class of pelagic marine and freshwater organisms, that fail to multiply from single cells in the absence of bacteria. The most widely used antibiotics are penicillin, streptomycin, chloramphenicol and to a lesser extent neomycin and the chlorotetracyclins. The wide range of bacteria usually found in field collections or enrichment cultures of algae renders single antibiotics less efficacious than mixtures. Too frequently the use of a single antibiotic merely reduces the species count, although complete success has on occasion been recorded with penicillin or streptomycin. Penicillin and streptomycin are to some extent complementary in that Gram-positive organisms tend to be sensitive to the

278

M. R. DROOP

former and Gram-negative to the latter, while chloramphenicol has a wide spectrum. Probably the most popular technique is to carry the algae through several transfers at one to three day intervals in a culture medium containing penicillin or streptomycin or a mixture of the two (Lewin, 1959). Some algae will continue to multiply in as high a concentration as 500 mg/litre penicillin. However, the method described below has been used routinely for more than ten years in the author's laboratory on a variety of organisms including Lamellibranchs, copepods, rotifera, protozoa, algae and bryophytes (Droop, 1967). It depends for its effect on a shorter exposure to a much higher concentration of a mixture. TABLE I Antibiotic mixtures (amountsin mg/ml, as prepared before making serial dilutions)

V Diatoms Benzyl penicillin sulphate Streptomycin sulphate Chloramphenicol Neomycin

8000 1600 200

..

VI

VII

Rotifers, Noctiluca, Tigriopus, Micromonas, etc. diatoms 8000 2000 ip 8

..

8000 1600 80 40

Algae vary in their tolerances to the various antibiotics (Provasoli, Pintner, and Packer, 1951); even closely related species can differ widely in their tolerance. The mixtures detailed in Table I were originally based on the tolerances of the organisms indicated to a 24 h exposure to the individual components, but they have found much wider application. Number VI has been especially useful. Since one usually requires quite small amounts on any one occasion it is convenient to prepare and store the mixtures in a dry state. The schedule requires six parallel test tube cultures, which should be as dense as possible but still multiplying. The required quantity of antibiotic mix is dissolved in a medium of similar composition to the cultures. Only one tube is required for the schedule, but it is advisable to prepare sufficient for two. The solution should be prepared on the day it is to be used. It is filter-sterilized, conveniently with an asbestos or membrane syringe filter, into a sterile tube to roughly the same volume as the cultures. Then a series of two-fold dilutions of this antibiotic solution is made with the algal cultures by mixing the tubes successively over a flame. One drop, no more, of an organic medium is now added aseptically to each tube to enhance the action of the penicillin

' XI. ALGAE

279

in the mixture. The tubes are numbered in order of decreasing concentration and incubated in the light. After 24 h transfers are made to antibiotic-free media. The fact that a medium has been suitable for bacterized cultures is no guarantee that it will support axenic growth. It is therefore advisable to prepare a variety of solutions for the subcultures. Six tubes of each are required, and if one works from Nos 1-6, in that order, a single Pasteur pipette can be used for the whole operation. It is of course essential to indicate on the subcultures both the medium and the origin of the transplant. All these cultures should be tested for sterility after a fortnight to a month. Typically, viable algae will be found in the subcultures from antibiotic tubes 3-6 and viable bacteriq in those from tubes 5 and 6. It is always advisable to keep all the viable dgal lines for several transplants, for bacteria may take several months to become re-established after antibiotic treatment. Moreover, one cannot be sure (especially if the antibiotic mixture contained chloramphenicol) that algal growth in the first subculture will be continued in the next. The schedule is of course not invariably successful. In the event of failure the best course is merely to repeat it using material hitherto unexposed to the treatments. If that fails it is worth trying the viable culture having received the highest antibiotic dose. With persistent failure the simplest course is to re-isolate from nature. Only in the last resort should it be necessary to spend time in tailoring antibiotics to the bacteria and algae. Moulds do not give much trouble as a rule, especially in, marine media. They are best eliminated by the washing method beforehand.

IV. MAINTENANCE The minimal apparatus requirements for maintaining a small collection of algae is a convenient window not receiving direct sunlight, a few gross of test tubes, a pressure cooker, cotton-wool, Pasteur pipettes and a dispensing balance, but some form of artificial light source and temperature control is needed for experimental work. Obviously, the smaller the volume of individual cultures the less space will a culture collection take up. The most convenient vessel will probably be found to be 16 x 160 mm Pyrex tubes without rims. The larger pelagic diatoms and dinoflagellates, and of course, multicellular algae, are more conveniently carried in Erlenmeyer flasks than test tubes; and should larger volumes be required penicillin pans, Roux flasks and the like can be employed. In general, however, the inconvenience of large vessels is not conducive to the maintenance of good order in a stock collection. The carbon dioxide requirement of algae precludes the use of tight-fitting screw-capped containers, while loose caps, which are very useful on occasion,

280

M. R. DROOP

are not absolutely safe over a long period and should not be used for stock cultures. Failing the marketing in this country of tight-fitting polypropylene caps, permeable to carbon dioxide and oxygen but not to water vapour, there remains the old-fashioned non-absorbent cotton wool as the most satisfactory stopper for algal culture vessels.

A. Artificial illumination Unless other conditions are optimal it will be found that a rather low level (1000 lux) of artificial illumination gives the best results. One thousand lux is far below the levels that most physiologists would employ, for strains of Chlorella or Euglena gracilis for instance, but is high enough for maintenance and is tolerated by most algae. No artificial illuminant is really satisfactory and species will be encountered that appear to require natural daylight. Tungsten lamps give a continuous spectrum but one comparatively richer in longer wavelengths than sunlight, and without ultraviolet emission, which is one of its advantages. More or less elaborate steps have to be taken to dissipate the heat generated by tungsten lamps (Pringsheim, 1946; Jitts et al., 1964). Fluorescent light is generally most satisfactory because it is cool, but the spectrum consists of a number of discrete emission bands, which in general do not coincide with the regions of maximum absorption by chlorophyll. “Warm White” is probably better than “Daylight” or “Cool White” since it passes less ultraviolet.

B. Temperature control In temperate climates bacteria-free cultures can be maintained at “room temperature”, but cultures containing a mixed microbial flora are best kept at 12”-15”C. Most algae grow very poorly in the laboratory below 10°C irrespective of their natural habit. In warmer climates some form of cold room or cabinet is needed. A very convenient cabinet can be constructed in Handy Angle or Dexion faced with 1& in. expanded polystyrene boards, illuminated from below through a double-glazed base. A domestic refrigerator replacement unit (4 ft3 capacity) will maintain 9 ft3 at any temperature between 5” and 27°C with 160 W fluorescent lighting 12in. below the base. Illumination from the side, as found in some commercial continental and American cabinets, cannot possibly give equal illumination to all the cultures. Control is effected by a mercury-toluene thermometer, which, for really trouble-free operation over long periods, should be made to control the relay by interrupting a light beam rather than an electrical circuit directly, although satisfactory results are obtained if the interrupted current is limited to a few hundred micro-amperes. In either case the mercury should be covered by medicinal

281

XI. ALGAE

paraffin to protect the meniscus. The design of the trigger circuits depends of course on the characteristics of the relay; the circuit illustrated in Fig. 1B is suitable for a 1.5 kQ, 22 mA hot wire vacuum relay, Type F202/10, acting upon the heater circuit directly. With this polarity the photo device is more or less fail-safe, but should it be necessary to control the cooler circuit directly the polarity of the trigger can be reversed by connecting up the OCP 71 as in Fig. 1A. The extremelylow temperature differential characteristic of the mercury-toleune thermometer will not of course be realized in -6V

P

A

-33

v

0

B

FIG.1. Phototransister-operated Schmidt trigger with 1.5 kaZ hot wire vacuum relay switch as load. A and B: alternativewirings for alternativepolarity of operation.

air, but if there is good circulation in the cabinet the differential can be reduced to less than a degree; the fan should be able to move 25-50 ft3lmin. Sealed contact thermometers are easier to operate than the mercury-toluene ones but are not always reliable. Cartridge thermostats are robust and a good deal 4heaper and are capable of differentials of less than a degree. Both devices can with advantage be used with the trigger circuit shown by omitting the OCP 71 and connecting the thermostat between earth and either the base or collector of the OC 75 according to the polarity required. For physiological work requiring very small temperature differentials there is no substitute for an illuminated, temperature-controlled water bath. A mercury-toluene thermometer controlling a cooling coil and a 500 W heater in conjunction with a small bubble gun (200mllmin) for mixing

282

M. R. DROOP

will give a differential of less than a quarter of a degree in a 6ft3 bath. Much smaller differentials are possible with more vigorous mixing. The base may be a quarter-inch plate glass seated on a non-hardening putty, with illumination from below, but for higher light intensities the fluorescent lamps should be mounted in tubes through the body of the bath. Some form of scaffolding has to be provided to hold the culture flasks or tubes. For some applications, e.g., continuous systems and mass cultures, it is more convenient to pump temperature-controlled water through a jacket surrounding the culture vessel or through a heat-exchange coil within the culture itself. I n experimental work having temperature as one of the parameters one needs a number of temperature settings simultaneously. The problem is most elegantly solved by use of a temperature gradient. The apparatus utilizes the heat flow through an aluminium block which is warmed at one end and cooled at the other. The culture tubes may be placed in holes drilled in the block (Thomas et al., 1963; Jitts et al., 1964) or the block may form the base of a large agar “plate” (Halldal and French, 1958). In either case the layout is two-dimensional, so that a second parameter, usually light intensity, can be included in the experimental design.

C. Aeration and shaking No form of aeration or shaking is recommended for the mere maintenance of cultures where the objective is to preserve strains safely and with the minimum of labour. Nevertheless a still culture with the cells, more probably than not, settled in a mass at the bottom, creates the least favourable conditions for growth, with mutual screening and local nutrient depletion. Even at the best of times the rate of growth of heavy algal cultures is likely to be limited by the rate of diffusion of carbon dioxide into the medium. Shaking or rolling increases the solution rate and, by keeping the cells in even suspension, ensures that all are treated alike. Algae, however, can be sensitive to the frequency of shaking and growth may even be depressed by this treatment. The same may be said of aeration, particularly in the early stages of growth. The reasons for this are not altogether clear but are very likely connected with the pbysicochemical state of the iron supply (Droop, 1961a, 1962). Besides keeping the cells in suspension aeration effectively inqeases the surface area of the culture medium. Definite relations exist between the partial pressure of carbon dioxide in the air stream, the bicarbonate or carbonate concentration and the pH of the medium at equilibrium. They are approximatelylog [HCO3-] - log Pcoz = pH - 7.6 log [C03’-]

- log Pcoz = 2pH - 16.6

XI. ALGAE

283

The log PcoZ of normal air, containing approximately 0.03% carbon dioxide, is - 3.5. This formula is useful for setting up carbon dioxide generators (in which, for instance, air is bubbled through a solution of sodium carbonate or bicarbonate adjusted to the required pH with hydrochloric acid) and for stabilizing the pH of media bubbled with a mixture of air and carbon dioxide. Heavy algal growths prevent equilibrium being reached and it is then possible to give a much greater carbon dioxide enrichment without lowering the pH and to increase the yield thereby. Enrichments of over 0.5% carbon dioxide are impractical with chemical generators and in any case cylinder gas is often more convenient. If the mixtures are not purchased ready made, flow meters and constant pressure devices for the carbon dioxide and air streams will be needed to ensure adequate control of the mixture’s composition (see Platon, this Series, Vol. 2). Five per cent carbon dioxide is commonly employed by physiologists, but with marine media so high an enrichment entails acceptance of working pH values below 7.1 so as to avoid calcium carbonate precipitation. Spencer (1966) discusses the whole question of pH and COz control in marine systems. The aerating gas has of course to be saturated with water vapour and to be sterile. A 1 x 12 in. length of glass tubing tightly packed with cotton-wool forms a cheap and reliable bacterial filter, but there are membrane filters especially marketed for this purpose. A cotton-wool filter should be kept dry with 6-10 W at 6 V, dissipated in a coil of resistance wire wound round the filter. One such filter can lead to a manifold to serve many cultures. Silicone rubber will be found to be the most suitable material for tubing and bungs, being non-toxic and heat sterilizable. All apparatus including, and down stream of, the filter must of course be sterilized, and the clipped ends of any idle lines should be kept sterile by immersion in 70% alcohol,

D. Sterilization Empty glassware can be steam-sterilized at 25 lb for an hour, but culture media should be subjected to as gentle a treatment as is compatible with a reasonable chance of absolute sterility. Two minutes at 15 lb is sufficient for test tube media, and 10 min for flasks containing 500 ml. A domestic pressure cooker is suitable for small quantities. Certain media components react when autoclaved together (see Bridmn and Brecker, this Series, Vol. 3A). Glucose and other reducing sugars are hydrolysed in acid or alkaline solutions and it is sometimes necessary to autoclave them apart from the rest of the medium. Agar is also denatured when autoclaved in acid solution. Natural extracts, such as liver, yeast, beef and soil extracts throw precipitates when autoclaved with natural seawater, which should therefore be autoclaved apart from them. Ferrous salts or

284

M. R. DROOP

complexes, e.g., ferrous ammonium citrate, may profitably be filtersterilized and added to the medium aseptically. Filter-sterilization, either through asbestos or membrane filters, enables heat-labile components of the medium to be preserved, but generally filtration yields a poorer medium than the same formulation autoclaved. There are two possible reasons for this, both of which concern the supply of iron especially in defined media. First, the reducing conditions in the autoclave ensure that at least some of the iron is returned to the ferrous state; and second, most of the iron in these mildly alkaline defined media is colloidal, even when chelated. Such iron is liable to be removed by filtration (Droop, 1961a, 1962). Phosphorus and humic colloids can also be removed from media in this way. Thorough sterilization of culture media is not always necessary or even desirable for bacterized cultures. Fastidious species often do better in pasteurized media. This applies particularly to Pringsheim’s biphasic soil-water media whose physical properties are destroyed by autoclaving (Pringsheim, 1946), and to media containing full-strength natural seawater, which may precipitate badly.

E. Maintenance and sterility testing It requires a methodical routine to maintain even a small culture collection reliably, and time spent on labelling, sterility testing, etc., will not be wasted,

It is convenient to number strains consecutively regardless of taxonomy and to keep a catalogue €or all information relevant to each. A culture sheet is kept for each strain in the collection; this enables subculturing and sterility testing to be carried out in a systematic manner (Fig. 2) (see also Lapage et uZ., this Series, Vol. 3A). The routine is simple enough, and will be clear from the Figure. It ensures that subcultures are only made from otherwise unused sterility-tested cultures. A golden rule is that borrowings of stocks are never made from the collection for any purpose. When cultures are needed for experiments or examination extra subcultures are taken, preferably from used stocks. Maintenance should always have priority and no risks should be taken with stocks. The most sensitive sterility-test media are those that show up the largest variety of bacteria, Medium “E 6” (Provasoli et al., 1957), a mixture of the two media of Table XI mixed with an equal volume of seawater, can be used routinely for convenience, but it could with advantage be diluted tenfold with dilute seawater or saline. In general, a very dilute liquid medium containing glucose, peptone, soil and liver extracts, incubated at 22°C for a month is more sensitive than a stronger medium incubated for a shorter period at a higher temperature. Agars are less suitable €or sterility testing.

285

XI. ALGAE CULTURE SHEET

1964

Organism Prymnesium parvum

Strain No. 65

Medium S 50

Frequency monthly

Subcultures

I

Sterility Test

Source

Date

1

Result

2213

311 311

2414

I

d

2414

1212

2915 2016

Date

1

1317

2915

X

2213

2016

d

2213

1317 ( 1 )

d

-

I

Notes

cQntamn.discard.

2 cultures set up

2915 -

,

F. Chemical cleanliness Standards of cleanliness in both chemicals and glassware will vary with the uses to which the cultures are to be put. Reasonable standards should always be aimed at, but there is no point in being very much cleaner than necessary. For instance, it would be pointless cleaning glassware to a standard suitable for trace-metal studies if it is to be used for undefined media, or even defintd media prepared with unpuritied AnalaR standard chemicals. One is never happy about the use of detergents for washing glassware. Soap followed by dichromate-sulphuric acid, then a thorough rinsing in tap, then distilled, water is a preferred routine. If the culture media contain neither natural extracts nor chelating agents the glassware should be steeped in versine (Naz EDTA) before final rinsing, to remove any traces of chromium. The dichromate-sulphuric schedule is suitable for most purposes, although not for trace-metal studies. Inorganic AnalaR chemicals are sufficiently free of organic impurities to need no further purification for most applications, including vitamin studies, but inorganic impurities are not negligible. For this reason trace-metal

286

M. R. DROOP

requirements are more easily ascertained for fresh water than for marine species. For special methods for trace metals, see for instance Stout and Arnon (1939), Hewitt (1952), and Hewitt et al. (1965). Rinsing of pipettes should be done by hand rather than by automatic pipette washer. The microbial ecology of pipette washers is quite a study. Water for defined media is best double distilled with a crystal of potassium permanganate in an all-glass still. De-ionized water is also suitable provided the apparatus can be kept free of micro-organisms in its dead spaces. Tap water can be used for natural media if the local authority is not in the habit of chlorinating the supply.

V. CULTURE MEDIA An annotated selection of algal media is listed in Tables 11-XIII. Details of many of the older media are to be found in Kuffareth (1929), Pringsheim (1946), Bold (1942), Neish (1951), Allen (1952), Pringsheim (1951b), Provasoli et al. (1957) and Links et al. (1961), while formulations of some forty simply prepared media in use in culture collections throughout the world are given by the Committee on Cultures, Society of Protozoologists (1958). It is not possible to review the hundreds of formulations adequately in a reasonable space, and the media discussed below should be regarded as exemplary rather than representative. The practice of the leading exponents in the field should be consulted, bearing in mind always the particular aims and scope of their work.

TABLE I1 Undefined freshwater media (mineral)f. Reference

Designation

Notes ~

Chu, 1942 Pringsheim, 1946 Provasoliand Pintner, 1953

1-17

~

~

Defined media prepared with lake water Various, including soil/water or soil extract media Synura.Contains peat extract

t I n Tables 11-IX the designation “mineral” is given to media containing little easily metabolized organic material ;such media are suitable for bacterized cultures. Media labelled “organic” are only suitable for bacteria-free cultures.

287

XI. ALGAE

TABLE I11 Undefined freshwater media (organic) Reference

Designation

Pringsheim, 1946 Reynolds, 1950 Lewin, 1952 Pringsheim, 1952 Storm and Hutner, 1953 Provasoli and Pintner, 1953 Lynch and Calvin, 1953 Baker et al., 1955 Norris et al., 1955 Damforth and Wilson, 1957 Proctor, 1957 Ford, 1958 Moewus and Moewus, 1959 Bendix and Allen, 1962 Pringsheim, 1964 Fott and McCarthy, 1964 Pringsheim, 1966 Lewin et al., 1966

A, B

E

Notes Various Stigioclonium Chlamydomonas moewusii Ochromonas malhamensis Peranema Cyanophora, Peridinium ,Ochromoms Euglenagracilis, COz fixation Euglena gracilis Survey of photosynthesis Euglena gracilis Haematococcus phvialis Ochromonas Polytoma Chlorella BeggMtoa Acidophilic Volvocales Lampropedia Navicula pelliculosa

TABLE IV Defined freshwater media (mineral) Reference

-~

ltkistol Roach, 1926 Craig and Trelease, 1937 Allison etal., 1937 Chu, 1942 Fogg, 1942 Emerson and Lewis, 1942 Pratt, 1943 Algeus, 1946 Rodhe, 1948 Fogg, 1949 Osterlind, 1949 Hutner and Provasoli, 1951 Gerloffetal., 1950a,b Hutner et al., 1950

Designation

1-1 7

I-VIII A, B, C

1-1I I

Notes Chlorococcales Chlorella Cyanophyta, Iron sole trace metal Planktonic algae Anabaenrr. Unchelated Chroococcus Chlorella Chlorococcales Planktonic freshwater algae Ambaenrr Scenedesmw Phytoflagellates. Chelated trace metals and vitamins Cyanophyta Trace metal studies

288

M. R. DROOP

TABLE IV-continued Reference

Designation

Notes ~

Pringsheim, 1951b Lewin, 1951 Neish, 1951 Allen, 1952

Neeb, 1952 Sager and Granick, 1953 Provasoli and Pintner, 1953 Syrett, 1953 Kessler, 1953 Tamiya et al., 1953 Lewin, J . C., 1954 Norris et al., 1955 Allen and Arnon, 1955 Arnon et al., 1955 Kratz and Myers, 1955 Proctor, 1957 Miller and Fogg, 1957 Kessler et al., 1957 Hughes et al., 1958 Zehender and Hughes, 1958

3,4

V, PG

A, F, J, K, 10

Sorokin and Krauss, 1958 Allen, 1959 Chorney et al., 1959 Wise, 1959 N 2, N 2b Soeder, 1960 Zehender and Gorham, 1960 Griffiths et al., 1960 Meffert, 1960 Kiyohara et al.,1960 Reisner et al., 1960 McLachlan and Gorham, 1961 Schmidt, 1961 Lazaroff and Vishniac, 1961 Miychi and Tamiya, 1961 Soeder et al., 1962 Scotten and Stokes, 1962

ASM

N5

~-

Micrasterias. Also Molisch, Beijerinck, Knop, Detmer,Czurda,Chu Chlamydomonas chlamydogumo, C. dysosmos Chlorella. Also Emerson, Warburg, Kok, Molisch, Beijerinck Cyanophyta. Also Richter, Molisch, Warder, Pringsheim, Beijerinck, Drews, Allison, Emerson/Lewis, Chu Hydrodiction Chlamydomonasrheinhardii Cyanophora, Petidinium, Synura Chlorella. Citrate and Arnon’s A 4 Chlorella Chlorella Navicula pelliculosa Photosynthesis Anabaena cylindrica. N fixation Scenedesmus.Trace metals Cyanophyta Heamatococcuspluvialis Monodus and other Xanthophyta Chlorella spp. Microcystis Cyanophyta ,Xanthop hyta, Chlorophyta Chlorellaspp. Cyanidium (thermophil and acidophil) Chlorella in D2O Polytomella base Chlorella Microcystis Chlorella. Craig and Tralease, Fe EDTA Mo. Scenedesmus Tolypothrix Chlorella vulgaris base Microcystis. Provasoli’s ASP modified. Chlorella, synchronization Nostoc Chlwella, synchronization Chlorella Beggiatoa

+

+

289

XI. ALGAE

TABLE IV-continued Reference

Designation

Notes

Astasia base Chlamydomonaspallens Haematococcuspluvialis Chlamydomonasmundana base Lichen symbionts Chlorellabase Aphanocapsa Anacystis. Kratz and Myers modified Chlorella spp., photometabolism Pediastrum. Based on Chu 11 and Rodhe IV Chara Chlamydomonasmundana base Chara Scenedesmus

Buetow and Padilla, 1963 Pringsheim, 1963 Stross, 1963 Eppley et al., 1963 Watanabe and Kiyohara, 1963 Galloway and Krauss, 1963 Moyse and Guyon, 1963 Kumar, 1963 Syrett et al., 1963 Davis, 1963 Fosberg, 1965 MaciasR, 1965 Imahori and Iwasa, 1965 Marsh et al., 1965

TABLE V Defined freshwater media (organic) Reference

Designation

Hopkins and Wann, 1926 Hutchins, 1941 Anderson, 1945 Provasoli et al., 1948 Hutner et al., 1949 Hutner et al., 1950 Little et al., 1951 Allen, 1952 Cramer and Myers, 1952 Lewin, 1953 Barber et al., 1953 Hutneretal., 1953 Hutner etal., 1956 Hutner et al., 1957 Hall, 1957 Allen, 1959 Greenblatt and Schiff, 1959 Aaronson and Baker, 1959 Aaronson and Scher, 1960

1-111

Notes

Chlorella. Contains glucose and citrate Chilomonas paramoecium Prototheca Apochlorotic Euglena Euglena gracilis. Vitamin BIZassay Trace metal studies Polytomella Cyanophyta Euglena gracilis Chlamydomonas moewusii Ochromonas. Vitamin Biz assay Ochromms Euglena gracilis. Vitamin BIZassay Ochromms. High temperature medium Chilomonas Cyanidium, (thermophil and acidophil) Euglena gracilis Ochromonas Ochromonas spp, Euglena gracilis

290

M. R. DROOP

TABLE V-continued qlr

Reference

Designation

Notes Astasia. Synchronous culture Synura, Vobox spp, Phacus, Woloszynskia Pyrobotrys Brachiomonas and Sphaerellaceae Polytomella. Also Lwoff, Pringsheirn, Links Ochromonas danica. Vitamin BIZ assay Poteriochromonas Euglenagracilis Haematococcuspluvialis Chlamydomonas mundana Poteriochromanas Euglena gracilis. Hutner et al., 1956, modified Astasia Euglena gracilis, bleached Euglena gracilis. Auto- and heterotrophic studies Chlamydomonas mundana Prototheca. No trace metals Euglenagracilis assay media. Review

Padilla and James, 1960 Provasoli and Pinmer, 1960 Pringsheim and Wiessner, 1961 Droop, 1961b Linksetal., 1961 Baker et al., 1962 Isenberg et al., 1962 Hurlbert and Rittenberg, 1962 McLachlan, 1962 Eppley and Macias, R., 1962 Isenberg, 1962 Price and Vallee, 1962 Barry and Barry, 1962 Moriber etal., 1963 Klein et al., 1963 Macias R. and Eppley, 1963 Callely and Lloyd, 1964 Hutner et al., 1966

TABLE VI Undefined marine media (mineral) Reference

Designation

Notes

~

Miquel, 1890 AllenandNelson, 1910 Schreiber, 1927 Fcayn, 1934 Barker, 1935 Ketchurnand Redfield, 1938 Matudaira, 1942 Sweeney, 1951 Goldberg et al., 1951 Spencer, 1954 Droop, 1954 Sweeney, 1954 Lewin, R. A., 1954 Haxo and Sweeney, 1955 Norris et al., 1955

“Miquel”

Diatoms Based on Miquel Diatoms and flagellates “Erdschreiber” Flagellates Dinoflagellates Phaeodactylum. Based on Miquel Diatoms Dinoflagellates Diatoms Phaeodactylum E 3, E 13 Flagellates Dinoflagellates Stichococnu Dinoflageliates S, D Photosynthesis survey

29 1

XI. ALGAE

TABLE VI-continued Reference

Designation

Kinne-Diettrich, 1955 Wilson and Collier, 1955 Provasoliet al., 1957

Nordli, 1957a Provasoli, 1958

E3,E13

ASW 111, ASW 8

Guillard and Wangersky, 1958 Kainand Fogg, 1958 Lewinand Lewin, 1960 Boalch, 1961 A,A 1, A 3,A 7 Iwasaki, 1961 Guillard and Ryther, 1962 MA, MAV Droop Von Stosch, 1963 Von Stosch and Drebes, 1964 Jitts et al., 1964 Antiaand Watt, 1965 PPCMM-2

Notes Cyanophyta Gymnodiniumbreve Diatoms and flagellates.Also Schreiber, Feyn, Barker, Sweeney, Goldbeq et al., Spencer, Haxo and Sweeney, Wilson and Collier. Ceratium. Bacterized cultures Ulva Flagellates Asterionella Littoral diatoms, base Ectocarpus Porphyra Diatoms, mainly pelagic See p. 300 of this article. Also Marshall and Orr (1962) Asporagopsis Stephanopyxis and other diatoms Diatoms and flagellates Flagellates

TABLE VII

Undefined marine media (organic) Reference

Designation

Notes Porphyridium

Pringsheim and Pringsheim,

1949 Droop, 1954 Provasoli et al.,1957

E6 ASW 111,

Flagellates Diatoms and flagellates

STP, E 6 Nordli, 1957b Lewinetal., 1958 Droop, 1959 McLaughlin and Zahl, 1959 Hulbert et al., 1960 Boalch, 1961 Rahat and Reich, 1963 Antia and Kalmakoff, 1965

I

OX 7

Ceratium. Algal flour extract supplement Cylindrotheca Oxyrrhis marina,axenic culture Zooxanthellae Katodinium Ectocarpus Prymnesium p a m m Hemiselmis virescens

292

M. R. DROOP

TABLE VIII Defined marine media (mineral) Reference

Designation

Hutner, 1948 Lewin, R.A. ,1954 Ryther, 1954 Droop, 1955 Provasoli et al.,1957 Kainand Fogg, 1958 McLaughlin, 1958 Provasoli, 1958 Aldrich and Wilson, 1960 Boalch, 1961 Stewart, 1962 Jones, 1962 Takano, 1963 Provasoli, 1963 Provasoli and McLaughlin,

1963 Paasche, 1964 McLachlan, 1964 Taylor, 1964 Takano, 1965 Rahat and Jahn, 1965 Lewin, 1965

Notes

Phaeodoctylum Stichococcus Nunnochloris, Stichococcus S 36 Skeletonema cortatum ASP, ASP 2, Diatoms and flagellates.Also ASP 6,S 32,S 36 Ryther, Lewin Asterionella Chrysomonads ASW 8 Ulva Gymnodinium breve C, E, F,G Ectocarpus Calothrix, Nostoc Porphyridium Diatoms ASP 1, ASP 2, Seaweeds ASP 6,ASP 7, ASP 12,ASM MGC, AC, Dinoflagellates DV Coccolithushwlii Discusses various defined media CF 1 Flagellates, based on ASP 6 Diatoms Prymnesium Cylindrotheca

TABLE IX Defined marine media (organic) Reference Hutner et ul.,1950 Provasoli and Pintner, 1953 Lewin, 1955 Provasoliet al., 1957 Droop, 1958 Pintner and Provasoli, 1958 Droop, 1959 McLaughlin and Zahl, 1959

Designation

RC, DC, S 46 etc.

s 50 S 68,S 69

Notes

DuMliella Gyrodinium Prasiola Diatoms, flagellates. Also Vishniac, Lewin, Vishniac and Watson, and others Monochrysis lutheriand others PhormMTiumpersecinum Axenic Oxyrrhis m r i n a Symbiodinium and other Zooxanthebe

293

XI. ALGAE

TABLE IX-continued Reference

Designation

McLaughlin et al.,1960 Droop, 1962 Provasoli and McLaughlin, 1963 Soli,1963 Lewinand Lewin, 1967 Droop, 1968

Notes Dinoflagellates Skeletonema costatum Dinoflagellates Pelagic diatoms Apochloroticdiatoms Monochrysip, dry mix

S 76

S 88

TABLE X Some trace-metal solutiops Reference Hoagland and Snyder, 1933 Amon, 1938 Chu, 1942 Burkholder and Nickell, 1949 Hutnerand Provasoli, 1951 ProvasoliandPintner, 1953 Hutneretal., 1953 Sager and Granick, 1953 Allenand Amon, 1955 Droop, 1955 Hutner etal., 1956 Provasoli et al.,1957 Droop, 1958 Gdron

Designation

Notes

A-Z A4,B7,C 13

See also Arnon (1938)

S1 42

TM 2 45

S 1,T M 2, SW 2 etc.

Also Vishniac, Gjvodinium

TM 11B See Hughes et al. (1958) TABLE XI

N.E.R.C. Culture Centre of Algae & Protozoa, Cambridge, Curator E. A. George)

Two useful stock media (from the J

+

“e s” Soil extract Liver extract (OXL25) Bactotryptone(Difco) Glucose

mo3

KzHPOi MgSOi. 7Hz0 H2O

t Or to a light straw colour.

50 mgt

.. .. ..

200 mg

20 mg 20 mg 1.O litre

“Ochrmonas” a .

1.0g

1-05

1-0g

..

..

.. 1-0litre

294

M. R. DROOP

TABLE XI1 Examples of defined media (amountsper litre) Emerson and Lewis (1942)

1-0 g

KNOs NH4C1 KHaPO4 CaCl2 FeSOa

50 mg 1.og

20 mg 4 mg (Chu, 1942) No. 10.

40 mg

Ca(N0s)z KpHP04 FeCls

5 or 10 mg 0.8 mg 25 mg

MgS04 NazSiOs

25 m g (Rodhe, 1948) No. VIII

60 mg 5.0 mg 5.Omg

Ca(N0s)z KeHPOet

{Mz!2° NazSiOs f Autoclaved separately

1-0mg 20 mg

Tumiyu et al. (1953) 5g

1.25g 2.5 g 3 . 0 mg 1 -0ml

1.O ml (Provasoli et al., 1957) ASP 6 (marine)

NaCl MgS04.7HeO CaClz. 6H2O KC1 NaN03 Dipotassium glycerophosphate NaeSiOa. 9He0 Tris buffer P 8 metals Vitamins

24 8 8.0 g 0.8 g 0.7 g 0.3 g 0.1 g

70 mg 1.0 g 10 ml 1.O mi

295

XI. ALGAE

TABLE XII-continued

(Droop, 1958) S 50 (marine)

15 8 2.5 g

NaCI MgCl2.6HaO CaS04.2H20

0.5 g

KC1 0.4 g KN03 0.1 g KzHPO4 10 mg 0.5 g G1ycylglycine 0.25 g Glycine Sodium EDTA (Na2 ethylenediamhe tetraacetate) 50 mg Vitamin B12 0.1 Pg Thiamine 1*O mg “SW 2” Metals 5 ml “TM 11B” metals 10 ml (Humsr et ai., 1956) Euglena, Vitamin KHzPOs MgSOi .7HzO CaC03 Sucrose L-Glutamic acid DL-Aspartic acid DL-Malic acid Glycine Ammonium succinate Thiamine HC1 Metals 45 (PH 3.6)

, assay, dry mix

0.3 g 0 - 4g 80 mg 15 8 3.0 g 2.0 g 1-0g

2.5 g 0.6 g 0.6 mg 22 mg

(ProvasoIi and Pintner, 1960) VOIVOX Ca(NOd2 MgS04.7HeO Disodium glycerophosphate(hydrated) KCI GIycylglycine Vitamin Biz

Biotin P IV metals (Droop, 1961b) S 66 NaCl MgCla. 6Ha0 KCI CaS04.2H20

KNOs KBHPOI

296

M. R. DROOP

TABLE XI I-continued Glycylgly cine Glycine Citric acid Vitamin Bia Thiamine “TM 11B”metals

0.5 g 0-25 g 40 mg 0.1 CLg 0.1 mg 10 ml

TABLE XI11 Some trace-metal and vitamin solutions

(Amon, 1938) A 4 (used 1-0mli/litre) 2.86 g 1.81 g 0.222 g 79 mg 1-0litre

H3B03 MnC12.4H20 ZnSO4.7HaO CuSo4.5Hz0 H20 (Amon, 1938) B 7 (used 1 * O ml/litre) 85 % Moor NH4VOs Cr2Ka(S0&, 24H20 Nis04.6H~O Co(N03)2.6H20 Na2W04.2H20 Ti0 :(COO. COOK). 2HzW N/10 Has04

17-6mg 23 mg 96 mg 44.8 mg 49-4 mg 17.9 mg 73.7mg 1 - 0 litre

t

Precipitated with NH40H, filtered, and dissolved in N/10

(Amon, 1938) C 13 (used 1 -0ml/litre) AI(SOd3 As03

CdC12 SrS04 HgCh PbCh LiCl Rb2S04 NaBr KI NaF Nan%Oa Be(N03)2.3HzO HzO

31.7mg 6 - 6 mg 8 - 2mg 10.5 mg 6.8 mg 6 - 7 mg 30.6 mg 7 - 8 mg 6.4 mg 6 . 5 mg 11.1 m g 11a9 mg 0.104g 1 * O litre

XI. ALGAE

TABLE XIII-continued (Provasoli et al., 1957) P 8 metals (used 10 ml/litre) Na3 Versenol (Na3 hydroxyethylethylenediamine 3n triacetate) 200 mg Fe (as Cl-) 50 mg Zn (as C1-) 100 mg Mn (as Cl-) 1-0mg Co (as C1-) 2.0 mg Cu (as Cl-) 200 mg B (as H3BOs) 50 mg Mo (as Moor) 1.O litre Ha0 (Droop, 1958) TM 11B (used 10 mlllitre) 50 mg Fe (as sod2-) 5.0 mg Mn , I 0 - 5 mg Zn 0.5 mg c u 8, 0.05 mg co 0.05 mg Mo (aahoo4) 1-0litre N/10HzS04 (Droop, 1958) SW 2 (used 5 ml/litre) 6-5g KBr 1.3g SrCla. 6HzO 50 mg AlCls .6H& 20 mg RbCl 10 mg LiCl H a 0 5 mg KI 1 -0 litre Ha0 (Prowasoli and Pintner, 1960) P 4 metals (used 3-4 ml/litre) HO EDTA (hydroxyethylethylenedi-e triacetic acid) Fe (as Cl-) 40 mg Mn (as Cl-) 10 mg Zn (as C1-) 5.0 mg I-Omg Co (as Cl-) 5.0 mg Mo (asMoO4) 1 litre Ha0 (Hutnm et al., 1956) Metah 45 (dry mix) (used 22 mg/litre) FeS04(NH4)aS04.6HaO 14 mg ZnSOr. 7 H z 0 4.4 mg 1.55 mg MnS04. H a 0 cuSo4. 5Ha0 0.31 mg 0.48 mg cosO4.7Hzo HsB03 0.57 mg (NHo)eMo.rOao.4Hao 0.64mg Na3V04.16HaO 93 Pg

.

297

298

M. R. DROOP

TABLE XIII-continued

(Proerasoli et al., 1957) P 8a vitamins (used 1 .O mlllitre) Thiamine Biotin Vitamin Biz Folic acid p-Amino benzoic acid Nicotinic acid Thymine Choline dihydrogen citrate Inositol Putrescine 2HC1 Riboflavin Pyridoxamine 2HC1 Orotic acid Folinic acid Calcium pantothenate HzO

0.2 g 0.5 mg 50 P g 2.5 mg 10 m g 0 - 1g 0.8 g 0.5 g 1.0g 40 m g 5m g 20 mg

0.26 g 0.2 mg 0.1 g

1 litre

A. Soil-water media Biphasic media, introduced by Pringsheim (l946), have been used extensively for difficult freshwater species and for enrichment cultures of heterotrophs. Good, but unfertilized, calcarious soil gives the best results. It should be sieved and air-dried to remove animal remains and roots. The proportions are about 0.5 g to 10 ml of water in test tubes. Since autoclaving releases toxic materials from soil, the usual practice is to pasteurize the media at 100°C on three successive days. Supplements may be added to soilwater media: calcium carbonate if the soil is poor in alkali; phosphorus and nitrogen for species that tolerate higher concentrations of these elements. Pringsheim used the poorly soluble magnesium ammonium phosphate as a source of three essential elements. Ten to twenty milligrammes is quite enough for either enrichment, and should be placed in the tube before the soil. Enrichments for heterotrophs include starch or a grain of pearl barley or wheat. Pringsheim also used peaty soil for acidophiis. The effectiveness of soil-water media derives from the buffering properties of soil, which regulate both pH and the supply of nutrients, and the presence of a stable microbial population helps to create a naturalistic environment. Biphasic media are invaluable for morphological and taxonomic studies as they preserve morphological features, particularly of Chlorophyta, better than other media. The technique is not suitable and has thus found little application in the marine field.

XI. ALGAE

299

B. Soil extract (Tables 11, 111, VI, VII and XI) Originally introduced by Pringsheim (1912), extracts of soil have become something of a panacea for cultivating both freshwater and marine algae, and they can be autoclaved. Soil extract serves as a source of major and minor nutrients, vitamins and trace metals, and although the buffer capacity and poise are slight and heavy metal complexing is weak, humic acids are still the most effective way of providing these services for many difficult species. On the other hand, oceanic planktonic forms do not well tolerate humates derived from soil and concentrations have to be kept low (not more than 10-20 mg/litre). The Cambridge medium known as “e + s” is the basis of many formulations. It can be used as written (Table XI) or diluted with up to four volumes of distilled water, artificial or natural seawater, or even an organic medium such as the Cambridge “Ochromonas” medium. Soil-extract media, natural seawater and organic media are incompatible in the autoclave, so mixing should be carried out after cooling. It is convenient to carry autoclaved tubed stocks of “e + s” “Ochromonas” (or their equivalents), natural and artificial seawater and distilled water for mixing in the proportions required. The “Ochromonas” medium has a pH of between 5 and 6 when freshly prepared; that of “e+s” will be between 7 and 8, while that of fresh seawater lies between 8 and 8-5. A soil extract of lower pH can be prepared by infusing peat, or alternatively by acidifying neat soil extract (to pH 5-6) and using the filtrate. Preparation of soil extract Weak extracts can be prepared simply by infusing soil with boiling water. A preferable method is to extract with sodium hydroxide, filter clear and freeze-dry. A litre of black woodland soil is autoclaved (1 h, 15 lb) with 1-2 litres of water and 3 g NaOH, and allowed to stand overnight. Further amounts of hydroxide can be added if desired until extraction is complete, but the soil should not be autoclaved a second time. Extraction can be taken to be almost complete when the pH of the mixture remains above 10 on standing overnight. The supernatant liquid is decanted and filtered clear. This is best effected by allowing the liquid to percolate without pressure through its own clay bed supported on a Whatman paper in an ordinary oonical glass funnel. The filtrate is returned until such time as it shows no Tyndall effect. (The rate of flow will then be about a drop in ten seconds.) This extraction can yield 20-40 g soluble solids per lifre and a negligible quantity of clay colloids. The filtrate is freeze-dried and used as required.

C. Enriched natural water (Tables 11, 111, VI and W) Unenriched natural waters seldom contain sufficient nutrients for

300

M. R. DROOP

anything but the lightest growth by culture standards; in some ecological and nutritional investigations, however, enrichments have for the sake of verisimilitude to be kept to a minimum. Then there is considerable advantage in using the balanced natural medium as a base, as in some of Chu’s media prepared with lake water (Chu, 1942). Owing to the difficulty of preparing a satisfactory naturalistic artificial water, natural seawater has formed the basis of many media for marine algae from the complicated, but now completely outmoded, Miquel’s medium to the sophisticated enrichments of vitamins, trace metals, pH buffers and chelating agents, as used for instance by Guillard (Guillard and Ryther, 1962). A simple enrichment (MAV) suitable for bacterized cultures of a variety of phytoplankton has the following composition per litreKNOs KzHP04

FeSO4.7HzO MnSO4. HzO Sodium EDTA Vitamin Blz Thiamine

This nutrient solution is used at the rate of 1 ml to 1 litre of seawater. It is autoclaved with a little distilled water and added to the seawater after sterilization. If desired a few mg of soil extract may be included with the nutrients. When seawater enrichments include soil extract the media are referred to as “Erdschreiber”. The original Erdschreiber was due to Fgyn (1934). Many laboratories prefer to let the seawater age for a couple of months to allow time for the organic constituents to be mineralized by microbial action. More recently a more complete mineralization has been effected in a far shorter time by the action of ultraviolet light (Hamilton and Carlucci, 1966; Armstrong, Williams, and Strickland, 1966; Armstrong and Tibbitts, 1968).

D. Defined media (Tables IV, V, MII, Ix and Xn) In a sense there is no such thing as a defined medium, for although one may determine with any desired degree of accuracy the purity of individual constitual components, the physicochemical interactions between them, except in the simplest cases, defy analysis. Furthermore, once cells have been introduced the medium must lose all claim to being defined. The point is by no means merely academic. All media contain sources of N, P, Mg, Ca, S and Fe. Either nitrate or ammonium nitrogen or both are used. Ammonium is to be preferred on

XI. ALGAE

301

purely theoretical grounds as, given suitable conditions, it is usually assimilated in preference to nitrate and with the expense of less energy. On the other hand, ammonium salts can be toxic even in the mildly alkaline conditions preferred by many algae, while their assimilation causes pH to drop beyond the limits tolerated. Nitrate is, therefore, the safer source. Sulphur is always introduced as sulphate, and phosphorus usually as orthophosphate. The insolubility of calcium and iron phosphates occasionally creates difficulties, especially in marine media having a high calcium content. More serious is the insolubility of the ferric oxide to which all iron tends in aerobic media, and few algae can be grown in defined media lacking organic complexing agents. The first chelator, other than soil extract, to be introduced was citric acid, which in many ways is still the most satisfactory. Non-metabolizable chelators, such as EDTA, nitrilotriacetic acid (NTA), hydroxyethylethylenediaminetriacetic acid (HOEDTA), etc., have however largely replaced citrate. With the use of strong chelating agents to control iron there is more than a chance of causing deficiencies in other essential heavy metals, which are therefore introduced with the chelator. Iron is often simply added as citrate or versinate (Fe EDTA). If this is done there should be at least a tenfold to hundredfold molar excess of chelator over metal, depending on the chelator and the salinity of the medium (Droop, 1961a). It is convenient to prepare a “chelated metal mix”, a number of examples of which will be found in the publications of Hutner, Provasoli and Droop and elsewhere throughout the current literature. Hoagland’s and Arnon’s solutions are used extensively by physiologists (Tables X and XIII). Stock metal solutions keep better if acidified. Some form of soluble sodium silicate is employed in diatom cultures; waterglass is most convenient and is satisfactory provided the inorganic impurities which it presumably contains can be tolerated. For an addition of up to 100 mg/litre the pH of the culture solution should be lowered to between 4 and 6 before adding the waterglass and adjusted to the required pH afterwards. Too low a pH causes silicic acid to precipitate, while too high a pH brings down medium components. In any case the silicate should be stirred in slowly from a 10% solution, Marine media (Provasoli et al., 1957) require an artificial seawater addition. This need not be particularly naturalistic for algae. Indeed, advantage is to be gained by lowering the Ca and Mg concentrations, owing both to their effecton the heavy-metal equilibria mentioned above and to the adverse solubility products of calcium carbonate and calcium phosphate. Salinities of defined marine media lie between 15 and 25%. In general, the higher the salinity the more difficult it is to devise a satisfactory artificial seawater. Culture media, or rather the iron and phosphorus components thereof, are damaged by autoclaving if the pH is allowed to rise too far in the auto-

302

M. R. DROOP

clave, as happens if p H buffering is limited to the natural bicarbonate system. Organic buffers prevent this. T h e maximum buffer capacity should be at a little above the required pH setting: Tris buffer, di- or tri-glycine for pH 8; ethanolamine, maleic acid for pH 6.5;succinic acid for p H 5.5; fumaric acid for p H 4.5; and for p H between 9 and 10 various amino-acids, e.g., valine or glycine. p H is conveniently adjusted before autoclaving with 10% HC1 or NaOH. Some culture media can be prepared in bulk as a “dry mix”. Marine media contain sufficient bulk as to require no additive, but freshwater media should be triturated with a convenient quantity of insoluble powder. Powdered magnesium silicate serves very well, though it should be borne in mind that this substance finds use in absorption chromatography. Sugar alcohols such as mannitol or sorbitol are sometimes used; they are soluble though generally not metabolized by algae, but they are osmotically active. I n the preparation of dry mixes it is necessary to ensure that all the components are thoroughly mixed. This can be done dry in a mortar or as a slurry which is freeze-dried afterwards. T h e latter has the advantage that minor components are easily dispersed by solution in the liquid phase, The secret of good mixing, either dry or as a slurry, is to mix the larger bulk gradually into the smaller, and not vice versa. Unless the medium contains liquifying organic acids it is best to prepare slurrics on the acid side; the colour change of the ferrous-ferric equilibrium gives an indication.

1. Nutrient concentration Physiologist’s media tend to be limited in the range of algae they will support because the nutrients supplied by them tend to be restricted and too high in concentration while, on the other hand, truly naturalistic media support impractically low cell populations. By way of compromise, the following components Fcr litre of medium will serve as a point of departureGlucose Anhydrous sodium acetate N (as Nos, NH4, or amino)

Mg Ca

P S Fe Mn cu Zn co Mo buffer (“tris” or diglycine) chelator (EDTA or citric acid)

10 g 1.0 g 10 mg 5.0 mg 5.0 mg 1.0 mg 1.0 mg 0.5 mg 50 Pg 5.0 Pg 5.0 Pg

0.5 PU 0.5 Pg 0.5 g 50 mg

XI. ALGAE

303

2. Preservative for stock solutions Stock solutions may be kept free from micro-organisms with a drop of the following volatile preservative (Hutner and Provosoli, 195 1) : o-fluorotoluene, n-butyl chloride and ethylene chloride in proportion 1 : 2 : 2.

E. Solid media Many algae, for instance most chlamydomonads, Chlorococcales and pennate diatoms, can be carried on agar slopes; others, notably naked forms and pelagic species, do not tolerate solid media. T h e advantage of a solid medium is that adventitious contamination is localized and can be detected before it has spread throughout the culture so that regular sterility testing is not necessary. Agars are not suitable for nutritional studies requiring the use of defined media. Many of the mineral and organic formulations are suitable for use with agar and, unless the medium has an acid or very alkaline reaction, it is not necessary to autoclave the nutrients apart from the agar. Algae are more hydrated than bacteria and mostly do better in softer gels. Difco Bacto agar will set firm at a concentration of 0.9-1-0%, although care has to be exercised not to break the surface when making inoculations with a wire loop. “Sloppy” agar media (with approximately 0.50/, agar) are sometimes used for delicate species. They provide a gradient in oxygen and carbon dioxide tension downwards from the surface.

VI. SOME SPECIAL METHODS A. Continuous systems An obvious limitation of all batch cultures is that conditions must alter continually with growth of the culture, unless cell numbers are extremely small relative to the culture volume and nutrients available. This becomes very evident if mediuw and cell constituents are monitored during the life of a cuIture. Furthermore, an apparently constant specific growth rate is an insensitive indicator of constant conditions, for nutrient depletion can proceed quite far before it is reflected in a diminished growth rate (Droop, 1968; Tempest, this Series, Vol. 2). The mean physiological state of a population can, however, be stabilized by maintaining a constant rate of cell increase in constant conditions by means of some form of continuous-flow culture. One could achieve approximately constant conditions by increasing the volume of the culture with fresh medium in proportion to cell growth, as indeed Myers and Clark (1944) did in their original “turbidostat”. More practicable, however, is to

304

M. R. DROOP

employ a constant volume device and either regulate the flow to keep a constant cell density-the normal turbidostat (Phillips and Myers, 1954; Fogg et al., 1959; Munson, this Series, Vol. 2 ) 4 r keep a constant flow, when the cell density will equilibrate and remain constant-the so-called “chemostat” (Novick and Szilard, 1950; Monod, 1950; Evans et al., this Series, Vol. 2). The difference between the two systems is of practical rather than fundamental importance ;the operation of either fits the same mathematical framework with respect to limiting nutrients. I n both systems the true controlling factor is the limiting concentration of the medium in which the cells are growing. Under conditions of saturation with respect to all nutrients other factors come into play; these in any case determine the saturation levels for the various nutrients. These continuous culture equations are derived by Tempest (this Series, Vol. 2). A set of empirical equations can also be written for light-limited cultures; they differ from the nutrient equations because the influx of light is not dependent on the dilution rate of the culture, and there is mutual screening by the cells-

L = -1 pnh

&+i

(1)

which combine and reduce to

where i is the light incident per unit cell mass I the incident light on each cell in the absence of mutual screening, and Ki and k, are constants, and ,u is the specific growth rate and ,urn the maximum specific growth rate. Equation (3) has a unique solution for any of the three variables x, I and p, in terms of the other two. Now the controlling factor is I , and p can be altered by manipulating x, or x altered by manipulating ,u (keeping I constant the while). When carbon dioxide is the limiting nutrient similar considerations apply: we have, under steady-state conditions-

r being the rate of entry of carbon dioxide into the system, and Y the yield constant. T h e relation between Y and the concentrations of the various

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dissolved components of the carbon dioxide system is rather involved, but for any one component may be approximately statedr = r r n (I-:), so is the steady-state concentration in

the absence of cells and depends principally on the COZ tension of the aerating gas and the base content of the culture medium. (See the equation on p. 282.) Substituting, from equations (4)and ( 5 ) and the usual equation relating growth rate to limiting nutrient concentration1‘ = prn

(&)

we get-

(a quadratic, but only one of the roots has any physical meaning).

It will be noticed that both with light and carbon dioxide limitation, because the entry of the “nutrient” is independent of medium flow, there is no upper limit to x as ,u approaches zero. In practice other factors come into play. Control through x (turbidostat) is most sensitive when r is rather small and flow rates fast, whereas control through ,u (chemostat) is most sensitive when rates are small and numbers large (Herbert, 1959). Thus, the desired operating point is the factor deciding which of the two systems should be adopted. In the turbidostat cell density is sensed by a photocell which operates a solenoid through a relay to open a valve in the medium inflow line, so that when the cell density reaches a given value the culture is diluted with fresh medium. The longer the light path through the culture between photocell and energizing lamp the lower can the operating ceI1 density be set, a point of some importance in ecological applications (Maddux and Jones, 1964; see Munson, this Series, Vol. 2). In-the chemostat medium flow is controlled either by a peristaltic pump or other metering device. Capillary-gravity systems are seldom used now because of their tendency to block and their sensitivity to temperatureviscosity fluctuations. The necessary condition for specific growth rate to equal dilution rate in the reaction vessel is that the culture and medium should be mixed and of course there should be no wall growth. Failure to meet these conditions ‘3

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results in the non-equivalence of dilution rate and the specific growth rate (Herbert et al., 1956). Vigorous aeration and stirring and careful design of the reactor are necessary. There should be no corners or dead spaces for settlement to occur. A completely filled spherical vessel with acentric aeration is fairly efficient, but of course is limited to use with fairly low cell densities if light limitation is to be avoided. Apparatus for use with heavy cell densities is usually constructed on the annulus principle, the culture being constrained in a thin layer between two concentric cylinders with a water jacket between the culture and a third concentric cylinder with the fluorescent lamp at the centre (Myers and Clark, 1944, Fogg et al., 1959).

FIG.3. 250 ml chemostat. Air pump air saturator, and inoculating port on the reactor are not shown. -, culture medium; --, air; .-.-. , medium and air; SV, clamping solenoids operated by a timer: SF, sterilizing air filter. (Reproduced from Droop (1966) by permissionof the Council of the Marine Biological Association of the United Kingdom.)

It is not proposed to go into details of construction and refinements of the various apparatus used. T h e reader is referred to the authors mentioned above and to articles by Evans et al., Munson, and Ritica, this Series, Vol. 2. Useful information will also be found in Burlew (1953); MQlek (1958); Tunevall(1959); and MBlek et al. (1964).

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Fig. 3 illustrates a cheap but effective chemostat constructed of glass and silicone rubber, used to study the kinetics of vitamin BIZ limitation in a marine chrysophyte (Droop, 1966, 1968).

B. Synchronous cultures The harvest from continuous cultures grown in continuous illumination is, or should be, random with respect to the division cycle of the cells, SO that cell composition determined from chemostat or turbidostat material tells little about the composition of individual cells and nothing about its variation throughout the division cycle. T o overcome this methods have been developed for synchronizing the division cycle of whole populations SO that the resultant harvest is ideally homogeneous with respect to the cell division cycle. The technique for synchronizing continuous cultures is yet in its infancy. Synchronization is brought about either by mechanical selection of cells in a particular age, e.g., small recently divided cells, or by subjecting a random population to an environmental stress that halts the cycle at a particular point. In the first category one may list differential centrifugation, differential filtration, manual selection of dividing cells, spores, etc. In the second category are lowering the temperature or subjecting the cells to an alternating light and dark regime. The use of an environmental stress to initiate a cycle or entrain a natural rhythm is open to the possible objection that events thereafter may not necessarily be the same as would be found in individual cells of a steady-state culture. The details of methods used by the Japanese school, which rely on an initial homogeneous population obtained by differential centrifugation, will be found in Tamiya et al. (1961) and Morimura (1959). The separated cells of Chloreila,more than 80% of which will be 3 pm in diameter, are matured in the light, then incubated in the dark, when they divide synchronously. The daughters so formed can be used to start the cycle over again. This is one of the variety of regimes used by this school, the so-called DLD Cycle. In programmed light and dark regimes, originally developed by Lorenzen at Marburg (for references and details see Lorenzen, 1964), a random culture is subjected to a regular light and dark alteration, the details of which depend on the species and also on temperature and light intensity, growth occurring during the light and cell division during the dark period. The culture is diluted after each cycle with an appropriate volume of medium to keep a more or less constant cell count. Complete synchrony is claimed by this method. A very useful small review will be found in Hoogenhout (1962), while Tamiya (1966) and Pirson and Lorenzen (1966) discuss the whole question,

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its achievements, pitfalls and potentialities at some length. Zeuther (1964) and Cameron and Padilla (1966) also contain much useful information. For a general discussion of methods of investigating the growth cycle of micro-organisms see Helmstetter, this Series, Vol. 1. REFERENCES Aaronson, S., and Baker, H. (1959).J. Protozool., 6, 282-284. Aaronson, S., and Scher, S. (1960). J. Protozool., 7, 156-158. Aldrich, D. V., and Wilson, W. B. (1960). Biol. Bull. mar. biol. Lab., Woods Hole., 119,5744. Algeus, S. (1946). Bot. Notiser, 1946, 129-278. Allen,E. J.,andNelson, E, W.(1910).J.mm. biol. Ass. U.K.,8,421474. Allen, M. B. (1952). Arch. Mikrobiol., 17, 34-53. Allen, M. B. (1959). Arch. Mikrobiol., 32, 270-277. Allen, M. B., and Arnon, D. I. (1955). Pl. Physiol., Lancaster, 30,366-372. Allison, F. E., Hoover, S. R., and Morrison, H. J. (1937). Bot. Gaz., 98,433-463. Anderson, E. H. (1945).J. gen. Physiol., 28,287-327. Antia, N. J., and Kalmakoff, J. (1965). Fish Res. Bd Canada, Manuscript Report Series., No. 203, 1-24. Antia, N. J., and Watt, A. (1965). J. Fish Res. Bd Can., 22, 793-799. Armstrong, F. A. J., Williams, P. M., and Strickland, J. D. H. (1966). Nature, Lond., 211,481-483. Armstrong, F. A. J., and Tibbitts, S. (1968). J. mar. biol. Ass. U.K., 48, 143-152. Arnon, D. I. (1938). Am.J. Bot., 25, 322-325. Amon, D. I., Ichioka, P. S., Wessel, G., Fujiwara A., and Woolley, J. T. (1955). Physiologia PI., 8, 538-551. Baker, H., Hutner, S. H., and Sobotka, H. (1955). Ann. N.Y. Acad. Sci., 62, 351-376. Baker, H , Frank, O., Matovitch, V. B., Pasher, I., Aaronson, S., Hutner, S. H., and Sobotka, H. (1962). Analyt. Biochem., 3, 31-39. Barber, F. W., Baile, D. L., Troescher, C. B., and Huhtanen, C. N. (1953). Ann. N. Y.Acad. Sci., 56, 863-869. Barker, H. A. (1935). Arch. MikrobioZ.,6,157-181. Barry, S., and Barry, N. C. (1962). J. Protozool., 9, 395-400. Beijerinck, M. W. (1890). Bot. Ztg., 48, 725-786. Bendix, S., and Allen, M. B. (1962). Arch. Mikrobiol., 41, 115-141. Boalch, G. T. (1961).J. mar. biol. Ass. U.K., 41, 287-304. Bold, H. C. (1942). Bot. Rw.,8, 69-138. Bristol Roach, B. M. (1926). A m . Bot., 40,149-201. Buetow, D. E., and Padilla, G. M. (1963).J. Protozool., 10, 121-123. Burkholder, P. R.,and Nickell, L. G. (1949). got. Guz., 110,426-437. Burlew, J. S. (1953). (Ed.) “Algal Culture: from Laboratory to Pilot Plant”. Publs. Carnegie Instn., No. 600. Callely, A. G., and Lloyd, D. (1964). Biochem. J., 90,483489. Cameron, 0. L., and Padilla, G. M. (1960). (Eds.) “Cell Synchrony”. Academic Press, New York. Chomey, w., scully, N. J., Crespi, €3. L., and Katz, J. J. (1959). Biochim. biophys. Actu, 37,280-287. Chu, S. P. (1942). J. Ecol., 30, 284-325.

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Rodhe, W. (1948). Symb. bot. upsal., 10 (1)) 1-149. Ryther, J. H. (1954). Biol. Bull. mar. biol. Lab., Woods Hole., 106, 198-209. Sager, R., and Granick, S. (1953). Ann. N . Y. Acad. Sci., 56, 831-838. Schmidt, R. R. (1961). Expl. Cell Res., 23, 209-217. Scotten, H. L., and Stokes, J. L. (1962). Arch. Mikrobiol., 42, 353-368. Schreiber, E. (1927). Wiss. Meeresunters, N.F. 16 (10). 1-34. Soeder, C. J. (1960). FZoru,Jena., 148, 489-516. Soeder, C. J., Muller, I., and Ried, A. (1962). Vortr. GesGeb. Bot., dt. bot. Ges., N.F. 1,195-200. Soli, G. (1963). I n “Symposium on Marine Microbiology” (C. H. Oppenheimer, Ed.), pp. 260-274. C. C. Thomas, Springfield. Sorokin, C., and Krauss, R. W. (1958). PI. Physiol., Luncaster, 37, 37-42. Sorokin, C., and Myers, J. (1953). Science, N.Y., 117, 330-331. Spencer, C. P. (1954). J. mar. biol. Ass. U.K., 33, 265-290. Spencer, C. P. (1966). Botanicu mar., 9, 81-99. Stewart, W. D. P. (1962). Ann. Bot., 26,439-445. Storm, J., and Hutner, S. H. (1953). Ann. N.Y. Acad. Sci., 56, 901-909. Stout, P. R., and Arnon, D. I. (1939). Am.J. Bot., 26, 144-149. Stross, R. G. (1963). Can. J. Microbiol., 9, 33-40. Sweeney, B. M. (1951). Am. J. Bot., 38,669-677. Sweeney, B. M. (1954). Am.J. Bot., 41, 821-824. Syrett, P. J. (1953). Ann. Bot., N.S. 17,l-19. Syrett, P. J., Merrett, M. S., and Bocks, S. M. (1963).J. exp. Bot., 14, 249-264. Takano, H. (1963). Bull. Tokai reg. Fish Res. Lab., 37, 19-25. Takano, H. (1965). Bull. Tokui reg. Fish. Res. Lab., 44,17-24. Tamiya. H . (1966). A. Rev. PI. Physiol., 17, 1-26. Tamiya, H., Hase, E., Shibata, K., Mituya, A,, Iwamura, T., Nihei, T., and Sasa, S. (1953). I n “Algal Culture: from Laboratory to Pilot Plant” (J. S. Burlew, Ed.), pp. 204-232. Publs. Carnegie Instn., No. 600. Tamiya, H., Morimura, Y., Yokota, M., and Kunieda, R. (1961). PI. Cell. Physiol., 2,383-403. Taylor, R. W. (1964). Occ. Publs. Narragansett mar. Lab., 2, 17-24. Thomas, W. H., Scotten, H. L., and Bradshaw, J. S. (1963). Limnol. Oceunogr., 8,357-360. Tunevall, G. (1959). (Ed.) “Recent Progress in Microbiology”. Almqvist and Wiksell, Stockholm. Von Stosch, H. A. (1963). Int. Seaweed Symp., 4, 142-150. Von Stosch, H. A., and Drebes, G. (1964). Helgolander Wiss. Meeresunters., 11, 209-2 5 7. Watanabe, A., and Kiyohara, T. (1963). In “Microalgae and Photosynthetic Bacteria” (Jap. SOC.pl. physiologists, Eds.), pp. 189-196. University Press, Tokyo. Wilson, W. B., and Collier, A. (1955). Science, N . Y., 121,396395. Wise, D. L. (1959).J. Protozool., 6,19-23. Zehnder, A., and Gorham, P. R., (1960). Can.J. Microbiol., 6,645-660. Zehnder, A., and Hughes, E. 0. (1958). Cun.J. Microbiol., 4, 399-408. Zeuther, E. (1964). (Ed.) “Synchrony in Cell Division and Growth”. Interscience Publishers, New York.

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CHAPTER

XI1

Isolation, Growth and Preservation of Bacteriophages EVEBILLING* Department of Microbiology, University of Reading, England I. Introduction . 11. Isolation A. Growth conditions . B. Platingmethods . C. Separation and inactivation of bacteria D. Isolation from natural sources . E. Isolation from lysogenic and carrier strains F. Purification . 111. Preparation of High-titre Stocks . A. Plate method B. Broth method C. Temperate phages . IV. Preservation References .

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I. INTRODUCTION The aim of this Chapter is to describe, mostly for the novice rather than the specialist, some methods for the isolation, cultivation and preparation of stocks of bacteriophages which should be applicable to avariety of phagehost systems. For the most part they involve simple operations and require no complex apparatus or equipment. Those who seek more specific information about particular phages or their hosts may need to consult original papers; of particular value in this connection is a list of methods for study of specific phage-host relationships given by Eisenstark (1966) in a recent account of various bacteriophage techniques. A further comprehensive source of information is the survey of phage literature, 1917-1958, by Raettig (1958) and its supplement (in preparation). Alimited list ofreferencescoveringavariety ofgenera and species other than Enterobacteriaceae where isolption methods are described is given in Table I. ;*

Present address: East Malling Research Station, Maidstone, Kent, England.

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TABLE I Isolation methods for bacteriophages Bacteriophage Actinomycetes Agrobacterium radiobacter Azotobacter Bacillus sp. Bordatella bronchiseptica Brucella spp. Caulobacter and Asticcacaulis Chondrococcus columnaris Clostridium perfringens Cl. sporogenes Corynebacterium diphtheriae C . jlaccumjaciens Haemophilus injluenzae Lactobacillus spp. Mycobacterium spp. Myxococcus xanthus Nocardia Pasteurella haemolytica Photobacterium phosphorarm Plectonema borganum, a blue-green algae Pscudomonas syringae and related phytopathogens Rhizobium spp. Spirochaeta rosea Haemolytic Streptococcus Streptomyces venezuelae Xanthomanas malvacearum

Reference Anderson & Bradley (1964) Roslyckyet al. (1962) Duff & Wyss (1960) Meynell(l961) Rauch & Pickett (1961) Brinley-Morgan et al. (1960) Schmidt & Stanier (1965) Anacker & Ordal(l956) Kingsbury & Ordal(l966) Guelin (1955) Betz & Anderson (1964) Groman et al. (1958) Klement & Lovas (1957) Harm & Rupert (1963) de Klerk et al. (1965) Sellars et al. (1962). Burchard & Dworkin (1966) Anderson & Bradley (1964) Rifkind & Pickett (1953) Spencer (1960) Schneider et al. (1964) Cross & Garrett (1963) Schwinghamer (1965) Lewin (1960) Kjems (1960) Kolstad & Bradley (1964) Hayward (1964)

Okabe and Goto (1963) describe sources of phages for other plant pathogenic bacteria and further methods for their isolation. For information on selection of phages for typing purposes, see Parker, this Series, Vol. 7. I n practice, many people follow with little modification the methods described by Adams (1959) for coli-phages, and this Chapter must of necessity duplicate much of what is written in his classic handbook. Other valuable sources of practical information on bacteriophage techniques are Eisenstark (1966) and Meynell and Meynell(1965). All methods described here should be applicable to Enterobacteriaceae, but similar methods have been used with success by different workers using a wide variety of host bacteria grown on appropriate media. Failure to isolate a phage at the first attempt may in many cases simply be due to the use of unsuitable source material (see Section 110) but some strains of particular species are more susceptible to phage attack than others. Although isolation of virulent

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phages may often provc easy, specific enrichmcnt for phages with a particular morphology, nuclcic acid content, or host rangc is scldom a practical proposition. The isolation of temperate phages depends on the availability of suitable lysogenic and indicator strains and presents somewhat different problems (seeSection I IE).

11. ISOLATION

A. Growth conditions A medium which gives rapid growth of the host bacterium will normally be satisfactory for phage multiplication. For many non-exacting bacteria Difco, Oxoid or other routine nutrient media are satisfactory. A suitable ionic environment is important for rapid adsorption of phage to its host and in addition some have a specific requirement for divalent cations such as calcium or magnesium. Optimum cation concentrations are not the same for every phage-host system however, and for isolation special provision is seldom made, reliance being placed on the fact that the normal media ingredients are likely to provide an adequate though not necessarily optimal concentration of cations. This could mean, however, that in enrichment procedures for isolation, some phages are at a disadvantage and so unlikely to become dominant. For aerobic organisms and their phages adequate aeration is important for maximum growth in broth cultures; this also holds true for many facultative anacrobes, especially when grown in a medium containing no ferrnentablc carbohydrate, but for isolation the use of liquid media in shallow layers is normally satisfactory. For other operations using liquid media (e.g., propagation) shaking or bubbling devices may be essential unle'ss the medium can be used in very shallow layers. For plating, 15-20 ml of agar medium per 9 cm plate is often adequate, but the optimum depth of medium will depend on the phage-host system. Normally if the host ceases to multiply because of lack of nutrients, phage multiplication will also stop, so an adequate reservoir of nutrients may be particularly important in the case of some phages which form small plaques, to allow time for the plaques to reach visible proportions; up to 30 ml of medium per 9 cm plate may be necessary for some systems but may lead to drying difficulties. Plaque size is also affected by the strength of the agar gel; with a soft agar the phage will diffuse more rapidly and so produce larger plaques. Thus a combination of a soft agar layer containing phage and host overlying a deep layer of normal strength nutrient agar provides conditions likely to give the largest plaques (see agar Iuyer method, page 3 19). Different brands of agar vary in the strength of gel they produce; with Japanese agar, l.S()I is suitable for plating purposes, but with other agars

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l.Oyo is sufficient; for soft agar 0.6-0*7% of agar is the usual recommendation. When suspensions of bacteria are to be made from agar slants, a more uniform suspension is obtained if a firmer agar is used, e.g., 2.0% Japanese agar or its equivalent (Adarns, 1959). The temperature of incubation used for isolation may influence the type of phage selected. One which readily forms plaques at 25°C will not necessarily do so at 30" or 37"C, although the latter temperatures may be nearer the optimum for growth of the host organism. With some RNA phages the converse may be true (Dettori et al., 1963). Mostly however it is appropriate to incubate cultures at the optimum temperature for growth of the host. Time of incubation will depend on the phage-host system, the temperature used and the nature of the procedure; between 18 and 48 h is usual, but plaques may sometimes be visible in 4 h or less on plates, and clearing of broth cultures may also be observed within this period. B. Plating methods Three methods are commonly used for plating phages to give isolated plaques; each has advantages and disadvantages depending on the conditions. All plating should be done on a level surface to ensure an even depth of medium; plate glass or boards covered with black laminate which can be levelled are convenient; the latter forms a good background for observing plaques. Whatever the choice of method, the optimum amount of host inoculum is likely to be of the same order. The aim is to obtain a layer of bacterial cells which can multiply and produce a confluent lawn, not separate colonies; about 108 viable cells per plate should achieve this. The cultures should be in an active state of growth and may be in the form of broth cultures or suspensions prepared from agar cultures. The amount of inoculum required will depend on the culture concerned but it is usually convenient to keep within the range 0.02-0.5 ml per plate.

1. Pour plate method The host suspension is added to the molten agar medium and the mixture poured into a plate. Mixing before pouring gives more even lawns than pouring the medium onto the inoculum in the plate. It may be convenient to add a suitable dilution of phage suspension to the molten mixture before pouring, although phage multiplication may be restricted in the depth of the agar; alternatively the phage suspension can be streaked on the surface of the solidified medium using a loop or serial dilutions plated using a Pasteur pipette or a syringe. Droplets of moisture normally collect on the surface of freshly poured agar plates and may lead to confluence of plaques. To

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avoid this the surface can be dried before inoculation by placing plates in an incubator at 30-37°C for 14 to 2 h with the lids tilted or removed; this may be thought bad practice but bacterial or mould contamination is seldom a problem. However, the warnings of Feary et ul. (1964) about the dangers of laboratory contamination by RNA phages may be appropriate here; also the risk of aerosols from bubbler tubes used for aeration when phage stocks are being prepared in broth cultures (Eisenstark, 1966). When serial dilutions have been plated, it is advisable to spread the drops either by tilting or agitating the plate or by using a glass spreader to ensure that they are completely absorbed before incubation.

2. Surface plating method The surface of the agar medium in the plate should first be thoroughly dried, e.g., by incubating for 2 days closed at 37"C, overnight closed plus 2 h open at 37°C or 14 h open at 55°C. These recommendations err on the side of safety, but drying efficiency can be erratic and may be more difficult to achieve when plastic plates are used. For lawns, 2-3 ml of the suspension can be pipetted onto the surface and the surplus rapidly decanted. This usually gives more uniform growth than spreading a smaller amount of suspension (e.g., 0.2 ml) with a glass spreader. In either case, the liquid should be rapidly absorbed ; this is facilitated by inoculating plates while still warm and by leaving lids tilted until drying is complete. Depending on requirements, dilutions of phage suspension may be incorporated in the host inoculum or spotted onto the plate with a loop or Pasteur pipette when the agar surface has dried. This second inoculum must be absorbed before plates are incubated.

3. Agar layer (ooerluy) method For this method, which is probably used more than any other, phage and host are incorporated and multiply in a thin layer of soft agar which is poured on to the dried surface of a nutrient agar base. The soft agar (see Section IIA) may set prematurely if care is not taken. If transferred from bulk to tubes just before use, this should be done while the agar is still hot and the tubes should be transferred immediately to the holding water bath at 4647°C. A lower holding temperature (e.g., 4042°C) may be desirable for some heat-sensitive organisms (Wishart, unpublished data) which means that even greater care is needed; if tubes are held too long at these temperatures they will start to gel and poor plates will result. The amount of soft agar nprmally used varies from 1.5 to 3.0 ml, but the beginner may have difficulty in covering the surface of the base agar with the smaller quantity and 2.5 nil is satisfactory in most cases.

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Dilutions of phage suspension should be added to the series of tubes of molten soft agar first as the phage is likely to be less hcat-sensitive than the host. A drop of a suspension of host bacteria is added and the contents of the tube gently mixed, avoiding bubble formation. T h e mixture is poured as soon as possible over the surface of the agar plate which is rocked gently to ensure that the whole surface is covered. U p to 1 ml of phage suspension may be added to 2-5 ml of soft agar, providing the host inoculum is restricted to a drop (e.g., 108 cells contained in about 0.05 mi). It is particularly important for this method that both base and soft agar are poured with plates resting on a level surface.

C. Separation and inactivation of bacteria Most bacteria in a lysate are readily deposited by centrifugation leaving the phage in the supernatant, but some viable cells will remain and must usually be removed or destroyed.

1. Filtration Phage may readily adsorb onto the filter material (Seitz, sintered glass or membrane) and considerable or even complete loss of phage may occur, particularly if titres are low. Prewashing of filters with nutrient broth may help to reduce losses, but it is advisable to avoid filtering low titre phage suspensions. A Millipore Type HA filter (0-45pm) or one with an equivalent average pore diameter (apd) is suitable for sterilization of filtrates; for sintered glass filters an apd of 2 p m is recommended (Meynell and Meynell,

1965). 2. Chloroform A convenient way of sterilizing source material or lysates is to add about 0.5 ml of chloroform per 10 ml. This method is not effective against all bacteria and with some source materials the numbers of survivors may be high although they will not necessarily cause difficulty. There is also a risk that some phages will be inactivated, although decanting the phage suspension after a few minutes and aerating to remove remaining chloroform may prevent undue losses (Eisenstark, 1966). Prolonged contact appears to have no adverse effect on viability of stable phages and they may even be stored over chloroform, but there is evidence that certain properties of phages may be altered by exposure to this agent.

3. Heat Most phages are less susceptible to heat than their hosts and with a suitable choice of temperature it is sometimes possible to destroy host bacteria without seriously affecting phage titres.

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D. Isolation from natural sources 1. General Where the source material is a solid, an extract can be made, although this is not essential in enrichment procedures. It is usual to centrifuge extracts to obtain clear supernatants, and chloroform treatment is often applied at this stage to decrease the bacterial content, especially when extracts are plated direct. The necessity for enrichment and the quantity of source material to be used depend on the likelihood or otherwise of an initially high concentration of phage for the particular host. For example, Stolp and Starr (1964) used 50-1OOg soil plus 5 g CaC03 per 100ml of nutrient solution for the enrichment of phages for Xanthomonas spp. ; in contrast, when sewage influent was used for isolation of Pseudomonas aeruginosa phages by Bradley (1964), the presence of between 100 and several thousand phage particles per ml of sewage made enrichment unnecessary and discrete plaques were obtained by direct plating of 0.1 ml to 0.01 ml of sewage per plate. The enrichment method described by Adams (1959) for coli-phages can be modified to suit individual circumstances. He recommends that 1ml of a visibly turbid culture of the host organism and 1 ml of centrifuged pooled sewage be mixed well with 30 ml of nutrient broth and the mixture incubated overnight at 37°C. After enrichment, the mixture is centrifuged ; filtration or chloroform treatment could be detrimental at this stage, but one or other treatment is usually applied before plating with the host organisms for the detection of plaques. The method of plating used to obtain isolated plaques is a matter of choice, but small plaque phages will normally show to best advantage with the agar layer method, To avoid the necessity of making dilutions, a loopful of suspension can be streaked on the dried surface of a pour plate seeded with host in the same way as a streak plate is made for obtaining isolated colonies of bacteria. If a number of preparations are being screened, spotting drops on the surface of a dried, seeded agar plate may be a useful preliminary test for the presence of phage. Sometimes the enrichment culture only contains a low concentration of phage; in such cases plating up to 1 ml of supernatant may be necessary. Presence of phage may also be detected by observing lysis of a log phase broth culture to which supernatant or filtrate has been added, but unless there is a specific requirement for a phage that gives good lysis in a liquid culture and plaque formation is of secondary importance, it is usually more convenient to work with a phage which forms clearly visible plaques; good lysis in liquid cultures and clear plaques do not necessarily go hand in hand. On this basis, however, good plaque formers will be studied at the expense of poor plaque formers and some worthwhile phages may be discarded.

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Phages which form large plaques can present certain difficulties in quantitative work and in phage typing because of the area covered by each individual plaque. Also they tend to be fast lysers and to give low yields when propagated (Eisenstark, 1966). 2. Small DNA and RNA phages These include isometric and rod-shaped (filamentous) phages containing single-stranded DNA and isometric RNA phages. There is now a considerable amount of information on the characteristics of small coli-phages (Zinder, 1965; Hoffmann-Berling et aZ., 1967), but little specific guidance on methods for their isolation; in the past such phages have often been isolated by chance using normal enrichment techniques. Outside the Enterobacteriaceae a filamentous single-stranded DNA phage has been isolated for Pseudomonas aeruginosa by Takeya and Armako (1966), but no details about methods are described ;isometric RNA phages have also been isolated for this species, by Feary et al. (1964) from a culture which was lysogenic and yielded a typical temperate phage in addition, and by Bradley (1966b) using direct plating of untreated sewage influent. Several RNA phages have been isolated for Caulobacter species by Schmidt and Stanier (1965) by enrichment from sewage and pond water. Feary et al. (1964) warn of the care needed with handling RNA phages to avoid contamination of the laboratory with the attendant risk of infection of other cultures. A simple test for the presence of an RNA phage is the incorporation of 50-100 pg of ribonuclease (RNAse) in the top layer of agar-layer plates (Zinder et al., 1963; Bradley, 1966b); this will normally prevent plaque formation by RNA phages but not by DNA phages. The type of nucleic acid in phage may also be determined by a fluorescent staining method (Mayor and Hill, 1961; Bradley, 1966a, b). One property of these phages which has been exploited in their isolation is their small size. Bishop and Bradley (1965) used the following procedure, which is similar to that described by Paranchych and Graham (1962), for the isolation of two new RNA coli-phages. Sewage samples were shaken with an equal volume of chloroform for 10 min. The aqueous phase was then centrifuged at 4000 g for 15 min to remove large debris and 1 ml of supernatant placed on top of a 24ml sucrose gradient (20-25%, w/v) in TM1 buffer. Centrifugation was at 96,000 g (max) for 3 h. One ml portions of successive 2 ml fractions were plated with an F + or Hfr culture in duplicate (one plate containing 100 pg RNAse) using the agar-layer method. With the two samples of sewage used, the third fraction gave high plaque numbers on untreated plates but none on those containing RNAse, and a typical RNA phage was obtained in each case. Paranchych and Graham (1962) used a 4.5 ml 5 4 % sucrose gradient and centrifuged for 30 min

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at 30,000 rpm in a Spinco Model L centrifuge. They found RNA phages in

fourdrop fractions 15-20. A characteristic of RNA coli-phages is that many (if not all) are male specific, Whether this also holds true outside the Enterobacteriaceae remains to be seen. This specificity is associated with adsorption on to sex pili (fimbriae) (Crawford and Gesteland, 1964; Brinton, 1965; Lawn et al., 1967); Pseudomonas and Caulobacter RNA phages have also been shown to adsorb onto pili. This means that for isolation of these RNA phages success can only be expected if the host used in enrichments forms pili of the appropriatetype. At present, such knowledge is seldom available and chance must continue to play a part in most cases outside the Enterobacteriaceae. As with other phages, plaque formation by RNA coli-phages may depend on how the bacteria are grown and tested (Dettori et al., 1963). I n the agarlayer method, the turbidity of plaques varied from clear to very turbid, depending on the host strain used; with the surface-plate method, when a cross streak technique was used for screening of host-cell sex, clearing was not observed at 35"C, was poor at 37°C but good at 42°C.Plaque diameter may sometimes give an indication of the size of the phage but it is not always a reliable guide. In the case of isometric single strand DNA phages, plaque diameters may exceed 5 mm (Bishop and Bradley, 1965). Lawn et al. (1967) have suggested that sex pili in the Enterobacteriaceae fall into two groups, F-like and I-like, according to whether they resemble the sex pili produced by F + strains or those produced by strains carrying colIb factor. With the detailed knowledge that these workers had concerning piliation and the factors carried by different strains of different species, they were able to isolate a filamentous phage which adsorbs onto the tips of I-like but not F-like pili and appears to be restricted in host range to strains forming I-like pili. The phage was isolated from sewage by enrichment with a strain of Salmonella typhimurium carrying a factor which caused it to produce I-like pili. Selection of other Salmonella phages which might have been enriched at the same time was avoided by plating on E. coli K12 carrying the same factor. It remains to be seen how far this type of technique can be applied to other groups of bacteria when more is known about their ability to form sex pili and about factors they carry which determine synthesis of such pili.

E. Isolation from lysogenic and carrier strains Many bacteria carry phage, either integrated as prophage in the host genome (lysogenic strains) or in a non-integrated form (carrier strains); differences between these are discussed by Hayes (1964). A single strain may release more than one phage. Phages which can enter the prophage

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state are referred to as temperate phages as opposed to virulent phages which are unable to lysogenize. The presence of such phages may remain unnoticed in the original host until it is mixed with or plated on a strain (referred to as an indicator strain) which is readily lysed. Both RNA and the larger DNA phages have been demonstrated in carrier strains; true lysogenic strains which carry temperate phage in the prophage state have so far only yielded larger DNA phages. Both carrier and lysogenic strains may produce a low concentration of phage during exponential growth as a result of lysis of a small proportion of cells. The amount of phage released depends on the phage-host system, but it may be influencedby the conditions of growth. I n some lysogenic systems, the majority of cells may be induced to lyse by exposure to ultraviolet light (see under Propagation, Section IV). Temperate phages normally produce turbid plaques because a proportion of cells is lysogenized instead of lysed, but clear plaque mutants which have lost the ability to lysogenize may sometimes be observed. The host range of temperate phages tends to be more restricted than that of virulent phages isolated from natural sources and many cultures may have to be screened if phage lysing a particular strain is sought; there is more likelihood of success if closely related species or strains are used. There are two simple methods for screening large numbers of strains. For the first, broth cultures of potentially lysogenic cultures at the end of their log phase are treated with chloroform and a loopful of each spotted on to plates seeded with the indicator strain; alternatively, supernatants of centrifuged cultures may be used. After incubation, plaques or clear areas may be seen in the areas spotted. Clearing may be a result of bacteriocin activity and dilutions of chloroform-treated or centrifuged broth cultures of these strains should be plated; only phages will produce individual plaques at the higher dilutions. Alternatively, strains may be patched on nutrient agar plates (glass not plastic) and, after incubation, killed by putting about 0-5 ml of chloroform in the lids of inverted plates. The plates are kept closed for about 15 min and then partly open for a further 30 min to allowthe chloroform toevaporate. A layer of soft agar containing the indicator strain is then poured over the surface. After incubation, any strains around which clear areas are observed can be tested as described earlier to distinguish phage from bacteriocin activity. Isolated plaques can be picked and purification can proceed as for virulent phage.

F. Purification Isolation plates may contain a mixture of several phages so, to ensure that a single type is selected, it is advisable to replate at least twice. A well-

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isolated plaque, which should contain progeny from a single phage particle, is stabbed with a sterile wire or toothpick which is then rinsed in about 1 ml of broth. Dilutions of this suspension are replated, to obtain isolated plaques. After a further picking and replating a plaque is picked into sterile broth and this suspension can be used for the preparation of stocks. The number of phage particles likely to be picked up by a wire or toothpick will vary with different phages, but a single 2 mm plaque may contain as many as 107-109 phage particles (Anderson, 1948).

111. PREPARATION OF HIGH-TITRE STOCKS For small quantities a plate method is more often used than the broth method and is more likely to give high-titre stocks. With both methods success may sometimes only be achieved after considerable experience with the particular phage and its host under different conditions;with some phagehost systems, particularly in the case of temperate phages, it may prove impossible to obtain high titres, but with most it should not be difficult to achieve a titre of at least 109 plaque-forming units (pfu) per ml, and with some up to 1012 pfu per ml may be obtained without difficulty. Repeated propagations, particularly in broth, should be avoided because of the risk of selection of mutant phages. A change of host carries the added danger of host-induced modifications.

A. Plate method The surface plate or the agar layer method may be used; 25-30 ml of medium per plate is recommended. With both methods a phage-host mixture which gives nearly confluent lysis at the time of maximum growth of the host is likely to give highest titres because this means that the maximum number of bacteria will have had opportunity to yield phage. A preliminary titration should ensure that this degree of confluence is achieved. For harvesting, about 3 ml of nutrient broth is added and allowed to stand for a period at room temperature; recommendations here by different workers with different phages vary from 20 min to 5 h. The phage should be harvested by decanting or pipetting the extract carefully from the surface preferably without disturbing the agar, because agar removed during this process may be difficult to eliminate later. A second extraction may yield a suspension with a titre almost as high as the first (Wishan, unpublished data). After centrifugation, remaining viable bacteria may be removed by filtration or by chloroform treatment, bearing in mind the attendant risks of both procedures. If desired, chloroform treatment may be applied earlier either by adding to the suspension or by inverting dishes (glass not plastic) for 15 min over 0.5 ml of chloroform in the

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lid. Treatment before centrifugation may cause unlysed cells to release phage and so increase the yield.

B. Broth method Again experience of the phage-host system is necessary for maximum yields as the time of adding phage to host and of harvesting may be critical. Adequate aeration by shaking or bubbling is essential for high yields, but moderate yields may be obtained using shallow layers. Adams (1959) recommends a procedure involving only a single cycle of infection, i.e., enough phage is added to a culture near the end of the log phase to ensure that nearly every cell is infected simultaneously. This means that a fairly high-titre preparation (about 109 pfu/ml) is required to start with. A more economical method is to allow for several cycles of infection, e.g., adding about 103 pfu/ml to a culture containing about 108 cells/ml. Not all phages will give complete clearing of broth cultures, but there will usually be a marked drop in turbidity. Harvesting may be possible within an hour if only one cycle of infection is involved; it should not be delayed too long after lysis has occurred. Chloroform treatment may be applied before or after centrifugation as with the plate method. Although on a small scale the plate method is often preferred to the broth method, for large-scale production of both low and high-titre phage deep culture methods will often be preferred. Whatever method is used for the preparation of stocks, the preparations will contain a considerable amount of bacterial debris. Eisenstark (1966) recommends the addition of host antiserum followed by centrifugation to remove flagella, capsular material and other unwanted bacterial residues. C. Temperate phages With temperate phages it may be difficult to obtain high-titre preparations because on infection of a sensitive host a proportion of cells will be lysogenized instead of lysed. By altering the growth conditions, it may be possible to reduce the frequency of lysogenization, e.g., by raising the temperature of incubation, using a low multiplicity of infection (a phage: bacterium ratio of less than one), or altering some other physiological factor (Jacob and Wollman, 1959). An added difficulty with temperate phages is that stocks may lose activity more rapidly than is the case with virulent phages, so it may be necessary to prepare them immediately before use. If the phage is inducible, exposure to ultraviolet light will cause release of phage from the majority of cells in a culture, but stocks prepared in this way may differ in behaviour from those produced by other means (Meynell and Meynell, 1965).

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1. Cultural methods

A well aerated lysogenic culture will release some phage spontaneously, the amount varying according to the cultural conditions, but the titre of such preparations is likely to be low. An alternative is to use a broth culture of the indicator strain in a way similar to that recommended for virulent phages, but bearing in mind that optimum phage: host ratio may be rather different €or temperate phages. The agar-layer plate method may also be used for preparation of stocks, but because of background growth the extent of lysis cannot readily be assessed as is the case with virulent phages, and only experience may show what conditions give the highest yields. According to Meynell and Meynell (1965) this method suffers from the disadvantage that it selects clear plaque mutants.

2. Iuduction For induction of inducible lysogenic bacteria by exposure to ultraviolet (UV) light, cells are normally centrifuged and resuspended in a phosphatebuffered salt solution. This is because UV light is absorbed by nitrogenous constituents of nutrient broth. If suspensions are too concentrated, cells may shield one another, so it may be advisable to distribute the material in several open glass dishes or watch glasses so that the depth does not exceed 2 mm and to rock during exposure or use a magnetic stirrer. The efficiency of the process also depends on the wavelength of the light and the intensity of illumination. A wavelength of about 253.7 nm appears to be most suitable and is similar to that found most efficient for sterilization of bacterial suspensions and inactivation of viruses (Adams, 1959). Examples of suitable lamps are : Hanovia Chromatolite low pressure mercury lamp, used at a distance of 75 cm (Fry, 1963); 15 W General Electric germicidal lamp at 56 cm (Eisenstark, 1966); 30 W Phillips TUV lamp at 75 cm (Ijavics et aE, 1967). The energy output of such lamps may vary and factors which affect this are discussed by Meynell and Meynell(1965) who also emphasize the care needed to avoid exposure of the eyes either to direct or reflected UV light. After irradiation for about 2 min or longer, depending on the phagehost system, the cell suspension must be transferred to a suitable medium (concentrated to avoid undue dilution) and incubated in shallow layers or with aeration to give optimum conditions for phage multiplication. After about 2 h, if lysis is not apparent earlier, chloroform treatment can be applied and, after centrifugation, the supernatant titrated.

328

EVE BILLING

IV. PRESERVATION T h e stability of a phage will depend on the suspending fluid. I n a medium containing protein or in broth they usually remain viable for long periods when stored at 4”C, but in chemically defined media or in water some are very unstable. Presence of a few ,ug/ml of protein in a medium prevents “surface inactivation” which is most marked when dilute phage suspensions are agitated; in some cases where a salt solution is used the presence of divalent cations such as Mg or Ca at a concentration of 10-3 M may greatly increase stability (Adams, 1959). Other substances which may affect stability in salt solutions rather than in broth are detergents, oxidizing agents, heavy metals and chlorine; it is important to ensure that special care is taken with the cleansing of glassware and the water used for preparing solutions. When stocks are prepared in nutrient broth, crude lysates of virulent phages (if kept free from contamination) will normally survive in screwcapped bottles or other closed containers for months or years at4”C,although titres will gradually decrease. Storage over chloroform is possible in many cases; this avoids the risk of growth of contaminating bacteria if containers are to be opened frequently. Clark (1962) compared several methods for preserving crude Iysates of a variety of phages but none appeared to have any advantage over simple storage at 4°C in screw-capped vials. Losses on freeze-drying were sometimes high. Keogh et al. (1966) found that phages of lactic streptococci had a high degree of stability when stored at - 18°C after quick freezing at about -7O”C, although in many cases storage at 4°C was equally satisfactory. Alternate freezing and thawing may affect the viability of some phages but there was little evidence of this in the case of these streptococcal phages. REFERENCES Adams, M. H. (1959). “Bacteriophages”. Interscience, New York. Anderson, T. F. (1945).J. Bact., 55, 651-665. hacker, R. L., and Ordal, E. J. (1956).J. Bact., 70,738-741. Anderson, D. L., and Bradley, S. G. (1964). J. gen. Microbiol., 37, 67-72. Betz, J. V., and Anderson, K. E. (1964). J. Bact., 87,408-41 5. Bishop, D. H. L., and Bradley, D. E. (1965). Biochem. J.,95, 82-93. Bradley, D. E. (1 964). 9.gen. Microbiol., 35,471-482. Bradley, D. E. (1966a). J. gen. Microbiol.,44, 383-397. Bradley, D. E. (1966b).J. gen. Microbiol., 45,83-96. Brinley-Morgan, W. J., Kay, D., and Bradley, D. E. (1960). Nature, Lond., 188, 74-75. Brinton, C. C. (1965). Trans. N . Y. Acad. Sci., 27, 1003-1054. Burchard, R. P., and Dworkin, M. (1966). J. Bact., 91, 1305-1 3 13. Clark, W. A. (1962). Appl. Mictobiol., 10, 466-471.

XII. BACTERIOPHAGES

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Crawford, E. M., and Gesteland, R. F. (1964). Virology, 22, 165-167. Crosse, J. E., and Garrett, C. M. E. (1963). J. appl. Buct., 26, 159-177. Davies, D. R., Arlett, C. F., Munsen, R. J., and Bridges, A. (1967).J.gen. Microbiol., 45,329-338. Dettori, R., Maccacaro, G. A., and Turri, M. (1963). G. Microbiol., 11, 15-34. de Klerk, H. C., Coetzee, J. N., and Foune, J. T. (1965).J. gen. Microbiol., 38,35-38. Duff, J. T., and Wyss, 0. (1961). J. gen. Microbiol., 24, 273-289. Eisenstark, A. (1966). in “Methods in Virology (Maramorosch, K. and Koprowski, H., Eds). Academic Press, New York and London. Feary, T. W., Fisher, E., and Fisher, l’ N.. (1964).J. Bact., 87, 196-208. Fry, B. A. (1963). J . gen. Microbiol., 31, 297-309. Groman, N. B., Eaton, M., and Rosher, Z. K. (1958).J. Buct., 75, 320-325. Guelin, A. (1955). Annls. Inst. Pusteur, Puris, 75, 485.496. Harm, W., and Rupert, C. S. (1963). Z.VererbLehre,94, 336-348. Hayes, W. (1964). “The Genetics of Bacteria and their Viruses”. Blackwell, Oxford. Hayward, A. C. (1964). J . gen. Microbiol., 25, 287-298. Hoffman-Berling, H., Kaetner, H. C., and Knippers, R. (1967). Adu. Virus Res., 12,329-370. Jacob, F., and Wollman, E. L. (1959). In “The Viruses” (Burnet, F. M. and Stanley, W. M., Eds). Academic Press, New York and London. Keogh, B. P., and Pettingill, G. (1966). Appl. Microbiol., 14, 421-424. Kingsbury, D. T., and Ordal, E. J. (1966).J. Buct., 91, 1327-1332. Kjems, E. (1960). Actu path. microbiol. scund., 49, 199-204. Klement, Z., and Lovas, B. (1959). Phytopathology,49,107-112. Kolstad, R. A., and Bradley, S. G. (1964).J. Bact., 87, 1157-1161. Lawn, A. M., Meynell, E., Meynell, G . G., and Datta, N. (1967). Nature, Lond., 216,343-346. Lewin, R. (1960). Nature, Lond., 186, 901. Mayor, €1. D., and Hill, N. 0. (1966). Virology, 14, 264-266. Meynell, E. (1961). J . gen. Microbiol., 25, 253-290. Meynell, G. G., and Meynell, E. (1965). “Theory and Practice in Experimental Bacteriology”. Cambridge University Press, London and New York. Okabe, N., and Goto, M. (1963). Ann. Rev. Phytoputh., 1, 397-418. Parenchych, W., and Graham, A. F. (1962). y, cell. comp. Physiol., 60, 199-208. Raettig, H. (1958). “Bacteriophagie”. Fischer, Stuttgart. Rauch, H. C., and Pickett, M. J. (1961). Cun.J. Microbiol., 7 , 125-133. Rifkind, D., and Pickett, M. J. (1953). J. Buct., 67, 243-246. Roslycky, E B., Allen, 0. N., and McCoy, E. (1962). Can.?. Microbiol., 8, 71-78. Schmidt, J. M., and Stanier, R. Y. (1965). J. gen. Microbiol., 39, 95-107. Schneider, J. R., Diener, T. O., and Safferman,R. S. (1964). Science, 144,1127-1130. Schwinghamer, E. A. (1965). Aust. J. bid. Sci., 18, 333-343. Sellers, M., Baxter, W. L., andRunnals, H. R. (1962). Can.J.Microbiol., 8,389-399. Spencer, R. (1960). J . Bact., 79, 614. Stolp, H., and Starr, M.P. (1964). Phytopath. Z.,51, 4424.78. Takeya, K., and Amako. K. (1966). Virology, 28, 163-165, Zinder, N. D. (1965). A. Rev.Microbiol., 19. 455-472. Zinder, N. D., Valentine, R. C., Roger, M.,.and Stoeckenius, W. (1963). Virology, 20,638-640.

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Author Index Numbers in italicsrefer to the pages on which references are listed at the end ofeach chapter.

A Arronson, S., 289, 290, 308, 310 Abou Akkada, A. R., 140, 141, 145, 146,148 Abram, D., 175, 182 Adams, M. E., 21, 39, 50 Adams, M. H., 316, 318, 321, 326, 327, 328,328 Adier, H. E., 216, 218, 219, 220, 231, 232, 261,262, 264,267 Afshar, A,, 218, 261 Akabori, S., 181,183 Aldrich, D. V., 292, 308 Alford, R. H., 227, 266 Algeus, S., 287, 308 Allen, E. J., 290, 308 Allen, M. B., 54, 57, 61, 62, 63, 64,75, 273, 277, 286, 287, 288, 289,293,308 Allen, 0. N., 316,329 Allgeier, D. L., 83, 113 Allison, F. E., 56, 63, 75, 287, 308 Altemeier, W. A., 91, 113 Amako, K., 322, 329 Amesz, J., 63, 73, 75, 76 Anacker, R. L., 316,328 Anagnostidis, K., 24, 49 Anderson, C. G., 97, 114 Anderson, D. L., 316, 328 Anderson, D. R., 257,258,261 Anderson, E. H., 289, 308 Anderson, K. E., 316, 328 Anderson, T. F., 325,328 Andrewes, C. H., 213, 261 Andrews, B. E., 223, 250, 261 Angelotti, R., 91, 112 Annotation, 216, 261 Antia, N. J., 291, 308 Argaman, M., 240, 265 Arisoy, F., 218, 219, 232, 261, 267 Arlett, C. F., 327,329 Armstrong, D., 219, 220, 261 Armstrong, F. A. J., 300, 308

Arnon, D. I., 61, 62, 64, 75, 286, 288, 293, 296, 308, 313 Arthur, W., 288, 310 Atkinson, D. E., 22, 49 Avigan, J., 240, 265

B Baalen, C. van, 62, 77 Baalsrud, K., 19, 49, 50 Baalsrud, K. S., 50 Baars, J. K., 21, 39, 50 Baas-Becking, L. G. M., 169, 170, 171, 182 Bach, M. K., 289, 293, 295, 297, 310 Baile, D. L., 289, 308 Baird-Parker, A. C., 152, 159 Baker, H., 287, 289, 290, 308, 310 Baker, R. L., 213, 266 Bakos, K., 219, 222, 234, 262 Balitskaya, R. M., 75, 76, 77 Barber, F. W., 289, 308 Barile, M. F., 217, 226, 227, 229, 237, 257,258,261 Barker, H. A., 290,308 Barnes, E. M., 143, 148, 149, 154, 158, 159,159,160 Barrett, F. C., 154, 259 Barry, N. C., 290,308 Barry, S., 290, 308 Batty, I., 94, 95, 96, 102, 112, 113 Baumgartner, J. G., 173, 182 Bavendamm, W., 6, 7, 8, 9, 18, 20, 50 Baxter,W. L., 316, 329 Beerens, H., 152,153,154,155, 157,195 160 Beger, H., 25, 50 Beijerinek, M. W., 54, 76, 276, 308 Belt, M., 289, 310 Bendix, S., 287, 308 Benn, E. C., 217, 264

332

AUTHOR INDEX

Bensky, B., 290, 311 Beran, K., 306, 311 Bergey, 4, 5 , 11, 12, 23, 26, 50, 152, 159 Berkman, J. I., 290, 310 Bernsteinas, J. P., 162, 168 Betts, A. O., 216, 261 Betz, J. V., 316, 328 Beveridge, W. 1. B., 152, 159, 216, 261 Bhalla, N. P., 218, 262 Biedermann, M., 74, 76 Biliett, F. S., 81, 113 Bishop, D. H. L., 322, 323, 328 Bissett, K. A., 18, 50 Bjerknes, Clara A., 39, 50 Blackburn, T. H., 140, 141, 145, 146, 148 Bladen, H. A., 145, 149 Blecker, H. H., 73, 76 Block, M. R., 169, 182 Boalch, G. T., 291, 292, 308 Bocks, S. M., 289, 313 Bodey, G. P., 229, 261 Boidin, A. G., 219, 232, 261 Bold, H. C., 286, 308 Boothroyd, M., 95, 113 Boring, J., 171, 182 Borrel, 212, 218, 261, 265 Bose, S. I<., 66, 67, 68, 69, 70, 72, 76 Bosher, 2. K., 316,329 Bovell, C., Jr., 22, 23, 42, 51 Bowman, R. O., 56, 77 Bradley, D. E., 316, 321, 322, 323, 328 Bradley, S. G., 316, 328, 329 Bradshaw, J. S., 282,313 Breed, R. S., 80, 108, 113, 185, 209 Brewer, J. H., 83, 89, 113 Bridges, A., 327, 329 Bridges, A. E., 88, 113 Bridges, J. B., 217, 264 BridrC, J., 212, 218, 219, 261 Brighton, W. D., 250, 261 Brinley-Morgan, W. J., 316, 328 Brinton, C. C., 323, 328 Bristol Roach, B. M., 287, 308 Broberg, G., 134, 149 Brock, M. L., 164, 167 Brock, T. D., 162, 164, 167 Brooks, M. E., 96, 113 Brown, H. J., 171, 175, 182 Brown, M. S., 217, 267

Brown, T. M., 243, 244, 261 Brown, T. McP., 251, 253, 261, 263 Brugger, J. E., 288, 310 Bruin, A. S. de, 87, 114 Bryant, M. P., 127, 132, 137, 139,144, 146,148,149 Buchner, H., 87, 113 Buckley, L. S., 218,219,231,262 Buetow, D. E., 289,308 Bullis, C., 223, 229, 265 Bullivant, S., 240, 262 Bunker, H. J., 19, 50 Bunt, J. S., 54, 55, 56, 76 Burchard, R. P., 316, 328 Burkholder, P. R., 293, 308 Burlew, J. S., 306, 308 Burris, R. H., 22, 23, 40, 50 Burton, S. D., 18, 50 Busby, W. F., 287, 311 Butler, H. M., 105, 113 Butlin, K. R., 21, 39, 50 Buttiaux, R., 181, 182

C Caldwell, D. R., 146, 149 Callely, A. G., 69, 76, 290, 308 Calvin, J., 287, 288, 290, 312 Calvin, M., 287, 311 Cameron, 0. L., 308 Campbell, A., 236, 264, 265 Campbell, A. D., 229, 230,267 Campbell, L. L., 22, 51, 162, 167 Campbell, L. L., Jr., 22, 50 Campbell, R. C . , 216, 261 Canchola, J., 217, 222, 265, 266 Cannon, D. A., 81, 114 Cardella, M. A., 90, 91, 115 Carlucci, A. F., 300, 309 Cai-michael, L. E., 218, 262, 264 Carr, N. G., 63, 65, 76, 77 Carski, T. R., 217, 237, 238, 262 Castel, M. M., 152, 153, 157, 159 Cataldi, M. S., 18, 32, 50 Cato, E. P., 155, 160 Chalquest, R. R., 220, 236, 261 Chandler, V. L., 88, 113 Chang, V., 237, 265 Chanock, R., 217, 266

AUTHOR INDEX

333

Chanock, R. M., 212, 216, 217, 222, Crowther, S., 227, 262 226,227,229,237, 246, 248, 250, 252, Cruickshank, R., 233, 262 Csonka, G. W., 217,262 261, 262, 265, 266, 267 Charles, A. B., 154, 158, 159, 159, 160 Cutchins, E., 90, 91, 113 Cho, K. Y., 173, 182 Cholodny, N., 25, 50 Chorney, W., 288, 308 D Chou, H., 146,149 Dack, G. M., 81, 113 Chowdray, J. E., 223, 237, 266 Dafaalla, E. N., 230, 262 Christian, J. H . B., 172, 182 Danielsson, D., 219, 222, 234, 262 Chu, H. P., 213, 220, 257, 261, 265 Chu, S. P., 60, 76, 286, 287, 293, 294, Darbyshire, J. H., 218, 262 Darnforth, W. F., 287, 309 300,308 Clark, H. W., 249, 253, 256, 257, 261 Datta, N., 323, 329 Davidson, I., 218, 266 Clark, L. B., 303, 306, 312 Davidson, M., 217, 262 Clark, W. A., 328, 328 Davies, D. R., 327, 329 Clarke, P. H., 97, 114 Clyde, W. A., Jr., 217, 222, 257, 258, Davies, J., 223, 261 Davis, H. S., 196, 209 261,262,264 Davis, J. S., 289, 309 Cobb, H. D., 62, 76 Davis, R. E., 143, 149 Coble, D. W., 251, 263 Dawson, P. S., 218,262 Coetzee, E. F. C., 106, 115 Dawson,P. S. S., 148, 149 . Coetzee, J. N., 316, 329 Debonera, G., 218, 262 Cohen, J. S., 22, 23, 40, 50 De Cicco, B. T., 23, 50 Cohen-Bazire, G., 74, 76 Deeb, B. J., 220, 262 Cole, B. C., 220, 235, 262 Del Giudice, R. A., 217, 237, 238, 262 Cole, R., 240, 262 Dellinger, J. D., 218, 264 Coleman, G. S., 22, 50 Denny, F. W., 257, 264 Collier, A., 291, 313 Dettori, R., 318, 323, 329 Collins, V. G., 1, 3, 17, 33, 34, 37, 50 Dhanda, M. R., 218, 262 Colmer, A. A., 19, 50 Dick, A. T., 229, 230, 267 Colmer, A. R., 24, 52 Dick, H. M., 217, 264 Committee on Cultures, 286, 309 Committee upon Anaerobic Bacteria Diener, T. O., 316, 329 Dienes, L., 240, 241, 262 and Infections, 81, 113 Dierks, R. E., 220, 235, 262 Connole, M. D., 218, 262 Dinter, Z., 216, 219, 222, 234, 262, 266 Conti, S. F., 74, 76 Disraely, M . N., 90, 114, 158, 160 Coons, A. H., 94, 113 Dodson, A. N., 274, 309 Cordy, D. R., 218, 219, 232, 261, 262 Corse, J., 217, 262 Doetsch, R. N., 145, 149, 181, 183 Cosenza, B. J., 213, 255, 265 Doman, N. G., 68, 76 Cottew, G. S., 216, 218, 219, 231, 232, Donati, E. J., 221, 264 261, 262, 264, 267 Donatien, A., 212, 218, 219, 261 Craig, F. N., 287, 309 Dorff, P., 13, 14, 15, 25, 50 Cramer, M. L., 289, 309 Dowell, V. R., 91, 113 Crawford, E. M., 323, 329 Downing, T. O., 221, 264 Crawford, Y. E., 227, 238, 262, 264 Doy, C. H., 173,182 Creech, H. J., 94, 113 Drebes, G., 291, 313 Crespi, H. L., 288, 308 Drews, G., 74, 76 Crosse, J. E., 316, 329 Driscoll, S. G., 228, 264

334

AUTHOR INDEX

Droop, M. R., 61, 62, 64, 77, 273, 274, 276,278,282,284,286,290,291, 292, 293, 294, 295,297, 298,301, 303, 306, 307,309,312 Duff, J. T., 316, 329 Dugan, P. R., 24, 50 Dujardin-Beaumetz, 212, 218, 265 Dulbecco, R., 233, 262 Duncan, D. W., 19, 50 Dundas, I. D., 171, 175, 176, 182 Dussault, H. P., 174, 178, 179, 180, 182, 183 Dworkin, M., 200, 209, 316, 328 Dyer, D. L., 63, 73, 76

E Eadie, J. M., 137, 139, 140, 149 Eaton, M., 316, 329 Eaton, M. D., 217, 246, 262, 264 Edgson, F. A., 218,266 Editorial Board, 212, 262 Edward, D. G. ff., 212, 213, 217, 219, 220, 227, 235, 252,261, 262, 265 Ehrlich, H. L., 25, 52 Eimhjellen, K., 175, 180, 183 Eisenstark, A., 315, 316, 319, 320, 322, 326,327,329 Ellis, D., 18, 50 Elsworth, E., 306,310 Emard, L. O., 90, 113 Emerson, R., 287, 294, 309 Engwall, C., 91, 114 Eppley, R. W., 289, 290, 309, 311 Erdag, O., 218, 219, 231, 232, 261,262, 267 Erne, H., 231,238,262 Ertel, I. J., 223, 229, 265 Etchells, J. L., 173, 180, 183 Etheridge, J. R., 218, 224, 230, 264 Exell, G., 65, 76 Eymers, J. G., 68, 70, 76

F Fabricant, J., 212, 220, 236, 252, 261, 262,263 Fallon, R. J., 213, 222, 263

Farber, J. F., 221, 264 Farlow, W. G., 170, 183 Farrell, J., 161, 162, 167 Faust, L., 18, 32, 33, 50 Fay, P., 63, 76 Feary, T. W., 319, 322,329 Fedorov, V. D., 57, 69, 77 Ferguson, L. C., 219, 232, 263 Field, L. E., 218, 261 Fievez, L., 152, 159, 159 Fildes, P., 81, 82, 85, 113, 114 Filfus, J., 289, 293, 310 Fincher, M. G., 218, 261 Findlay, G. M., 220, 263 Finegold, S. M., 159, 160 Fish, F. F., 196, 209 Fisher, E., 319, 322, 329 Fisher, T. N., 319, 322, 329 Fitzgerald, G. P., 56, 60, 61, 72, 76, 287,309 Fitzgerald, W. A., 219, 227, 235, 262 Flannery, W. L., 181,183 Fogg, G. E., 56, 57, 76, 287, 288, 291, 292, 304, 306, 309, 310, 311 Foggie, A., 232, 264 Fogh, H., 237, 263 Fogh, J., 237, 263 Ford, D. K., 217, 263 Ford, J. E., 287, 309 Forsberg, C., 289, 309 Fortner, J., 87, 113 Foter, M. J., 91, 112 Fott, B., 273, 287, 309 Foune, J. T., 316, 329 Fowler, R. C., 251, 253, 261, 263 Fox, H., 217, 266 Fox, H. H., 216, 261 Foyn, B., 290, 300, 309 Frank, H. A., 22, 50 Frank, O., 290, 308, 310 Franklin, A. L., 289, 310 Fred, E. B., 88, 114 Freeze, H., 164, 167 French, C. S., 282, 309 Freundt, E. A., 212, 217,220, 252, 253, 256,262,263 Friedewald, W. T., 227, 266 Friedman, M., 219, 220, 261 Fritsch, F. E., 270, 309 Fry, B. A., 327,329

335

AUTHOR INDEX

Fujita, Y., 64, 73, 75, 76, 288, 310 Fujiwara, A., 288, 308 Fukumi, H., 182,183 Fuller, R. C., 69, 74, 76 Furness, G., 213, 244, 263

G Gafford, R. D., 63, 73, 76 Galloway, R. A., 289, 309 Galmiche, J. M., 289, 311 Gannon, J., 25, 51 Garrett, C. M. E., 316, 329 Gee, R., 289, 309 Geitler, L., 196, 209 Georgala, D. L., 95, 113 Gering, R. K., 288, 312 Gerloff, G. C., 56, 60, 61, 72, 76, 287, 309 Gest, H., 57,65,66, 67, 76, 77 Gesteland, R. F., 323, 329 Gibbons, N. E., 171, 172, 175, 176, 177, 181,182,183 Gibbs, M., 289, 311 Gilbert, H. D., 193,210 Gill, V., 237, 265 Goldberg, E. D., 290, 309 Goldberg,H. S., 154,158,159,159,160 Goldstein Brouwers, G. W. M., 87, 114 Golightly, L., 220, 235, 262 Goodwin, R. F. W., 216, 219, 232, 233, 234, 238, 239, 246, 254,263 Gordon, A. M., 217, 264 Goret, P., 181, 183 Gorham, P. R., 63, 73, 76, 77,288, 293 310,311,313 Goto, M., 316, 329 Gould, J., 56, 77 GourIay, R. N., 230, 263 Gowland, G., 98, 115 Grabowski, M. W., 229, 261 Grace, J. B., 18, 50 Graf, W., 196, 210 Graham, A. F., 322, 329 Granick, S., 288, 293, 313 Grayston, J. T., 245, 250, 264 Green, J. H., 162, 168 Greenblatt, C. L., 289, 309 Grieg, A. S., 219, 220, 232, 263 Griffiths, B. M., 18, 52

Griffiths, D. J., 288, 309 Grinsted, E., 89, 91, 113, 153, 160 Groman, N. B., 316, 329 Grossman, J. P., 22, 50 Guelin, A., 316, 329 Guillard, R,R. L., 291, 300, 309 Guillaume, J., 152, 153, 154, 155, 159, 160 Gusev, M. V., 73, 76 Gutekunst, R. R., 216, 261 Guthrie, R. S., 218, 261 Gutierrez, J,, 143, 149 Guyon, D., 289,312

H Haas, H., 251, 263 Hackett, D. P., 64, 77 Haig, D. A., 218, 231, 264, 267 Hale, H. H., 218, 263 Hall, E. R., 22, 50 Hall, F. J., 257, 258, 267 Hall, H. E., 91, 112 Hall, I. C., 80, 81, 90, 113 Hall, R. P., 289, 309 Hallaway, M., 63, 76 Halldal, P., 282, 309 Halliwell, G., 139, 145, 149 Halvorson, H. O., 175, 176, 178, 182, 183 Hamdy, A. H., 219, 232, 263 Hamilton, R. D., 300, 309 Hanert, H., 24, 50 Hanko, E., 218,263 Hansen, P. A., 181, 183 Happold, F. C., 19, 50 Harbourne, J. F., 218, 225, 263 Hardman, Y., 25, 50 Hare, R., 51, 113 Harm, W., 316, 329 Harris, R. F., 16, 51 Harrison, F. C., 170, 183 Hart, R. K., 218, 262 Hartman, H. A., 218, 263 Hase, E., 288, 294, 313 Haskins, C. P., 287, 289, 292, 310 Hattori, A., 64, 73, 75, 76, 288, 310 Havinga, A., 286, 290,311 Haxo, F. T., 290,309 Hayes, W., 323, 329

336

AUTHOR INDEX

Hayflick, L., 212, 216, 217, 226, 227, Hudson, J. R., 216, 218, 219, 224, 230, 231,264 229, 237, 239, 241, 243, 251, 252, Hughes, E. O., 288, 293, 310, 313 259,261, 262,263 Hughes, J. E., 22, 51 Hayward, A. C., 316, 329 Huhtanen, C. N., 289, 308 Hayward, N. J., 81, 89, 97, 113 Huijsmans-Evers, A. G. M., 217, 222, Heer, A. A., 83, 113 264 Heidenreich, P.,97, 114 Hulbert, E. M., 291,310 Heller, C. L., 82, 113 Hungate, R. E., 118, 119,125, 126, 127, Helmboldt, C. F., 218, 263 132,134,135,137,139,142,144,148, Hendley, D. D., 68, 70, 75, 76 149 Henrici, A. T., 25, 50 Hunter, D., 218, 225, 263 Henry, H., 81, 113 Hurlbert, R. E., 290, 310 Henry, M., 181, 183 Hutchins, J. 0.)289, 310 Herbert, D., 305, 306, 309, 310 Hutchinson, D., 213, 263 Herderschee, D., 217, 233, 263 Hutchinson, M., 19, 50 Herick, W. van, 217, 262 Hutner, S. H., 21,37, 38, 39, 50, 58,65, Hershenov, B., 290, 311 66, 67, 76, 77, 287, 289, 290, 292, Hewitt, E. J., 286, 310 293, 295, 297, 303, 308, 310, 312,313 Heyningen, W. E. van, 81, 113 Hill, E. 0.) 91, 113 Hill, N. O., 322, 329 1 Hirsch, A,, 89, 91, 113, 153, 160 Ichinose, H., 155, 160 Hoagland, D. R., 293, 310 Ichioka, P. S., 288, 308 Hoare, D. S., 16, 51, 63, 76 Imahori, K., 289, 310 Hoare, M., 218, 264 Imshenetsky, A. A., 194, 204, 210 Hobbs, B. C., 92, 113 Inglis, J. M., 223, 261 Hobbs, G., 91, 97, 99, 111, 115 Hobson, P. N., 137,139, 140, 141, 143, Ingraham, J. L., 162, 168 Ingram, G. C., 171, 183 144,145,146,147,148,148, 149 Ingram, M., 170, 171, 183 Hodgkins, W., 181, 183 Innes, J. R. M., 221, 264 Hoffman-Berling, H., 322, 329 Isenberg, H. D., 290, 310 Hoffman, C. E., 289, 310 Iwamura, T., 288, 294, 313 Hofstad, M. S., 220, 267 Iwanami, S., 182, 183 Holdeman, L. V., 155,160 Iwasa, K., 289, 310 Holmes, P. K., 178, 183 Iwasaki, H., 291, 310 Holm-Hansen, 0.) 60, 64, 76 Holt, S. C., 74, 76 Holton, R. W., 54, 63, 73, 76 J Hoogenhout, H., 63, 73, 75, 76, 307, Jackson, D. K., 222, 263 310 Jackson, J. F., 19, 50 Hoover, S. R., 63, 75, 287, 308 Jacob, F., 326,329 Hopkins, E. F., 289, 310 Jahn, T. L., 292, 312 Home, R. W., 257, 258, 261, 267 Jain, N. C., 218, 264 Hornsey, H. C., 171, 183 James, D. M., 286, 310 Hospodka, J., 306, 311 Hottle, G. A., 213, 264 James, T. W., 290, 312 Howard, B. H., 134, 149 James, W. D., 216, 246, 248, 250, 261, Howell, D., 218, 266 266 Heye, KR., 170, 183 Jasper, D. E., 218, 264 Huck, R. A., 218, 261 Jeantet, 212, 261

337

AUTHOR INDEX

Jeffers, E. E., 197, 210 Jensen, K. E., 222, 264, 267 Jitts, H. R., 280, 282, 291, 310 Johansson, K. R., 90, 113 Johnson, K., 217, 266 Johnson, S. D., 218, 261 Johnston, L. A. Y., 218, 266 Johnstone, K. I., 19, 50 Jones, D. M., 217, 264 Jones, G. L., 19, 52 Jones, I. D., 173, 180, 183 Jones, R. F., 292, 305, 310, 311 Jones, R. N., 94, 113 Jouan, 212,261 Joubert, L., 181,183 Jukes, T. H., 289,310

K Kaetner, H. C., 322, 329 Kagan, G. Y., 240, 264 Kain, J. M., 291, 292, 310 Kalmakoff, J., 291, 308 Kanarek, A. D., 220, 262 Karavaiko, 19, 51 Karnauchow, P. N., 87, 114 Katz, J. J., 288, 308 Katznelson, H., 174, 175, 183 Kaufman, L., 108, 113 Kawatomari, T., 108, 114 Kay, D., 316, 328 Keane, M., 65, 66, 67, 76 Keil, F., 18, 50 Kellerman, K. F., 170, 183 Kelton, W. H., 245, 246, 259, 260, 264 Kennedy, M. E., 170, 183 Kenny, G. E., 220, 245, 250,262, 264 Keogh, B. P., 328, 329 Kerr, K. M., 236, 264, 265 Kessler, E., 288, 310 Ketchum, B. H., 290, 310 Kim, K. S., 257, 258, 262, 264 Kingsbury, D. T., 316, 329 Kinne-Diettrich, E.-M., 291, 310 Kistner, A., 23, 42, 50, 148, 149 Kitchell, A. G., 171, 183 Kiyohara, T., 64, 73, 76, 288, 289,310, 313 Kjems, E., 316, 329 Klebahn, H., 170, 183

Kleckner, A. L., 220, 264 Klein, M. R., 290; 310 Klement, Z., 316, 329 Klerk, H. C. de, 316, 329 Kleineberger, E., 220, 234, 263, 264 Kleineberger-Nobel, E., 212, 216, 218, 234,253,264 Knight, B. C. J. C., 227, 262 Knight, B. C. J. G., 227, 265 Knight, V., 217, 266 Knippers, R., 322, 329 Knittingen, J., 182, 183 Knyszynski, A., 217, 265 Kocur, M., 181, 183 Koffler, H., 22, 52 Kohn, J., 145, 149 Kolstad, R. A., 316, 329 Kondrat'eva, E. N., 57, 75, 76 Kondratieva, E. N., 57, 69, 77 Kono, M., 181,183 Koprowski, H., 259, 263 Kornberg, H. L., 65, 76 Kramer, N. C., 251,263 Kratz, W. A., 61, 62, 64, 73, 75, 76, 77, 288,310 Krauss, M. P., 56, 76 Krauss, R. W., 288, 289, 309, 213 Kravetz, H., 217, 266 Kraybill, W. H., 227, 238, 262, 264 Kreig, N. R., 24, 48, 51 Krzemieniewska, H., 192, 194, 210 Krzemieniewski, S., 192, 194, 210 Kucera, S., 48, 50 Kuffareth, H., 286, 310 Kumar, H. D., 289, 310 Kundsin, R. B., 228, 264 Kunieda, R., 307, 313 Kunisawa, R., 74, 76 Kurihara, Y.,137, 139, 140, 149 Kushner, D. J., 171, 173, 178, 182, 183

L Lachance, R. A., 174, 183 Lackey, J. B., 18, 25, 50, 52 Laidlaw, P. P., 81, 113 Langer, P. H., 218,264 Larsen, H., 17, 20, 21, 37, 50, 57. 59, 69, 72, 77, 170, 171, 173, 182, 183

338

AUTHOR INDEX

Lascelles, J., 60, 65, 66, 67, 74, 76, 77 Lawn, A. M., 323,329 Laws, L., 218, 219,262, 264 Lazoroff, N., 19, 50, 56, 77, 288, 310 Leach, R. H., 218, 225, 263,264 Lees, H., 16, 19, 51 Lemcke, R. M., 212, 217, 220, 221, 229,252,262,264 Lewin, J. C., 273, 287, 288, 291, 292, 293,310,311 Lewin, R., 316, 329 Lewin, R. A., 64, 76, 77, 270, 273, 278, 287,288,289,290,291,292,293,311 Lewis, C. M., 287, 294,309 Lewis, K. H., 91, 112 Lewis, W. M., 173, 180, 183 Lieske, R., 18, 33, 51 Lifshitz, Y., 217, 265 Lilly, H. A., 91, 113 Lindberg, R. B., 81, 90, 91, 113, 114 Lindhal, I. L., 143, 149 Links, J., 286, 290, 311 Linquist, W. E., 218, 261 Liston, J., 181, 183 Litsky, W., 25, 51 Little, P. A., 289, 311 Livingston, C. W., Jr. 234, 265 Lloyd, D., 290, 308 Lloyd, L. G., 218, 231, 262 Lloyd-Jones, C. P., 286,310 Lockhart, W. R., 87, 113 Lochhead, A. G., 174, 175, 183 Lockwood, S., 289, 310 London, J., 19, 51 Longley, E. O., 216, 218, 231, 264 Lorenzen, H., 307,311, 312 Lovas, B., 316, 329 Low, I. E., 246, 264 Lowbury, E. J. L., 91, 113 Lubart, K. J., 65, 66, 67, 76 Lucas, D. R., 96, 114 Ludiarn, G. B., 217, 264 Lunceford, C. D., 213, 228, 229, 266 Lundgren, D. G., 24, 49, 50,51 Lutsky, I. I., 220, 264 Lwoff, A., 274, 311 Lyalikova, N. N., 19, 51 Lyell, A., 217, 264 Lynch, V. H., 287, 311 Lyons, M. J.. 213,265

M McAllister, C. D., 280, 282, 291, 310 McArthur, J. M., 134, 149 Maccacaro, G. A., 318, 323, 329 MacCallum, F. O., 220, 263 McCance, M. E., 176, 183 McCarthy, A. J., 273, 287, 309 McCartney, J. E., 147, 149 McClung, L. S., 80,81,88,97,113,114 McCoy, E., 80, 88, 114, 316, 329 McCrea, C. T., 218, 262 McCurdy, H. D., 199, 202, 207, 210 McDaniel, H. R., 56, 77 McDonald, J. C., 199, 210 McDougall, E. I., 134, 149 McDougalI, I. E., 146, 149 McFaden, B. A., 22,49 Macfarlane, R. G., 97, 114 McGee, 2. A., 241, 264 Macias, R. F. M., 289, 290, 309, 311 McIntosh, J., 81, 82, 85, 113, 114 Mackay, J. M. K., 219, 232, 264 Mackenzie, R. D., 220, 263 Mackereth, F. J. H., 3, 51 Mackie, T. J., 147,149 McLachan, J., 63, 73, 77, 288, 290, 292, 311 McLaughlin, C. B., 83, 113 McLaughlin, J. J. A., 61, 62, 64, 77, 284, 286, 291, 292, 293, 294, 297, 298, 301, 310, 311, 312 MacLennan, J. D., 81, 114 MacLeod, A. K., 230, 232, 263, 265 McLeod, J. W., 87, 114 MacLeod, R. A., 169. 183 MacMurtrey, M. J., 213, 244, 263 McVay, L. V., 81, 114 Maddux, W. S., 305, 311 Malek, I., 306, 311 Mallows, J. H., 171, 183 Man, J. C. de, 143, 149 Manchee, R. J., 246, 267 Maniloff, J., 213, 257, 264, 265 Mann, S. O., 137, 139, 140, 141, 143, 144, 145, 147, 149 Manten, A., 59, 77 Marchisotto, J., 293, 311 Mare, C. J., 219, 233, 264 Markham, F. S., 220, 264

339

AUTHOR INDEX

Markham, R., 156, 160 Marsh, H . V., 289, 311 Marshall, J . D., 90, 91, 108, 114, 115 Marshall, J . H., 87, 114 Marshall, S. M., 291, 311 Martin, G. W., 193, 210 Martinec, T., 181, 183 Maruo, B., 181, 183 Marx, R., 74, 76 Mascoli, C. C., 236, 264 Mason, R. P., 90, 91, 113 Matovitch, V . B., 290, 308 Matthews, A. D., 87, 114 Matudaira, T., 290, 311 Mayor, H . D., 322, 329 Medical Research Council, 108, 114 Medrek, T . F., 143, 149 Meffert, M. E., 288, 311 Meiklejohn, G., 217, 262 Meiklejohn, J., 16, 51 Mercer, E. H., 173, 182 Merrett, M. S., 289, 313 Meynell, E., 316, 320, 323, 326, 327,

329 Meynell, G. G., 316, 320, 323, 326, 327,

Moriber, L. G., 290, 311 Morimura, Y., 307, 311, 313 Morita, Y., 18, 50 Morowitz, H . J., 213, 257, 264, 265 Morris, H . J., 56, 63, 75 Morrison, H . J., 287, 308 Morselli, M.-F., 290, 310 Mortimer, C . H., 3, 51 Morton, H. E., 227, 265 Morton, V., 219, 220, 261 Moskal, P. A., 91, 114 Mossel, D. A. A., 87, 114 Moyse, A., 289, 312 Muellar, W . S., 25, 51 Mufson, M . A., 237, 265 Mulder, E. G., 24, 25, 47, 51 Miiller, I., 288, 313 Munsen, R. J., 327, 329 Murphy, W . H., 223, 229,265 Murray, E. G. D., 80, 108, 113, 185,

209 Myers, J., 61,62, 64,73,75, 76, 77, 273,

288,289,303,304,306,309,310,312, 313 Mylroie, R. L., 127, 132

329 Middlebrook, J . B., 56, 77 Miles, A. A., 89, 97, 103, 113, 114, 248,

205 Miles, E. M., 97,114 Miller, A. B., 159, 160 Miller, J . D. A., 22, 51, 288, 304, 306,

309,311 Miller, L. P., 22,40, 51 Miltimore, J . E., 134, 149 Ming, P.-M. L., 228, 264 Minken, J., 217, 263 Miquel, P., 290, 311 Mishustin, E. N., 194, 210 Misra, R. A., 248, 265 Mituya, A., 288, 294, 313 Miyachi, S., 288, 311 Moewus, F., 287, 311 Moewus, L., 287, 311 Monod, J., 304, 311 Moore, M. L., 91, 114 Moore, R. B., 63, 76 Moore, R. W., 234, 265 Moore, W . E. C., 1 5 5 , 160 Moriarty, D. J . W., 19, 50

N Nagler, F. P. O., 96, 97, 114 Nasri, M. El, 218, 231, 246, 265 Nathan, H., 289, 310 Naylor, P. G. D., 87, 114 Neeb, O., 288, 312 Neimark, H . C., 241, 265 Neish, A. C., 286, 288, 312 Nelson, E. W., 290, 308 Nelson, J . B., 213, 220, 265 Newing, C. R., 230,232, 265 Newman, J . A., 220, 235, 262 Newnham, A. G., 213, 265 Nicholas, D. J . D., 19, 50 Nickell, L. G., 293, 308 Nicol, C. S., 217, 265 Niel, C. B. van, 20, 21, 37, 51, 57, 58,

59,77 Nielsen S. W., 218, 263 Nihei, T., 288, 294, 313 Nisbert, D. I., 219, 232, 264 Nocard, 212, 218, 265 Nordli, E., 291, 312

340

AUTHOR INDEX

NorBn, B., 199, 200, 210 Norris, L., 287, 288, 290, 312 Norris, R. E., 287, 288, 290, 312 Novick, A., 304, 312 Nowak, A., 293, 311 Nowak, J., 212, 265 Nunley, J. W., 24, 48, 51 0

Oakley, C. L., 97, 114 Okabe, N., 316, 329 Oleson, J. J., 289, 311 Olson, N. O., 236, 264, 265 Omata, R. R., 90, 214, 158, 160 Omata, S., 180, 181, 183 Onishi, H., 176, 177, 183 Onofrey, E., 169, 183 Onore, M., 73, 76 Ordal, E. J., 196, 210, 316, 325, 329 Organick, A. B., 220, 264 Ormerod, J. G., 65, 66, 67, 77 Omerod, K. S., 65, 66, 67, 77 Orr, A. P., 291, 311 Orr, J. H., 81, 98, 112, 114 Orskov, J., 253, 265 Ostelind, S., 287, 312 Osteux, R., 155, 160 Otterlin, S. E., 218, 263 Overbeck, J., 24, 49, 51 Oxford, A. E., 144, 149

P Paasche, E., 292, 312 Packer, L., 278, 312 Padilla, G . M., 289, 290, 308, 312 Pankevicius, J. A., 221, 264 Pankhurst, E. S., 22, 51 Parenchych, W., 322,329 Parish, H. J., 81, 114 Parker, E. T., 162, 168 Parker, W. H., 218, 262 Pasher, I., 290, 308 Payne, J. I., 172, 183 Pearce, J., 63, 76, 77 Pene, J . J., 241, 265 Pepper, R. E., 88, 113 Peterson, J. E., 199, 210

Peterson, R. A., 289, 310 Petit, H., 154, 155, 259, 160 Pettingill, G., 328, 329 Pfennig, N., 20, 21, 34, 51, 57, 58, 59, 68, 70, 74, 76, 77 Phaup, J . D., 25, 51 Phillips, J. N., 304, 312 Philpott, D. E., 291, 311 Pickett, M. J., 316, 329 Piercy, S. E., 216, 265 Pillai, C., 234, 265 Pintner, I. S., 278, 286. 287, 288, 290, 292, 293, 295, 297, 312 Pipes, F. J., 213, 244, 263 Pirson, A,, 307, 312 Pittman, K, A., 146, 149 Plastridge, W. N., 218, 231, 262, 263 Pochmann, A., 18, 51 Pomeroy, A. P., 219,232, 233, 234, 238, 239,263

Pomeroy, B. S., 220, 235, 262 Posnick, D. J., 159, 260 Postgate, J. R., 22, 50, 51 Pounden, W . D., 219, 232,263 Powelson, D., 22, 52 Prager, J., 293, 311 Prasad, R., 64, 76 Pratt, R., 287, 312 Price, C. A., 290, 312 Pringsheim, E. G., 25, 51, 53, 54, 55, 77, 273, 274, 280, 284, 286, 287, 288, 289, 290, 291, 298, 299, 312 Pringsheim, O., 291, 312 Proctor, V. W., 287, 288, 312 Provasoli, L., 61, 62, 64, 77, 278, 284, 286, 287, 288, 289, 290, 291, 292,293, 294,295,297,298, 301, 303,310,312 Purcell, R. H., 217, 222, 229, 246, 265, 267 Purdom, M . R., 143, 145, 149

R Raettig, H., 315, 329 Rahat, M., 291, 292, 312 Rajanikant, R. P., 251, 263 Rasmussen, G., 217, 263 Rauch, H. C., 316, 329 Razell, W. E., 19, 51

341

AUTHOR INDEX

Razin, S., 212, 213, 217, 219, 220, 227, 234,240,241,251,252,255,262,263, 265,266,267 Redfield, A., 290, 310 Redmond, H. E., 234,265 Reed, G. B., 81, 98, 112, 114 Reich, K., 291, 312 Reimann, B. E., 287,311 Reisner, G. S., 288, 312 Renault, L., 181, 183 Rettger, L. F., 80, 81, 90, 114, 115 Reynolds, N., 287, 312 Rhijn, G. R. van, 217,263 Rhoades, H. E., 234,265 Rhodes, M. E., 51 Ried, A., 288, 313 Rifkind, D., 217, 266, 316, 329 Riggs, D. B., 229, 261 Rittenberg, S. C., 290, 310 Roberts, D. H., 220,266 Roberts, R. J., 227,265 Robertson, M., 81, 89, 114 Robillard, N. F., 237, 238, 262 Robinson, I. M., 137, 146, 149 Robinson, J., 181, 183 Rockwell, G. E., 87, 114 Rodhe, W., 287, 294,313 Rodriguez, E., 289, 310 Roger, M., 322, 329 Rogers, H. J., 19, 50 Rogosa, M., 143, 144, 149 Rogul, M., 241, 264 Romanova, A. K., 68, 76 Romansky, M. J., 237, 265 Rose, A. H., 161, 162, 163, 167, 168 Rosenthal, L., 87, 114 Roslycky, E. B., 316, 329 Ross, G. 1. M., 289, 293, 295, 297, 310 ROSS,M. A., 221, 264 Ross, R. F., 234, 266 Rothblat, G. H., 240, 266 Rottem, S., 241, 251, 266 Roux, 212, 218, 265 Rowen, R., 173, 183 Rucker, R. R., 196, 209, 210 Ruiter, M., 217, 266 Runnals, H. R., 316, 329 Rupert, C. S., 316, 329 Ruys, A. C., 217, 222, 263, 264

Ryschenkow, E. I., 243, 266, Ryther, J. H., 291, 292, 300,309,313

S Sacks, T. G., 253, 263 SafTerman, R. S., 316, 329 Sager, R., 288, 293, 313 Sakazaki, R., 182, 183 Salimbeni, 212, 218, 265 Saltman, P., 289, 309 Sanders, M., 289, 310 Sandholzer, L. A., 95, 115 San Pietro, A., 57, 76 Santer, M., 19, 52 Sasa, S., 288, 294, 313 Saunders, G. F., 22, 51 Schaffner, Y., 152, 153,159 Schatz, A., 22, 23,42,51, 287, 289, 292, 310,312 Scher, B., 58, 77 Scher, S., 58, 77, 289, 308 ScherRel, A., 212,266 Schiff, J. A,, 289, 309 Schimke, R. T., 237,261 Schlegel, H. G., 59, 68, 77 Schmidt, J. M., 316, 322, 329 Schmidt, R. R., 288, 313 Schneider, J. R., 316, 329 Schreiber, E., 290, 313 Schroder, J., 74, 76 Schwabacher, H., 96, 114 Schwinghamer, E. A., 316, 329 Scoog, F., 287, 309 Scotten, H. L., 18, 51, 64, 77, 282, 288, 313 Scully, N. J., 288, 308 Sehgal, S. N., 172, 175, 183 Seiffert, G., 96, 114 Sellers, M., 316, 329 Senterfit, L. B., 216, 222, 261, 267 Serrano, B. A., 251, 263 Shaposhnikov, V. N., 57, 69, 75, 77 Sharma, G. L., 218, 262 Sharpe, M. E., 143,149 Sheagren, J. N., 217, 267 Shepard, M. C., 213,217,224,228,229, 246,259,260,266 Shewan, J. M., 181, 183 Shibata, K., 288, 294, 313

342

AUTHOR INDEX

Shiio, I., 181, 183 Shoetensack, H. M., 220, 234, 266 Shoetensack, M., 220, 234, 266 Silverman, M. P., 24, 49, 51 Simmons, G. C., 218, 266 Simon, J., 234, 265 Singh, B. N., 195, 210 Sistrom, W. R., 74, 75, 76, 77 Skerman, V. B. D., 13, 16, 19, 22, 23, 24, 26, 27, 28, 29, 30, 31, 34, 39, 42, 43, 44, 45, 51, 59, 77 Skinner, F. A., 31, 51 Skoog, F., 56, 60, 61, 72, 76 Skuja, H., 6, 7, 8,9,10, 14,15, 20, 51 Slanetz, L. W., 81, 90, 114 Slavin, G., 218, 266 Slyter, L. L., 148, 149 Smith, C. B., 227,266 Smith, F. B., 181, 183 Smith, H. J., 16, 51 Smith, H. W., 91, 114 Smith, L. D. S., 81, 102, 108, 114 Smith, N. R., 80, 108, 113, 185, 209 Smith, P. F., 213, 240, 247, 248, 251, 266 Smith, P. H., 118, 132, 142, 149 Smith, W., 148, 149 Smith, W. E. E., 304, 306, 309 Snyder, J., 229, 261 Snyder, W. C., 293, 310 Sobeslavsky, 0.)222, 267 Sobotka, H., 287, 290, 308 Soeder, C. J., 288, 313 Sokolova, G. A., 19, 51 Soli, G., 292, 313 Solntzeva, L., 194, 210 Somerson, N. L., 212, 246, 248, 250, 252, 262, 266, 267 Somerville, R. G., 217, 264 Sommers, L. E., 16, 51 Soriano, S., 16, 31, 51, 64, 77 Sorokin, C., 273, 288, 313 Southworth, J. M. L., 91, 115 Spencer, C. P., 283, 290,313 Spencer, R., 316, 329 Spray, R. S., 81, 87, 89, 90, 108, 114, 115 Sprunt, D. H., 81, 114 Srinivasan, V. R., 175, 176, 182 Stanbridge, E., 237, 239, 263

Stanier, R. Y., 74, 76, 194, 210, 316, 322,329 Stanley, S. O., 163, 168 Starkey, R. L., 18, 21, 33, 34, 39, 51,52 Starr, M. P., 74, 77, 321, 329 Steabben, D. B., 220, 264 Stephens, K., 280, 282, 291, 310 Sterne, M., 96,113 Stewart, S. M., 223, 237, 254, 266 Stewart, W. D. P., 292, 313 Stoeckenius, W., 173, 183, 322, 329 Stokes, E. J., 85, 115, 217, 266 Stokes, J. L., 64, 77, 162, 168, 288,313 Stokstad, E. L. R., 289, 310 Stolp, H., 321, 329 Storm, J., 287, 313 Stouffer, R. M., 221, 264 Stout, H. A., 22, 52 Stout, P. R., 313 Street, H. E., 288, 309 Strickland, J. D. H., 280, 282, 291, 300, 308,310 Strong, D. H., 91, 115 Stross, R. G., 289, 313 Stuart, P., 218, 261, 262, 266 Stuart, R. D., 223, 266 Stukus, P. E., 23, 50 Stula, E. F., 218, 263 Summers, R., 146, 148, 149 Sundheim, L. H., 290, 310 Suzuki, S., 155, 160 Sweeney, B. M., 290, 309, 313 Swift, H. F., 243, 244, 261 Switzer, W. P., 219, 220, 233, 234, 264, 266 Syrett, P. J., 288, 289, 313 Sziiard, L., 304, 312

T Taha, M. S., 56, 61, 62, 77 Tahon-Castel, M. M., 155, 159 Takano, H., 292, 313 Takeya, K., 322, 329 Tamiya, H., 288, 294, 307, 311, 313 Taniguchi, S., 181,183 Taylor, C. E. D., 106, 114 Taylor, J. H., 216, 261 Taylor, R. W., 292, 313

343

AUTHOR INDEX

Taylor-Robinson, D., 213, 216, 217, 218, 222, 229, 231, 234, 246, 250, 265,266,267 Tchan, Y. T., 56, 77 Telling, R. C . , 306, 310 Temple, K. L., 24, 52 Terent'eva, 2. A., 68, 76 Thaxter, R., 196, 210 Thjotta, Th., 182, 183 Thomas, J. H., 154, 160 Thomas, L., 217, 262 Thomas, M., 21, 39, 50 , Thomas, W. H., 274, 282,309, 313 Thompson, J. F., 288, 312 Thresher, C. L., 288, 309 Tibbitts, S., 300, 308 Tittsler, R. P., 95, 115 Toabe, R., 97, 114 Tobin, B., 217, 264 Tourtellotte, M. E., 218, 231, 262, 263 Tourteliotte, M. F., 213, 255, 265 "release, S. F., 287, 309 Trentini, W. C., 74, 77 Tret'yakova, A. N., 54, 55, 77 Trimble, R. B., 25, 52 Troescher, C. B., 289, 308 Trussell, P. C., 19, 50, 51 Tully, J., 219, 220, 261 Tully, J. G., 213, 217, 219, 220, 234, 267 Tunevall, G., 306, 313 Turner, A. W., 212, 216, 218, 229, 230, 255,267 Turner, H. C., 250, 267 Turri, M., 318, 323, 329 Twort, D. N., 51, 113 i)

U Ueno, T., 180, 181, 183 Ushijima, T., 155, 160

V Valdesuso, J., 222, 265 Valentine, R. C., 322, 329 Vallee, B. L., 290, 312 Vaughn, R. H., 90, 113, 115 Veen, W. L. van, 24, 47, 51 Verloop, A,, 286, 290, 311

Vernon, L. P., 57, 76 Vishniac, W., 19, 52, 56, 77, 288, 310 Vogt, M., 233, 262 Volcani, B. E., 173, 183, 287, 311 Von Stosch, H. A., 291, 313

W Waitz, S., 25, 52 Waksman, S. A., 18, 33, 52 Walden, C. C . , 19, 50 Walker, N., 16, 31, 51 Walker, P. D., 94, 95, 96, 102, 112, 113 Walker, T. J., 290, 309 Walls, B. E., 246, 248, 250, 266 Wangersky, P. J., 291, 309 Wann, F. B., 289, 310 Wansor, J., 290, 310 Ward, J. R., 220, 235, 262 Warrack, G. H., 96, 113, 114 Wassink, E. C., 59, 68, 70, 76, 77 Watanabe, A., 60, 64, 73, 76, 77, 288, 289,310,313 Watson, R. F., 243, 244, 261 Watson, W. A., 218, 219, 231, 323, 261, 262,267 Watt, A., 291, 308 Weaver, R. H., 108, 113 Webber, M. M., 174, 175, 183 Webster, D. A., 64, 77 Weiss, J. E., 80, 115 Welch, F. V., 213, 261 Wentholt, H. M. M., 217, 266 Wertlake, P. T . , 243, 266 Wessel, G., 288, 308 West, G. S., 18, 52 Wetzler, T. F., 90, 91, 108, 114, 115 Whisenand, A., 290,309 White, D., 19, 50 White, D. C . , 146, 349 White, R. J., 216, 261 Whittlestone, P., 216, 219, 232, 233, 234, 238, 239, 246, 254, 257, 258, 216,263,267 Wieringa, K. T., 56, 77 Wiessner, W., 290, 312 Wildy, P., 81, 113 Williams, J. H., 289, 311 Williams, M. H., 218, 231, 267 Williams, P. M., 300, 308

344

AUTHOR INDEX

Williams, R. E. O., 217, 250, 261, 262 Willis, A. T., 81, 88, 91, 97, 98, 99 108, i l l , 115 Willoughby, L. G . , 3, 50 Wilson, B. W., 287,309 Wilson, C. E., 221, 264 Wilson, E., 22, 52 Wilson, W. B., 291, 292, 308, 313 Wilson, W. J., 87, 115 Windsor, G. D., 250, 261 Winogradsky, S., 3, 6, 8, 17, 18, 20, 37, 52, 194, 210

Wise, D. L., 288, 313 Wittler, R. G., 212, 241, 252, 262, 264 Wolfe, R. S., 18, 32, 33, 48, 50, 52 Wolff, S. M., 217, 267 Wolin, M. J., 148, 149 Wong, D., 217, 222, 229, 246, 265 Wong, D. C., 222,265,267 Wong, S. C., 220, 264 Wood, S. C . , 237, 265 Woods, G. T., 234, 265 Woolley, D., 19, 52 Woolley, J. T., 288, 308 Wollman, E. L., 326, 329

Wright, B. M., 81, 115 Wright, D. N., 213, 264 Wyss, O., 316, 329

Y Yamamoto, R., 218, 220, 261, 262, 267 Yevich, P. P., 221, 264 Yoder, H. W., 220, 221, 267 Yokota, M., 307, 313 York, G. K., 90, 115 Youatt, J. B., 19, 50 Young, V. M., 217,243,266,267 Yu, B., 219, 220,261

z Zahalsky, A. C., 65, 66, 67, 76, 290, 310 Zahl, P. A., 291, 292, 293, 310, 311 Zarafonetis, C . J. D., 223, 229, 265 Zehnder, A., 288, 293, 310, 313 Zeuther, E., 313 Zhukova, R. A., 194, 210 Zinder, N. D., 322,329 Zyl, J. G. van, 148, 149

Subject Index A Abomasum, 134 Acetate algae, 270 Acetic acid, in rumen fluid, 135 Achromatiaceae, 5 Achromatium, 5, 6 enrichment culture for, 18 isolation and cultivation of, 16-18 A. oxaliferum, 6 A. mobile, 6 Acidophilic algae, 273, 298 Actidione, see also Cycloheximide, 56, 191,230 Actinomyces, phages of, 316 Aeration, of algal cultures, 282-283 Aerobacter aerogenes, lysis by myxobacteria, 195-196 “Aerobacter circle plates”, for myxobacteria, 196 Aerobes, see also under sgecific names, oxygen removal by, 87 redox potential of systems in, 117 spore-forming, elimination of, 90 thermophilic, 166 “Agnotobiotic” algal cultures, 272 Agrobacterium radiobacter, phage of, 316 Algae, see also under specific names, also Blue-green, etc., acidophilic, 273, 298 aerating gas for, 283 aeration and shaking of cultures of, 282-283 agar methods for manipulation of, 276-277 “agnotobiotic” cultures of, 272 amino acids and, 270-271, 272 ammonium ion utilization by, 270,

300 antibiotic techniques for, 277-279 antibiotic tolerance of, 278 “axenic” cultures of, 272 biphasic media for, 284, 298 blooms of, 18 15

buffering of media for, 302 carbon dioxide for, 270, 282-283 carbon nutrition of, 270 centrifugal concentration of, 274 “chemostat” culture of, 304 continuous culture systems for, 303307 culture media for, 286-303 defined, 287-290, 294-296, 300303 undefined, 286-287 culture sheets for, 284, 285 defined media for, 287-290, 294-296, 300-303 desiccation resistant, 273 enriched water media for, 299-300 enrichment techniques for, 273 “Erdschreiber” media for, 300 glassware cleaning for, 285-286 growth factors for, 272 hexose-utilization by, 270 illumination of, 280 iron for, 271-272, 282, 284, 301 isolation methods for, 272-279 light-limited cultures of, 304 maintenance of, 279-286 manipulative techniques for, 273-277 media for culture of, see under culture, 286-303 media sterilization, 284 micro-pipette techniques for, 276, 277 moulds during isolation of, 279 myxobacterium parasitizing, 196 N.E.R.C. Culture Centre for, 293 nitrate utilization by, 270, 300 nitrogen nutrition of, 270-271, 300301 nutrient concentration in media for, 302 nutrition, synopsis of, 270-272 oxidative assimilation of carbon compounds by, 270

346

SUBJECT INDEX

Algae-cont. pasteurization of media for, 284, 298 p H and ion assimilation by, 270, 271 phosphorous nutrition of, 271-272, 301 phototaxis in isolation media for, 276 plating techniques for, 274, 276-277 purines as nitrogen source for, 271 record sheet for cultures of, 284, 285 sexual, 273 silicon nutrition of, 271, 301 single cell manipulation of, 274 “sloppy” agar media for, 303-308 soil extract media, 299 soil-water media for, 303 special methods for, 303-308 sterility-test media for, 284 sterilization of media for, 283-284 sugar-utilizing species of, 270 sulphur nutrition of, 271, 301 synchronization techniques for, 307 synchronous cultures of, 307-308 temperature of storage of, 280-282 thermophilic, 273 trace metals and, 272, 286, 293, 296297 “turbidostat” cultures of, 303 vitamins for, 298 washing techniques for, 273 water for media for, 286 p-Aminobenzoic acid, for Rhodopseudon m a s palustris, 68 Ammonia-oxidizing autotrophs, 4 isolation and cultivation of, 12-1 6, 31 medium for, 28-29,31 Ammonia production, from threonine by anaerabes, 156 Amoebobacter, 5, 8, 19-20 A. bacillus, 8 A . roseum, 8 Ampicillin, in mycoplasma isolation, 228 Amylolytic rumen bacteria, 140 Anabaena, media for, 287, 288 A. cylindrica, 53 growth of, 62 media for, 288 A . variabilis, 53 carbon dioxide requirement of, 63

glucose incorporation by, 63 light intensity for growth of, 73 nitrogen source for, 62 stock culture maintenance of, 60 Anacystis, medium for, 289 A . nidulans, 53, 63 culture purification of, 56 light and, 73, 75 stock culture maintenance of, 60 Anaerobes, agar plates for, 152 anaerobiosis for, 81-88 bile stimulation of, 154, 157 carbon dioxide for, 86 carbohydrate fermentation by, 154 colonial appearance of, 95-96 contaminants of, 93 continuous culture of, 148 cultural characteristics of, 96 cultural techniques for, 89-104 differential heating for isolation of, 104 enrichment culture of, 103 Gram-negative, non-sporing, 152160 antibiotic sensitivity of, 158 biochemical tests for, 154-1 58 gelatin liquefaction test for, 158 general precautions for, 152-153 growth tests and morphology of, 153-1 54 hydrogen sulphide production by, 157 indole production by, 157 isolation techniques for, 158-1 59 media for, 153-1 59 methods for, 152-1 60 Gram-reaction of, 93 growth methods for, 81-89, 152-153 growth tests for, 153-154 haemolytic, 96 halophilic, 173 histotoxic, 101 indole production by, 101 inoculation of media with, 91-92 isolation and purification of, 102-104 liquid media for, 152-1 53 media for, 89-90,111-112,153-159 methods for, 80-115, 152-160 morphology of, 93

SUBJECT INDEX

Anaerobes-cont. motility of, 95 neurotoxic, 101 non-haemolytic, 96 oxygen toxicity to, 95 pathogenic, 101-102 pure cultures of, 92-93 Reinforced Clostridial Agar for, 153 Robertson’s broth for, 101 rumen of, see aZso Rumen bacteria, 133-149 selection techniques for, 90-91 serology of, 102 spore morphology of, 94 stock cultures of, 92 spore-forming, techniques for, 80115 strict, colonies of, 129 cultivation by roll tube method, 117-132 dilution of, 130 media for, 126-127 media sterilization; 1 2 7 428 oxygen-free media for, 120,-126 picking colonies of, 129-130 pure cultures of, 128-1 31 redox potentials for, 118 roll tube cultivation of, 117-132 storage of cultures, of, 131 surface culture isolation of, 102103 swarming growth of, 103 thermophilic, 166 threonine deaminase test for, 154, 155 toxins of, 96 VL medium for, 153 volatile fatty acids from, 154-1 55 Anaerobic environment, production of, see a h Oxygen removal, 81-88 Anaerobic jars, Baird and Tatlock, 82, 83, 84 electrically operated, 82, 84 Gram-negative anaerobes, for, 152 McIntosh and Fildes, 81-82 opening of, 86 setting-up of, 83-85 Anaerobic media, see also under Anaerobes and Media, rumen bacteria, for, 135-146 158

347

Anaerobic rumen bacteria, see under Rumen bacteria Anaerobic techniques, see also under Anaerobes and Anaerobiosis, strict anaerobes, for, 117-132 Anaerobiosis, carbon dioxide and, 86 degrees of, 117 indicators for, 85-86 liquid to gas volume for, 119 methods for achieving, 81-88 oxygen and, see also under Oxygen, 117-1 20,120-126 oxygen-free media for, 120-1 26 procedures for, 120-132 reducing agents for, 88, 127 zone in freshwater lakes, 2 Animals, pathogenic anaerobe testing with, 101-102,110 Antibiotics, algae, for purification of, 277-279 anaerobes, as selective agents for, 91 Bacteroides resistance to, 158 mycoplasmas, action on,213,228,229 myxobacteria, of, 201 myxobacterial culture, use in, 191 uni-algal cultures, use for, 56 Antifoam agents, for autotrophic media, 40 Antigens, mycoplasmal, 250, 252 Antisera, towards mycoplasmas, 221222 Antitoxins, towards anaerobes, 96, 97, 101 Aphanocapsa, medium for, 289 Archangiaceae, illustrations of, 186, 187 medium for, 199 purification of, 202 Archangium spp., characteristics of, 189 fruiting bodies of, 205 isolation of, 192, 193 maintenance of, 209 A. primigenum, illustration of, 186, 187 stalked cysts of, 205 Arginine deiminase, see L-Arginine iminohydrolase L-Arginine iminohydrolase, 237

348

SUBJECT INDEX

Arthritis, mycoplasmas and, 218, 219, 220,221 Artificial rumens, 147-148 Asparagopsis, medium for, 291 Astasia, 270 media for, 289, 290 Asterionella, media for, 291, 292 Asterococm canis, 220 A . mycoides, 212 Asticcaulis, phage of, 316 Athiorhodaceae, aerobic growth of, 74 elective medium for, 58 growth conditions for, 73-74 growth media for, 37-39, 65-68 isolation and culture of, 21-22, 57-59 light intensities for, 74 pigmentation of, 74 vitamin requirements of, 68 Autotrophic way of life, see also Autotrophs, 25-26 Autotrophs, see also under specijic names, environmental site in nature of, 1-3 facultative, 26 freshwater lakes, in, 1-26 groups of, 4-12 isolation and cultivation of, 12-25 media and methods for, 26-49 oxygen deficit and, 2 sampling methods for, 3 sulphate reducing, 2 “Axenic” cultures of algae, 272 h i d e , as selective agents for anaerobes, 90 Azotobacter, phage of, 316

B Baars’ medium, for Desulpbibrio, 39 Bacillariophyceae, sugar-utilizing, 270 Bacillus sp., phage of, 316 B. stearothwmophilus, as a thermophile, 164 Bacteria, see itnde~spec@ names and categories Bacteriochlorophyll, 57, 65,74 Bacteriocin, 324 Bacteriological diagnosis, see under Diagnosis -

Bacteriological samplers, 3 Bacteriophages, 317-325 aeration for growth of, 317 adsorption to host of, 317 carrier strains of host, from, 323-324 chloroform treatment of, 320 DNA type, 322-323, 324 enrichment procedures for, 317, 321 filamentous, 323 filtration of, 320 fimbriae, adsorption onto, 323 freeze-drying for preservation of, 328 growth conditions for, 317-318 heat treatment of, 320 high-titre stocks, preparation of, 325327 host bacteria for, 316 inducible, 326, 327 ionic environment for, 317 isolation of, 317-325 lysogenic strains of host, from, 323324 plaque size of, 317, 323 plating methods for isolation of, 318 preservation of, 328 purification of, 324-325 RNA type of, 318, 319, 322-323, 324 separation from bacteria, 320 sex pili, adsorption onto, 323 sources for isolation of, 321-323 temperate, 322, 324, 326-327 typing purposes, for, 316 virulent, 324 Bacteroidaceae, see also Gram-negative anaerobes, antiobiotic sensitivity of, 158 bile stimulation of, 154, 157 biochemical tests for, 154-158 carbohydrate fermentation by, 154 dyes, reactions to, 158 fatty acid production by, 153-154 gelatin liquefaction by, 158 growth tests and morphology of, 153154 hydrogen sulphide production by, 157 indole production by, 157 isolation techniques for, 158-1 59 media for, 152-1 53, 158-1 59

349

SUBJECT INDEX

Bacteroidaceae-cont . methods for, 152-160 threonine deaminase test for, 154, 155 Bacteroides spp., dyes, inhibition by, 158 isolation media for, 159 media for, 153-154 methods for, 1 52-1 60 B. amylophilus, carbohydrates fermented by, 137 B. fragilis, growth tests for, 153-154 B . melaninogenicus, growth tests for, 152-154 isolation medium for, 159 Bacteriolytic myxobacteria, 195-196 Bacto PPLO broth, for mycoplasmas, 226 Bark, of trees, myxobacteria from, 192 Beggiatoa, cultivation of, 5 , 6 enrichment culture media for, 64 isolation and cultivation of, 16-18 media for, 287, 288 pure cultures of, 18, 33 B . alba, 6 B . arachnoidea, 6 B. minima, 6 B . mirabilis, 6 Beggiatoaceae, 5 Beggiatoales, 5, 6 Benzyl penicillin, mycoplasmal isolation using, 228 Benzyl viologen, as redox indicator for media, 127 Bicarbonate, in algal nutrition, 270, 282 Bile, stimulation of anaerobes by, 154, 157 Biochemical tests, anaerobes, for, 154-158 rumen bacteria, for, 145-146 Biotin, algal growth, for, 272 Athiorhodaceae, for, 68 Biphasic media, for algae, 284, 298 Blood agar, see under Fresh, Horse, Ox, etc. B.T.L. anaerobic jar, 82, 83, 84 Blooms, of algae, 18 Blue-green algae, see also Myxophyceae and under specific names, clumping during growth of, 73

culture media for, 55, 60-64 culture purity criteria for, 57 filamentous, 54 growth of, 53-76 growth media for, 60-64 heterotrophic, 63 isolation and culture of, 53-57 large-scale production of, 63 light intensity for, 73, 75 maintenance of cultures of, 59-60 micro-elements for, 62 “morphogenetic substance” for, 56 phototactic movements of, 54 physical conditions for growth of, 69-73 sulphide and, 59 temperature for growth of, 73 trace-elements for, 54 Bordatella bronchiseptica, phage of, 316 Borrelomyces, see also Mycoplasma, 212 Bovine mycoplasma strains, 229-232 Brachiomonas, medium for, 290 Brilliant green, anaerobe resistance to, 158,159 Brines, see also under Salt, Sodium chloride, etc., halophiles from, 169, 171, 180 Brucella spp., phage of, 316 Bryophytes, purification of, 278 Buffers, for algal media, 302 Butyric acid, rumen fluid, in, 135 Gram-negative anaerobes, and, 154 Butyrovibrio, continuous culture of, 148

C

Calothrix, medium for, 292 C. elenkinii, nitrogen for growth of, 62 Caprine mycoplasma strains, 23 1-232 Capryllate, enrichment for Rhodospirillum using, 58 Capsule staining, 94 Carbohydrate fermentation, Gram-negative anaerobes, by, 154 rumen bacteria by, 145 Carbon dioxide, algal nutrition, in, 270, 282-283, 304 anaerobic jars, addition to, 86

350

SUBJECT INDEX

Carbon dioxide-cont. Chlamydomonads, solid media for, 303 blue-green algae, for, 63 Chlamydomonas chlamydogama, media control of, in algal cultures, 282for, 288 283 C. moewusii, media for, 287, 289 generators, 282, 283 C. mundana, media for, 289, 290 Gram-negative anaerobes, and, 152, C.pallens, medium for, 289 C. rheinhcrdii, media for, 288 159 rumen bacteria, for, 134, 136, 146 C. pulsatilla, acetic acid requirement of, strict anaerobes, gas phase for, 119270 120 Chloramphenicol, algal purification Carbon-dioxide-bicarbonate buffer, for with, 277,278 Chorella, anaerobic media, 126, 132 illumination for, 280 Carbon monoxide, observation and oxidation of, 45 media for, 287, 288, 289 C. vulgaris, medium for, 288 production method for, 42 Chlorobacteriaceae, bacteria oxidizing, 11, 23,42-44 elective medium for, 58 Carboxydowtonas, 11 isolation of, 23 enrichments of, 54 group, 11 media for, 42-44 Carotenoids, growth media for, 37, 69 blue-green algae, of, 75 illustrations of, 10 halophiles, of, 171 isolation and culture of, 20-21, 57mycoplasmal synthesis of, 213 58 Cat, mycoplasmas of, 220 physical conditions for growth of, Caulobacter sp., 73-74 stock culture maintenance of, 60 Nostoc, interaction with, 56 Chlorobium, 10, 11 phages of, 316, 322, 323 Caulobacteriaceae, isolation and cultivation of, 20-21 medium for, 35-36 group, 12 illustrations of, 14 Chlorobium chlorophyll, 57 isolation and cultivation of, 24 Chlorobium limicola, media for, 69 Cell envelope, of halophiles, 173 C. thiosulphatophilum, media for, 69 Cell membrane, of mycoplasmas, 213, Chlorochromatium, 11, 20-21 240,242,258 C. glebulum, 10 Cellulolytic rumen bacteria, Chlorococcales, fermentation substrates of, 137 media for, 287 media for, 139, 146 solid media for, 303 tests for, 145 sugar-utilizing, 270 Cellulolytic myxobacteria, sources of, Chlorogloea fritschii, 194-195 heterotrophic growth of, 63 Cellulose-containing media, for myxostock cultures of, 59 bacteria, 198-1 99 Chlorophyta, Cephaloridine, 21 3 cyclic photosynthesis in, 270 Ceratium, medium for, 291 medium for, 288 Chara, medium for, 289 naturalistic morphology of, 272 “Chemostat” culture, of algae, 304, 305, Chloropseudomonas ethylicum, 306 carbon sources for, 57, 69 Chilomonas. medium for. 289 light intensity and, 74, 75 C. paramoecium, medium for, 289 Chl&otetracycl;ns, for algal purification, Chlamydobacteriaceae, 12, 13, 24, 47 277

SUBJECT INDEX

Cholesterol, mycoplasmal cell membrane, in, 213, 240 mycoplasmal growth and, 213, 235, 250 Chondrococcus spp., characteristics, of, 189 isolation of, 192 C. columnaris, agar for isolation of, 197 fish parasite, as, 192, 196 phage of, 316 C.coralloides, eubacteriolytic nature of, 195 Chondromyces sp., characteristics of, 190 enbacteriolytic forms of, 195 fruiting body morphology of, 205 isolation of, 192 maintenance of cultures of, 207, 208, 209 C. apiculatus, illustrations of, 188, 189 C. catenulatus, illustration of, 188, 189 C. crocatus, illustration of, 188, 189 Chromatophores, algae lacking, 270 Chromatium, 5,9 isolation and cultivation of, 19-20 media for, 35, 36, 68, 70 Chromatiurn D , 68 C. minus, medium for, 68 C . okenii, medium for, 69 C. warmingii, medium for, 69 Chronic respiratory disease of poultry, mycoplasmal, 221 Chroococcus, media for, 287 Chrysomonads, medium for, 292 Chrysophyta, silicon for, 271 Citric acid, as iron complexer, 272 Clathrochloris, 10, 11, 20-21 C . hypolimnica, 10 Cloning methods, for mycobacterial purification, 238-239 Clonothrix, 12, 13, 24-25 C. fusca, 13 Clostridia, see also under specific names, anaerobic, 93, 109-110 animal inoculation and protection tests for, 101 classification of, 108-1 11 1585

351

culture from wound material, 104105 diagnosis of, 97-98 identification scheme for, 100 non-pathogenic, 80, 108 non-saccharolytic, 101 oxygen tolerant, 103 pathogenic, 101-102, 104-107, 108111 reactions on lactose-egg yolk-milk agar, 99 sqccharolytic, 101 wound examination for, 104, 105106,107 Clostridial infections, diagnosis of, 104107 Clostridial myonecrosis (gas gangrene), diagnosis of, 105 Clostridium bifermentans, culture from pathological material of, 106 identification scheme for, 100 indole production by, 101 key to, 109 reactions of, 99, 110 Cl.bifermentans, -sordellii group, characteristics of, 97, 98 C1. botulinum, classification of, 80 Cl. sporogenes, differentiation from, 95,101 identification of, 100, 101, 109 reactions of, 97, 100, 110 Cl. butyricum, capsule staining of, 94 reactions of, 99, 110 C1. capitovale, reactions of, 99, 110 C1. carnis, key to, 109 C1.chauvoei, identification of, 94 key to, 109 reactions of, 110 CI.cochlearium, reactions of, 99, 110 Cl. fallax, reactions CI. fallax, reactions of, 99, 110 Cl. histolyticum, aerotolerant growth of, 93 culture from pathological material, 106,107 diagnosis of, 98

352

SUBJECT INDEX

C1. histolyticum-cont. identification scheme for, 100 key to, 109 reactions of, 99, 110 CE. innominatum, reactions of, 110 C1. multifermentans, reactions of, 110 C1. oedematiens, anaerobic growth of type D, 92 animal inoculation tests for, 102 cultural characteristics of, 97, 98, 99 diagnosis of, 98 Gram-reaction of, 93 identification scheme for, 100 immuno-fluorescent staining of, 95 key to, 109 pathological material, from, 106 reactions of, 99, 110 wound examination for, 105, 107 C1. perfringens, phage of, 316 Cl. septicum, identification of, 94, 100 key to, 109 reactions of, 99, 110 wound examination for, 106, 107 C l . sordellii, identification scheme for, 100 indole production by, 101 key to, 109 pathological material, from, 106 reactions of, 99,110 C l . sphenoides, reactions of, 99, 110 Cl. sporogenes, Cl. botulinum, differentiation from, 95,101 C1. welchii infections, effect on, 108 identification of, 100 pathological material, from, 106 phage of, 316 reactions of, 97, 99, 110 swarming growth of, 103 C1. tertium, aerotolerant growth of, 93 identification of, 100 key to, 109 reactions of, 99, 110 Cl. tetani, colonial appearance of, 95 Gram-reaction of, 93 identification of. 100 immuno-fluorescent staining of, 95

indole production by, 101 isolation by surface culture of, 102 pathogenicity of, 102 reactions of, 99, 110 spore morphology of, 94 swarming growth of, 102 tetanus, and, 104 wound material, from, 105 C1. tetanomorphum, reactions of, 99,

110 CI. welchii, animal inoculation tests for, 102 capsule staining of, 94 colonial appearance of, 95 cultural characteristics of, 96, 97, 98 diagnosis of, 98 enrichment culture of, 103 food poisoning with, 106 haemolysis by, 96 identification scheme for, 100 infection and capsulation of, 105 isolation of, 91 key to, 109 pathological material, from, 106 reactions of, 99, 110 selective agents for, 91 spores of, 94, 103, 104 toxins of, 96 Cocci, characteristics for anaerobic, 93 Coccolithus huxlii, medium for, 292 Coliform bacilli, oxygen removal using, 87 Coli-phages, 316, 321, 322 Colonies, anaerobes of, 129 growth measurement of, 248 mycoplasmas, of, diameter of, 248 illustrations of, 214 microscopy of, 252-254 Conjunctivitis, mycoplasmas and, 217, 220 Contagious aglactia, mycoplasmas and, 212, 218, 219, 221, 231 Contagious bovine pleuropneumonia, mycoplasmas and, 212, 216, 21 8 Contagious caprine pleuropneumonia, mycoplasmas and, 216, 218,

..

311 ~

353

SUBJECT INDEX

Continuous culture, algae, of, 303-307 “artificial rumens” and, 147-148 microalgae, for, 75 mycoplasmas, of, 250 rumen as, 134 Cooked-meat medium, for anaerobes, 89,112 Cooked meat particles, for anaerobiosis, 89 Copepods, purification of, 278 Corynebacterium diphtheriae, phage of, 316 C.flaccumfaciens, phage of, 316 Crenothrix, 12, 13 isolation and cultivation of, 24-25 C. polyspora, 13 Crenotrichaceae, 12 Crystal violet, anaerobe resistance to, 158 Culture media, see under Media and under names of micro-organisms Cyanidium media, for, 288, 289 Cyanophora, media for, 287, 288 Cyanophyceae, heterotrophic colourless, 64 Cyanophyta, agar technique for, 277 colourless, 64 media for, 287, 288, 289, 291 Cyclic photosynthesis, in algae, 270 Cycloheximide, see also Actidione, 191, 192 Cylindrogloea, 10, 11 isolation and cultivation of, 20-21 C . bacterifera, 10 Clyindrotheca, media for, 291, 292 Cysteine, reducing agent for media, 89, 127, 136, 137, 138, 139, 140, 141, 142, 144 Cytophaga, 187 Cytophagaceae, 187

D Dafaalla’s medium, 230, 231 Dark-field microscopy, of mycoplasmas, 255 Deamination of amino acids, 145 “D” catalyst, see under Engelhard

Deep-culture tubes, for photosynthetic bacteria, 20 Deoxyribonucleic acid, see also DNA, mycoplasmal media, for, 227 Desiccation-resistant algae, 273 Desulphovibrio, anaerobic incubation of, 40 isolation and cultivation of, 11, 21-22 isolation media for, 39-40 pure cultures of, 39-40 Diagnosis, of cloutridial infections, 104107 clostridial myonecrosis, 105-106 food poisoning and, 106 tetanus, 104-105 Diaminopimelic acid, 173 Diatoms, media for, 290, 291, 292, 293 Dichromate-sulphuric acidcleaning, 285 Differential heating techniques, 104 Diluting fluids, for rumen bacteria, 136 Dinoflagellates, media for, 290, 292, 293 Diseases, mycoplasmal, 212, 216-221 Dithionite, as reducing agent for media, 127 DNA phages, 322-323,324 Dog, mycoplasmas from, 219-220 Dunaliella, medium for, 292 Dung, myxobacteria from, 192, 193-194 myxobacterial media containing, 198

E Ectocarpus, medium for, 291, 292 EDTA, for algal media, 272, 301 Eggerthella conz‘exa, see Bacteroides fragilis Egg-yolk agar, 96, 97, 98, 103, 110, 111 Egg-yolk emulson, 111 Eh, media, of, 142 rumen, of, 134 Ehrlich’s reagent, 157 Elective culture, see under Enrichment and name of micro-organisms concerned Electron microscopy, of mycoplasmas, 242,257-258 Electrophoretic analysis of cell proteins, 241,251

354

SUBJECT INDEX

Elementary bodies, of mycoplasmas, 213,253,254,255 Engelhard “D” catalyst, for anaerobic jars, 84 Enrichment culture, 273 algae, for, 273 anaerobes, of, 103 blue-green algae, of, 53-57 hydrogen-reducing bacteria, for, 23 nitrite-oxidizing bacteria, for, 16 Nitrosomonas spp., for, 13 photosynthetic bacteria, for, 19-20, 20-2 1 sulphur-oxidizing bacteria, for, 17 thiobacilli, for, 18,19 Winogradsky cylinders for, 3 Enterobacteriaceae, sex pili and phages of, 323 Enzootic pneumonia of pigs, as mycoplasma1 disease, 216, 219 Epilimnion, 1 , 2 “Erdschreiber” media, for algae, 300 Erythema multiforme, mycoplasmas and Erythema multiforme, mycoplasmas and, 217,223 Erythromycin, as mycoplasma inhibitor, 213 Eschrichia coli, oxygen removal by, 142 Euglena gracilis, illumination for, 280 media for, 287,289,290 Eugleninae, cyclic photosynthesis in, 270 F Facultative autotrophs, 26 Facultative psychrophiles, 162 Fatty acids, Gram-negative anaerobes, from, 154155 rumen, of, 135 Febrile respiratomy tract disease, mycoplasmas and, 217 Fermentation, media, 99, 112 rumen bacteria, by, 134,135,137, 143, 145 Fermenters, for thermophiles, 166 Ferribacterium, 12, 15 isolation and culture of, 24-25

F . dubium, 15 Ferrobacilius, 12 isolation and culture of, 24-25 F. jerro-oxidans, 49 Ferrous sulphide, formation in nature, 2 Filamentous blue-green algae, isolation of, 54 Filamentous iron organisms, 3 Fildes and McIntosh indicator, 85 Filter-sterilization, of media, 127, 284 Flagellates, media for, 290,291,292 Flexibacter sp., cultivation of, 64 Flexibacteria, blue-green algae and, 64 Fluorescent antibody technique, anaerobes, for, 94 mycoplasmas, for, 237 Fluorescent light, for algae, 280 Fluorescent staining method, for nucleic acid, 322 Food poisoning, 106 Formic acid, in rumen fluid, 135 Foster’s autotrophic medium, 45 Freeze drying for preservation of cultures, mycoplasmas, of, 260 myxobacteria, of, 207-208 Fresh blood agar, 96, 111 Freshwater lakes, autotrophs in, 1-3 Friedinger water bottle, 3 Fruiting bodies, of myxobacteria, 187, 189-191,205,208-209 Fulgostatin, 56 Fusiformis sp., colonies of, 95 F . melaninogenicum, colonial pigmentation of, 96 F. nodosus, growth tests for, 152, 154 Fusijcrmis, characteristics of, 93, 96 Fusobacterium, biochemical tests for, 154 dye resistance of, 158 growth tests for, 153-1 54 medium for isolation of, 158 methods for, 152-160 F. polymorphum, 152

G Gallionella, 12, 14 isolation and cultivation of, 24-25, 48-49

SUBJECT INDEX

355

H G .ferruginea, 14 Habitat-simulating media, for rumen G . tenuicaulis, 14, 48 bacteria, 136-1 38 Gas dispersion ball, porous, 41 Gas gangrene (Clostridial niyonecrosis), Harmatin, as pigmerit of Fusiformis sp., 96 diagnosis of, 105 “Gaspak System”, for hydrogen genera- Haematococcus pluaialis, media for, 287, 288,289,290 tion, 83 Haemolysins, production by anaerobes, Gelatin, 96 anaerobe reaction on, 98,110,145,158 liquefaction test, 145,158 Haemolysis, caused by anaerobes, 96 Gelatinase, 98 Haemophilus influenzae, phage of, 3 16 Genito-urinary tract sepsis, mycoplas- Half-antitoxin plates, preparation of, 91-92 mas and, 222 uses of, 97,98,106 Gentian violet, for anaerobe selection, Ilalobacterium cutirubrum, 90 growth measurements for, 179 Giesma stain, for mycoplasmas, 253 halophile, as, 170 Glassware, for algal culture, 285-286 radiation resistance of, 171 Glucose, synthetic medium for, 176 clostridial fermentation of, 99, 110 reducing agent for anaerobiosis, as, 89 H . halobium, Glucose-gelatin medium, 98,112 halophile, as, 170 Glycerol media, for rumen bacteria, 143 synthetic medium for, 176 H . salinarium, Goats, see also Caprine, halophile, as, 170 mycoplasmas and diseases of, 212, 281-219,221,231 malic dehydrogenase of, 178 Gram-negative anaerobes, see also under synthetic medium for, 176 Anaerobes, 13. trapanicum, 170 Halophilies, see also under specific names, biochemical tests for, 154-1 58 general precautions for, 152-1 53 169-183 anaerobic, 173-174 growth tests and morphology of, 153-154 carotenoid pigments of, 171 cell envelopes of, 173 isolation techniques, for, 158-1 59 classifications for, 170, 171 media for, 153-159 methods for, 152-160 definitions of, 169, 170 non-sporing, methods for, 152-160 extreme, 170-180 Gram-reaction, for anaerobes, 93 growth requirements CJf, 171-174, Green photosynthetic bacteria, see 180-1 81 Chlorobacteriaceae and under ionic requirements of, 171-172, 180 isolation and enumeration of, 177-1 78 specific names I Green sulphur bacteria, see under large-scale production of, 178-179 Sulphur bacteria and under maintenance of, 179 specific names media for, 174-177, 181 Griess-Llosvay reagents, for nitrite, 31 moderate, 180-182 Growth, see also under names of specific occurrence of, 170-180 micro-organisms, pathogenic, 182 mycoplasmas, of, 245-249 production on a large scale of, 178-1 79 Growth factors, see also Vitamins, 14 red pigmented, 170,171, 172,173,175 Growth media, see under Media 177,178,180 Gymnodinium breve, media for, 291, 292 Halophilic bacteria, see also f-lalophiles, Gyrodinium, medium for, 292 169-1 83

356

SUBJECT INDEX

Hapalosiphon laminosus, medium for, 54, 63 Haploangium, 190 Hayflick's media, for mycoplasmas, 226, 23 1 Heated-blood agar, 95 Heat-exchangers for study of thermophiles, 165 Hemiselmis virescens, medium for, 291 Herbivores, rumen bacteria and, 133 Histotoxic clostridia, 101 Hoagland trace element solution, 36 Horse-blood agar, use of, 95, 96, 103 Horse serum media, for mycoplasmas, 226,228 Human-serum agar, use of, 96, 97 Humans, mycoplasmas from, 222-224 Humates, as iron complexers, 272 Hungate roll-tube technique, see under Roll-tube Hutner's methods and media, for sulphate-reducing bacteria, 37-39 Hydrodyction, media for, 288 Hydrogen, anaerobic jars, for, 82-83 anaerobic media, for, 127 oxidation of, 45 Hydrogen bacteria (hydrogen-oxidizing, 11 isolation and cultivation of, 22-23 media for, 40-42 Hydrogenomonas, 11 isolation and cultivation of, 22-23 media for, 40-42 Hydrogen sulphide, Gram-negative anaerobes, from, 157 natural production of, 2,157 reducing agent for media, as, 127 rumen bacteria, from, 145 sulphur bacteria and, 17, 19 H .facilis, 40,41 Hypolimnion, 1,2,16-17

Immuno-fluorescent staining, of anaerobes, 94,102 Incubators, for low temperatures, 163 Indicator media, for anaerobes, 103 Indicator strain, of phage host, 324 Indole, production by anaerobes, 101, 110,157 Indole-3-acetic acid, blue-green algae, and, 57 Infections, human clostridial, 104-107 Infectious aglactia, mycoplasmas and, 212 Infectious catarrh of rodents, mycoplasmas and, 221 Inoculation, of anaerobic media, 91-92 Inorganic sulphur, bacteria oxidizing, 5 Inspissated serum medium, for anaerobes, 98 Intestinal flora, Gram-negative anaerobes from, 152 Ionic requirements of halophiles, 171172,180 Iron, algal nutrition, in, 271-282 metallic, as reducing agent, 89 Iron compound-oxidizing bacteria, 12 isolation and cultivation of, 24-25 media for, 47-49 Iron cycle, 26 Iron organisms, filamentous, 3 Iron-oxidizing thiobacilli, 19 Irradiation, of contaminated algal cultures, 56 Isolation techniques, see also under names of specific micro-organisms, algae, for, 272-279 anaerobes, for, 158-159 autotrophs, for, 12-25 bacteriophages, for, 317-325 Gram-negativeanaerobes, for, 158-159 halophiles, for, 177-178 mycoplasmas, for, 222-238 myxobacteria, for, 191-197 psychrophiles, for, 162-163

I Identification, see also under names of specific micro-organisms, clostridia, of, 100 rumen bacteria, of, 144-1 46

J Jenkin surface-mud sampler, 3, 4, 12, 17, 18, 19, 21, 23 Joint diseases, mycoplasmas and, 221

357

SUBJECT INDEX

K

I< & K plates, for myxobacteria, 192 Kanamycin, Bucterioides resistance to, 158,159 Katodinizim, medium for, 291 Kistner’s medium, for Curboxydomonus, 42

media for, 141 tests for, 145 Loeffler’smedium, for anaerobes, 98 Lucas semi-solid indicator, for anaerobiosis, 85 Lyophilization, see also Freeze drying, blue-green algal cultures, of, 60 photosynthetic bacteria, of, 60 Lysogenic hosts, phages from, 323-324

L Lactic acid, in rumen fluid, 135 Lactobacilli, maintenance of, 147 phages of, 316 rumen of, 143 Lactose, clostridial fermentation of, 99, 110 Lactose-egg media, 97, 99, 100, 111 Lakes, autotrophs in, 1-26 oxygenated zone of, 18 stratification of, 1-3 Lamellibranchs, purification of, 278 Lamprocystis, 5,8,19-20 L. roseo-persicina, 8 Lumpropediu, media for, 287 Larsen’s medium, 21, 37 Large-scale production, halophiles, of, 178-179 Microcystis aeruginosa, of, 63 L-foms, 216 differentiation from mycoplasmas, 239, 240, 251, 252 Lecithinases, 97 Leitz Heine system phase-contrast, 17 Leptothrix, 12,13 isolation and cultivation of, 24-25 media for, 47-48 pure cultures of, 48 L. discophoru, 48 L. ochracea, 13 L . sideropous, 13 Leucothrix mucor, cultivation of, 64 Lieske’s medium, for thiobacilli, 33 Light intensity, bluegreen algae, for, 73,75 photosynthetic bacteria, for, 73-74,75 Lipolysis, by clostridia, 97 Lipolytic rumen bacteria,

M Mackereth oxygen electrode, 3 Macromonas, 5 , 7 isolation and cultivation of, 18-19 M . bipunctata, 7 M . fusiformis, 7 M . minutissima, 7 M. mobilis, 7 Maintenance, see also under Stock cultures, and names of specific orgaqisms, algae, of, 279-286 halophiles, of, 179 mycoplasmas, of, 258-260 myxobacteria, of, 205-209 rumen bacteria, of, 147 Malic dehydrogenase, of a halophile, 178 Maltose, clostridial fermentatiqn of, 99, 110 Manganese compound-oxidizing bacteria, 12 isolation and cultivation of, 24-25 media for, 47-49 Marine bacteria, 169 Mastitis, as mycoplasmal disease, 218, 221 McIntosh and Fildes anaerobic jars, 40, 81-82 Meat-infusion-peptone broth, 89-90 Media, see also under numes of specific micro-organisms, algae, for, 286-303 anaerobes, for, 89-90, 111-112, 120128,153-159 autotrophs, for, 26-49 blue-green algae, for, 55, 60-64 Gram-negative anaerobes, for, 152153 halophiles, for, 174-1 77

358

SUBJECT INDEX

Media-cont. mycoplasmas, for, 225-237 myxobacteria, for, 198-202 photosynthetic bacteria, for, 58, 6569,70-72 rumen bacteria, for, 135-176 Meiklejohn’s medium, 16, 30 Mercury-toluene thermometers, 280281 Mesophiles, use of term, 161 Methane, method for producing, 44 oxidation of, 45 rumen, in, 134, 135 Methane-forming bacteria, 118 Methane-oxidizing bacteria, 11, 23-24 Methanobacterium ruminantium, 142 Methanogenesis, by rumen bacteria, 142 Methanomonadaceae, gases for, 43 group, 11 isolation and cultivation of, 22-23, 23-24 media for, 40-47 Methanomonas, 11 isolation and cultivation of, 23-24 media for, 44-47 Methylene blue, mycoplasmas resistant to, 227 ’ Micrococci, salt-tolerance of, 170 Micrococcus sp. No. 203, 180, 181 M . dentrificans, halophilic variety of, 181 M . hulodenitrificans, 180, 181 M. Zitoralis, as halophile, 170 M. morrhuae, as halophile, 171 M . radiodurans, 171 Microcystis, media for, 288 M . aeruginosa, 63, 73 Micro-elements, for blue-green algae, 62 Microscopy, of mycoplasmas, 252-258 Microstenae, media for, 288 Milk agar, anaerobes on, 98, 110 preparation of, 112 Milk medium,for halophiles, 174 Miller’s medium, for Desulphouibrio, 40 Minimal viable units, of mycoplasmas, 213 Mollicutales, suggested Class, 21 2 Monochrysis, medium for, 293

M . lutheri, medium for, 292 Monodus, medium for, 288 “Monoxemic” cultures, of algae, 272 “Morphogenetic substance”, for bluegreen algal growth, 56 Motility, of anaerobes, 95 Mouse, mycoplasmas from, 220 Mud, bacterial flora of, 1, 2 Nitrobacter sp. from, 16 Nitrosomonas sp. from, 12-16 sampling of, 3, 12 sulphur-oxidizing bacteria from, 1618 Muir’s capsule stain, 94 Mulder and van Veen’s medium, 47 Muramic acid, halophile cell envelopes and, 173 Mycobacterium spp. phages of, 316 Mycoplasma(s),212-267 acid formation by, 248 air bubbles, differentiation from, 242 animal sources, from, 224-225 antibiotic action on, 213, 228, 229 antigens of, 250, 252 antisera towards, 214, 221, 222 arginine metabolism by, 248 artefacts, differentiation from, 242244 arthritis and, 221 avian strains of, 235, 254 bacteria, differentiation from, 241-242 binary fission of, 213, 246, 257 biochemical studies for, 251-252 Boidin’s medium for, 232 bone marrow as source of, 229 bovine strains of, 229-232 BVF-OS medium for 229,230,231 canine strain isolation, 234, 235 caprine strain isolation, 231-232 carbohydrate metabolism by, 248 cat, from, 220 cell count of, 248-249 cell membrane of, 213, 240, 242, 258 cell wall, lack of, 212-213 characteristics of, 212-216 cloning methods for, 238-239 colonies of, 214, 248, 252-254 continuous culture of, 250 cross contamination of, 252

SUBJECT INDEX

Mycoplasma(s)-cont . culture purification of, 238-245 Dafaala’s medium for, 230, 23 1 dark-field microscopy of, 255 differentiation from artefacts, 239-244 digitonin sensitivity of, 240 diseases associated with, 216-221 dog, from, 219-220 electron microscopy of, 242, 257-258 electrophoretic analysis of proteins of, 241,25 1 elementary bodies of, 213, 253, 254, 255 feline strain isolation, 235 filaments in, 213,253,255 fluid droplets, differentiation from, 243 fluorescent antibody method for, 237 freezing cultures of, 259-260 freeze drying for maintenance of, 260 gaseous atmosphere for isolation of, 237 Giesma stain for, 253 glassware for culture of, 226 goats, from, 218-219 Grieg’s medium for, 232 growth of, 245-249 Hamdy’s medium for, 232 handling of, 249-258 Hayflick’s media for, 226, 231 human sources, from, 222-224 identification of, 221, 245 isolation methods for, 222-238 joint diseases and, 221 Kelton’s storage medium for, 259 L-forms, differentiation from, 239241, 251 latent pathogens, as, 224 Mackay’s medium for, 232 maintenance of cultures of, 258-260 man, from, 217 mastitis and, 221 medical or veterinary interest, of, 21 7-220 metabolic activities of, 221 metabolic units of, 248-249 methylene blue resistance of, 227 microscopy of, 252-258 minimal viable units of, 213 morphology of, 252-258

359

motility of, 21 3 mouse, from the, 220 name origin, 212 Nasri’s medium for, 231 nomenclature of, 21 2 non-antigenic medium for, 250 occurrence of, 212 ovine strain isolation, 232 ox, from, 218 pathogenic, 212,216-221, 222-224 p H for growth of, 227 phase contrast microscopy of, 255256 Pillai strains of, 231 porcine strain isolation, 232-233 pseudocolonies, differentiation from, 243 rat strains of, 220 rabbit strains of, 220 reproductive mode of, 246, 257 respiratory diseases and, 216,221,222 rodent strains of, 235 serological studies, for, 249-250 sheep, from, 219 Shepard’s storage broth for, 259-260 Shoetensack’s media for, 234 solid medium, growth on, 246-247 sources of, 216 species of, 221-222 specimens of, 222-225 stained preparations of, 256 sterols for growth of, 212, 213, 235, 240,250 storage of cultures of, 259-260 subculture of, 249 surfaces, growth on, 246, 254 swab samples of, 223, 224 synovitis and, 221 T-strain of, 213, 214, 221, 224, 228, 229,231,246,258 tendo-synovitis, and, 221 tissue cells, differentiation from, 242243 tissue cultures, in, 236-237 titres in liquid media, 246 total cell counts for, 249 transport medium for, 223 transportation of, 222-225, 260-261 turbidity and growth of, 247 type cultures of, 252

M ycoplasma(s)--cont. strains related to, 218-219 urea in media for, 221, 229, 248 titres in liquid media of, 246 veterinary interest of, 217-220 M . neurolyticum viable cell counts for, 248 penicillin inhibition of, 213 Mycoplasma agalactiae, rolling disease in mice, and, 220 contagious agalactia and, 218,219,231 M. orale, strains of, 217, 246, 259 var. bovis, mastitis and, 217 M . pharyngis, 217 M . arthritidis, arthritis and, 220, 221 M . pneumoniae, M . B3, colonies of, 214, 215 antigen, preparation of, 250 M . bovigenitalum, mastitis and, 218, 221 antiserum tests for, 222 M . bovimastitis, 218 colonies of, 214 erythromycin inhibition of, 213 M . bovirhinis, 218 M . canis, 219 human pathogen, as, 216, 217, 227 M . Capri, 218 isolation media for, 225, 227, 236, 237 maintenance of cultures of, 258 M . felis, conjunctivitis and, 220 metabolic activities of, 248 isolation of, 235 pH for, 227 M . fermentans, 217 surfaces, growth on, 246 titres in liquid media, 246 M. gallinarum, colonies of, 214 M . pulmonis, porcine strain of, 234 rodent pathogen, as, 220, 221 poultry pathogen, as, 221 surfaces, growth on, 246 M . gallisepticum, 246, 248, 257 M . salivariuni, 217 , 246 M . gateae, 220, 235 M. spumans, 219 M.granularum, 213, 219,221, 234 M . suipneumoniae, M . horninis, colonies of, 214, 215, 254 isolation media for, 234 electron microscropy of, 258 pelvic sepsis and, 217 enzootic pneumonia and, 219 p H for, 227 isolation media for, 232-233, 234, 237 surfaces, growth on, 246 metabolic activities of, 248 M . hyopneumoniae, see M . suipneumoniae, microscopic examination of, 256 216, 219 pneumonia of pigs and, 216, 225 M . hyorhinis, surfaces, growth on, 246 growth test with, 222 M. synoviae, joint disease and, 219, 221 isolation of, 235 purification problems with, 239 media for, 235, 236 M . iners, 234 NAD requirement of, 236 M . laidlaziii, tendo-synovitis and, 221 sources of, 213, 216, 217, 218, 219, Mycoplasma Reference Centres, 252 234 Mycoplasma Reference Laboratory, 252 sterol-independent growth of, 240 Mycoplasmatales, M . lipophiliae, 2 17 classification of, 212 M . maculosum, 219 nomenclature and type cultures of, M . meleagridis, isolation of, 235 252 M . mycoides, Myringitis, mycoplasmas and, 21 7 original isolation of, 212 Myxobacterales, 187 var. Capri (PG3), 231, 234 Myxobacteria, 182-210 var. mycoides, 216, 221, 224,225 aquatic sources of, 197 media for, 229, 230, 231 bacteriolytic forms of, 195-196 microscopy of, 253, 255 bark of trees, from, 192-193

SUBJECT INDEX

361

M yxobacteria-cont. M. fitlvirs, 186, 187, 205 M. stipitatus, 205 cellulolytic forms of, 194-1 95 cellulose media for, 198-199 M. xanthus, characteristics of, 187 defined media for, 200,201 contamination of, 191 liquid culture of, 204 cultivation and media tor, 198-205 M. stipitatus and, 205 cultivation problems with, 204-205 phage of, 316 culture conditions for, 203 Myxophyceae, see also Blue-green algae, cysts of, 205 development and purification of, defined media for, 199, 200 53-57 dung, from, 193 nitrogen fixation by, 54, 56, 62 dung media for, 198 pure cultures of, 53-57 freezing for maintenance of, 208 species commonly studied, 53 freeze drying for maintenance of, 207stock culture maintenance, 59-60 thermophilic, 54, 63 208 fruiting ability, loss of, 205 ultraviolet light for purification of, 56 fruiting bodies of, 187, 189, 191, 205, 208 growth of, 191-201 N isolation of, 191-197 liquid culture of, 203-204 Nagler reaction, 97 maintenance of cultures of, 205-209 Nannochloris, medium for, 292 media for, Nasri’s medium, for M. mycoides, 231 cultivation of, 198-201 Naumaniella, 12, 15, 24-25 isolation of, 194 N. neustonica, 15 occurrence of, 191 “Navel” rnycoplasma, 21 7 parasitic forms of, 189, 196-197 Navicula pelliculosa, media for, 287, 288 pigmentation change of, 205 Necrotic lesions, anaerobes and, 152 plant-debris as source of, 193 Neomycin, purification of isolates of, 202-203 algae, for purification of, 277, 278 soil as source of, 192 anaerobe selective agent, as, 91, 103 soil storage of, 209 Bacteroides resistance to, 158, 159 S t a n k plates for isolation of, 192, media, preparation of, 92 194-1 95 sulphate, 91,92,103 symbiosis with eubacteria, 204 Neonatal conjunctivitis, mycoplasmas Mycobacteriat-eubacterial association, and, 21 9 204-205 Ncurotoxic clostridia, 101 Myococcaceae, Nicotinic acid, for Athiorhodaceae, 68 genera of, 189 Nile-bluc reduction, by anaerobes, 156 illustration of, 186, 187 Nitrate, media for, 199-200 algae, for, 270, 300 occurrence of, 189 formation by Nitrobacter, 16 purification of, 202 reduction by rumen bacteria, 145-1 46 Myxococcus spp., Nitriloacetic acid, for algal mcdia, 301 characteristics of, 189 Nitrite, cultivation of, 200-201 accumulation by Nitrosomonas, 16 isolation of, 192, 195 determination of, 31 maintenance of cultures of, 207, 208, utilization by Nitrobacter, 16 209 Nitrite-oxidizing bacteria, 5 stiptate forms of, 205 isolation and cultivation of, 16

362

SUBJECT INDEX

Nitrobacter, isolation and cultivation of, 16 media and methods for, 30-31 Nitrobacteriaceae,4,5,16-17,29-32 Nitrogen, sources for blue-green algae, 62 Nitrogen cycle, 26 Nitrogen-fixing algae, 54,56,62,63 Nitrosorno~ssp., 4 association with Sorangium sp., 204 isolation and cultivation of, 12-16 media for, 28-29, 30 Nocardia, phage for, 316 Non-sulphur-purple bacteria, see under Purple Nostoc sp., growth stimulation by Caulobacter, 56-57 medium for, 288, 292 N . muscorum, heterotrophic growth of, 63 light intensity for, 73 “morphogenetic substance” for, 53, 56 nitrogen source for, 62 pH and growth of, 60-61 Nutrient gelatin medium, 112 0 “Obligate” psychrophiles, 162 Ochrobium, 12, 15 isolation and cultivation of, 24-25 0. tectum, 15 Ochromunas, media for, 287, 289, 293 0. a’unica, 290 0.malhamensis, 287 “Ochromonus” medium, for algae, 293, 299 Oil baths, for thermophile culture, 166 Omasum, 134 Omnivores, bacteria like those of rumen in, 133-134 Oral flora, anaerobes of, 152 Oscillatoria sp., 54 Otitis media, of rodents, 220, 221 Ox,see also under Bovine, blood agar, 96 mycoplasmas from, 218 Oxidation-reduction indicators, 127

Oxidation-reduction potential, 117 Oxygen, aerobic organisms for removal of, 8788 anaerobes, motility and, 95 removal for growth of, 81-88 anaerobiosis and, 117-120 halophiles, for growth of, 172 lake water, in, 1-2, 16-17, 18, 21 media for anaerobes and, 120-1 26 motility of anaerobes and, 95 palladium catalyst for removal of, 8186 permissible concentration for anaerobes, 118 redox potential of, 118 removal of, 120-1 21,122-1 23 Oxygen electrodes, Mackereth, 3 Oxygen tolerant clostridia, 103 Oxyrrhis marina, media for, 291, 292

P Palladinized asbestos, for oxygen removal, 82 Palladium catalysis, oxygen removal by, 81-86 Parasitic myxobacteria, 196 Pastarella haemolytica, phage of, 316 Pathogens, halophiles as, 182 mycoplasmas as, 212, 216-221 Pectin media for rumen bacteria, 143 Pediastrum, medium for, 289 Pelagic diatoms, media for, 291, 293 naturalistic morphology of, 272 Pelargonate, enrichment of Rhodospirillum using, 58 Pelodictyon, 10, 11 isolation and cultivation of, 20-21 Pelonema, 12,14 isolation and cultivation of, 24-25 P. tenue, 14 Peloploca, 12, 14 isolation and cultivation of, 24-25 P . ferruginea, 14 Peloplocaceae, 12, 14

SUBJECT INDEX

Pelvic sepsis in man, mycoplasmas and, 217 Penicillin, algal purification using, 277, 278 mycoplasmal resistance to, 213, 227 Peptostreptococci, media for, 143, 144 Pepostreptococcus elsdenii, 143, 144, 147 Peranema, media for, 287 Peridinium, media for, 287, 288 Phacus, medium for, 290 Phaedactylum, media for, 290, 292 Phages, see Bacteriophages Phase-contrast microscopy, of mycoplasmas, 255-256 Phenethyl alcohol, as selective agent for anaerobes, 91 Phenosafranine, as redox indicator, 127 Phormidium persecinum, medium for, 292 Phosphorus, algal nutrition, and, 271, 298,301 Photobacterium phosphoreum, phage of, 316 Photosynthesis, cyclic, 270 Photosynthetic bacteria, elective media for, 58 families of, 57 freshwater lakes, in, 2, 3 green, see also Chlorobacteriaceae, 57 growth media for, 65-69, 70-72 illumination intensities for, 73-74 isolation and elective culture of, 57-59 physical conditions for growth of, 73-74 purple non-sulphur, see also Athio, rhodaceae, 57 purple sulphur, see also Thiorhodaceae, 57 stock culture maintenance of, 60 temperatures for growth of, 74 vitamin requirements of, 68 Photosynthetic non-sulphur bacteria, 3, 11,21-22, 37-39 Photosynthetic pigments, 7 4 7 5 Photosynthetic sulphur bacteria, 2,5,11 green, 20-21 isolation and cultivation of, 19-20 media for, 34-39 red and purple, 19-20

363

Photosynthetic thermophiles, 167 Phototactic movement, of blue-green algae, 54 Phototrophic bacteria, growth of, 53-76 Phototrophs, see also Blue-green algae and Photosynthetic bacteria, 53-77 Phycocyanin, 75 Phycoerythrin, 75 Phytoflagellates, media for, 287 Pigmentation, halophiles, of, 170, 171, 172, 173, 175, 177,178,180 myxobacteria, of, 205 Pigments, photosynthetic, 74-75 Pigs, mycoplasmal disease of, 216, 225 Pianktonic algae, media for, 287 Plant pathogenic bacteria, phages for, 316 Plaque size, factors affecting, 318 Plating techniques, algae, for, 274, 276-277 phage isolation, for, 318-320 Plectonenm borganum, phage of, 316 Pleuropneumonia-like organisms (PPLO), see also Mycoplasmas, 212 Pneumo-enteritis, as mycopiasmal disease, 218 Pneumonia, ox, of, 218 primary atypical, 216, 217 rodents, of, 220, 221 Podangium sp., isolation of, 193 P. graciltpes, 188, 189, 205 P . lichenincolum, 196 Polyangiaceae, characteristics of, 190 illustration of, 186, 187 purification of isolation of, 202-203 Polyangium sp., characteristics of, 190 fruiting-body morphology of, 205 isolation of, 192 maintenance of, 209 P . cellulosum, identity of, 194 P.fuscum, 195, 207 P . vitellinum, 188, 189 Polymyxin B, as selective agent for anaerobes, 90,91

364

SUBJECT INDEX

Polysaccharide slime, of myxobacteria, 202,203 Polytoma, media for, 270,287 Polytomella, media for, 288, 289, 290 Pope and Skerman media, 13,19,26,27 Porphyra, 291 Porphyridium, media for, 291, 292 Poteriochromonas, media for, 290 Poultry, mycoplasmal disease of, 221 PPLO, see Pleuropneumonia-like organisms also Mycoplasmas, 212 Primary atypical pneumonia, mycoplasmasand, 216,217 Prophage, 323, 324 Propionic acid, detection, 156 rumen fluid, in, 135 Proteolysis, by clostridia, 99 Proteolytic rumen bacteria, media for, 140 tests for, 145 Protoplasts, differentiation from mycoplasmas, 240 Prototheca, 270 media for, 289, 290 Protozoa, purification of, 278 Prymnesium parvum, media for, 291-292 Pseudocolonies, differentiation from mycoplasrnas, 243 PseUdomonas, RNA Phages of, 322, 323 Pseudomonas strain 101, as halophile, 180 P. aeruginosa, phages of, 321, 322 P . syringae, phage of, 316 Psychrop hiles, cultivation of, methods for, 163-164 definition of, 161-162 “facultative”. 162 incubators for, 163-164 isolation of, 162-164 “obligate”, 162 oceans, from, 162 stock culture maintenance of, 163 water samples, in, 163 Pure cultures, myxobacteria, of 202-203 strict anaerobes, of, 128-1 31 Purple bacteria, non-sulphur, see also Athiorhodaceae, 57

sulphur, see also Thiorhodaceae, 57-58 Pyrogallic acid, for oxygen absorption, 46,87 PYrogallol, for oxygen removal, 73, 86-87 Pyrobotrys, acetic acid requirement of, 270 medium for, 290

R Rat, m ~ c o ~ l a s mfrom, a s 220 RCM, see Reinforced Clostridial Agar Red sulphur bacteria, 19-20 Redox indicators, for media, 127,137 Redox potentials, anaerobiosis and, 117, 118 mud, of, 3 Reducing agents, for anaerobic media, 88-89, 122, 127, 132, 137 Reinforced Clostridial Agar, 153 Resazurin, as redox indicator in media, 127, 137, 138, 139, 140, 141, 142,144 Respiratory diseases, mycoplasmal, 216, 217,221,222 Rhabdochromatium, 9 R.fusiforme,9 roseurn, Rhabdomonas, 5,9, 19-20 Rhizobium spp., phage of, 316 Rhodomicrobium vanniellii, bacteriochlorophyll of, 74 enrichment of, 58 growth medium for, 66, 67 physical conditions for growth of, 73 stock culture maintenance of, 60 temperature for growth of, 74 Rhodopseudomonas, 1 1, 21 R. capsulata, 65, 68 R. gelatinosa, 65, 68 R. isachenkoi, 65, 68 R . palustris, enrichment of, 58 growth medium for, 65, 66, 67 vitamins for, 68 R . spheroides, growth medium for, 65,66, 67 light intensity and bacteriochlorophyll of, 74

365

SUBJECT INDEX

R. spheroides-cont. temperature for growth of, 74 vitamins for, 68 Rhodospirillum, 11,21,38 R . fulvum, enrichment of, 58 R . molischianum, enrichment of, 58 R . rubrum, aerobic growth conditions for, 74 continuous culture of, 75 growth medium for, 38, 65, 66, 67 vitamins for, 68 Rhodotheca, 5, 8, 19-20 R. nuda, 8 R . conspicua, 8 “Rhodovibrio”, 38 Ribonuclease, test for RNA phages, 322 RNA phages, 314, 319, 322-323, 324 Robertson’s cooked-meat broth, for anaerobes, 101,103 Rodents, mycoplasmal pneumonia of, 221 Roll-tube method, for anaerobes, Gram-negative non-sporing, 152 strict, 117-132 Rotifera, purification of, 278 Rumen, bacteria of, see Rumen bacteria, 133149 fatty acids in, 135 Gram-negative anaerobes from, 152 Rumen bacteria, amylolytic, 140 biochemical tests on, 145 carbohydrate fermentation by, 137, 143,145 carbon dioxide for, 137 cellulolytic, 139-140 continuous culture of, 147-148 culture maintenance of, 147 defined media for, 146 dilution techniques for, 136 glycerol media for, 143 growth factors for, 135, 146 growth media for, 135-146 habitat-simulating media for, 135138 identification media for, 144-146 isolation of, 135-144 Iipolytic, 141 media for, 135-144, 144-146

methanogenic, 142 morphology of, 144-145 nitrogen requirements of, 136-137 non habitat-simulating media for, 143-144 pectin-containing media for, 143 proteolytic, 140-141 saponin-containing media for, 143 selective counting of, 138 solid media for, 139 stock culture maintenance of, 147 viable counting of, 136-138 xylan-containing media for, 143 Rumen fluid, 135,136,138 Ruminant saliva, 134, 137 Ruminants, digestive system of, 133 Rumination, 134 Ruminococci, 13 7 Ruminococcus a l h s , continuous culture of, 148

S Saccharolytic species, of anaerobes, 99 Salmonella, sp., phage of, 323 S. typhimurium, phage from 323 Salpingitis, as mycoplasmal disease, 218 Salt, see also Sodium chloride, halophiles and, 169, 171, 177, 178, 179-1 80 solar, 171, 177 Salt-tolerant organisms, 170, 180 Sample collection, autotrophs, for, 3 psychrophiles, for, 162 thermophiles, 167 Samples, bacteriological, 3 Saponin, 143 Saprospira sp., cultivation of, 64 Sarcina Zitoralis, temperature and growth of, 172 Scenedesmus, media for, 287, 289 Schatz and Bovell’s medium, 42 Serology,of anaerobes, 102 Serratia marcescens, oxygen removal by, 87 Serum agar, see u n d o Human Shake cultures, of anaerobes, 88 Shaking algal cultures, 282-283

366

SUBJECT INDEX

Sheep, mycoplasmal aglactia of, 212, 221, 231 Shepard’s A2 medium, 228 Shipping fever, as mycoplasma disease, 218 Siderobacter, 12,15 isolation and cultivation of, 24-25 S. duplex, 15 S. lineare, 15 siderocapsa, 12, 15 isolation and cultivation of, 24-25 S. c o r m t a , 15 S. geminata, 15 S. major, 15 S. Treubii, 15 Siderocapsaceae, 12, 15, 24-25 Siderocystis (Sideromonas) confervarum, 15 Sideroderma (Ferribacterium) dubium, 15 Sideromonas, 12, 15 isolation and cultivation of, 24-25 S.confmarum, 15 Silica gel media, for autotrophs, 28, 29, 30 Silicic acid sol media, for autotrophs, 29, 30 Silicon, algal nutrition and, 271, 301 “Singh’s Aerobacter circles” for myxobacteria, 192,195-196 Skeletonema costatum, 292, 293 Skerman’s medium, 16 Skerman’s method for gas mixing, 43 Skim milk medium, for anaerobes, 158 Skinner and Walker’s medium, 31 Slime, myxobacterial, 202, 203, 204 Sodium azide, as selective agent, 90 Sodium chloride, see also Salt, halophile requirement for, 171 Soil, myxobacteria from, 192,194 Soil media, for algae, 298, 299 Solar salt, halophiles from, 171, 177 Sorangiaceae, 186, 187, 189, 203 Swangium spp., cellulose media for, 198-199 fruiting body morphology of, 205 isolation of, 192, 193, 194 maintenance of, 207,208,209 Nitrosomonas association with, 204 pigmentation changes in, 205 s.cehlosum, 186, 187, 189

Sorbic acid, as selective agent, 90 Soriano and Walker’s medium, 16, 31 Sphaerellaceae, medium for, 290 Sphaerophorus spp., biochemical tests for, 154 dyes, resistance to, 158 isolation medium for, 159 media for, 153-154, 159 methods for, 152-160 S. necrophorus, 152 Sphaerotilusspp., 12,13 isolation and cultivation of, 24-25,47, 48 S. natans, 13, 15,47,48 Sphaerotilus-Leptothrix group, isolation of, 25 Spheroplasts, differentiation of mycoplasmas from, 240 Spirillaceae, 11,21-22,394 Spirochaeta rosea, phage of, 316 Spore-forming bacteria, aerobic, 96 anaerobic, 80-1 15 Spores, clostridial, heat-resistant, 103, 104 morphology of, 94 Sporocytophaga sp., exclusion from Myxobacteriales, 187 “Squire” mycoplasma, 218 Stains, immuno-fluorescent, 94-95 Muir’s, for capsule staining, 94 Stainer plates, for myxobacteria, f 92, 194-195,198-199 Starch gel electrophoresis, for cell protein analysis, 251 Starkey’s media, 34, 39 Steam pressure sterilization, 127-128 Stelangium sp., characteristics of, 189 isolation of, 193 Stephanopyxis, medium for, 291 Sterility-testing, of algal cultures, 284 Sterilization, algal media, of, 283-284 anaerobic media, of, 127-1 28 Sterols, see also under specific names, mycoplasmal growth and, 212, 213, 235,240,250 Stichococw, media for, 290,292

367

SUBJECT INDEX

Stigioclonium, media for, 287 Stock cultures, see also under Maintenance and names of spec$c micro-organisms, anaerobes, of, 92, 131 blue-green algae, of, 59-60 halophiles, of, 179 mycoplasmas, of, 258-260 Myxophyceae, of, 59-60 photosynthetic bacteria, of, 60 psychrophiles, of, 163 rumen bacteria, of, 147 Stratified freshwater lakes, autotrophs in, 1-26 Streptococci, 143, 147 Streptococcus spp., phage of, 316 S.bovis, from rumen contents, 143 Streptomyces venezuelae, phage of, 316 Streptomycin, 91 algae, for purification of, 277,278 anaerobes, as selective agent for, 91,

media and methods for, 32-34 Sulphur cycle, 28 Surface culture of anaerobes, 102-103 Swabs, mycoplasmas from, 223, 224 Swarming anaerobes, 95, 103 Symbiodinium, medium for, 292 Synchronization, of algal cultures, 307 Synechococcus lividus, growth of, 63, 73 Synovitis, mycoplasmas and, 221 Synura, media for, 288,290

T

T-strain mycoplasmas, bovine strain of, 231 colonies of, 214, 215, 221 erythromycin inhibition of, 213 isolation media for, 228, 229, 231 maintenance of cultures of, 258, 259260 man, from, 217 158-159 ox, from, 218 Strict anaerobes, seealso under Anaerobes specimens and transport of, 224 and names of specific microthallium acetate inhibition of, 228 organisms, titres in liquid media, 246 colonies of, 129 Temperate phages, 322, 324, 326-327 cultivation of, 117-132 Temperature blocks, for study of dilution of, 130 thermophiles, 165 media for, 120-1 26, 126-1 27 Temperature control, media sterilization for, 127-128 algal cultures, of, 280-282 oxygen-free media for, 120-1 26 thermophiles, for study of,164-167 picking colonies of, 129-130 Temperature gradients, in incubators, pure cultures of, 128-131 163 redox potentials for, 118 Tendo-synovitis, mycoplasmas and, 221 roll-tube cultivation of, 117-132 Tetanus, bacteriological diagnosis of, storage of cultures of, 131 104 Succinic acid, in rumen fluid, 135 Tetracyclines, mycoplasmal inhibition Sucrose, clostridial fermentation of, 99, by, 213 110 Thallous acetate medium, for mycoSulphadiazine, as selective agent, 91 plasmas, 226 Sulphate-reducing autotrophs, 2,811 Thermocline, isolation and cultivation of, 21-22 bacteria in, 2, 3, 21 media for, 39-40 definition of, 1,2-3 pure cultures of, 3 9 4 0 Thermop hiles, spore-forming, 22 agar plates for, 166-1 67 Sulphur-oxidizing bacteria, 5 cold shock of, 167 enrichment methods for, 17, 32 contamination of, 167 isolation and cultivation of, 16-1 8 definitions for, 161-1 62 18-1 9 fermenters for, 166

368

SUBJECT INDEX

Thermophiles-cont. heat exchangers for study of, 165-1 66 isolation of, 164 photosynthetic, 167 sample collection of, 167 Thermophilic algae, 54,63,73,273

Thiamine, algal requirements for, 272 Athiorhodaceae, for, 68 Thiobacilli, anaerobic, 19 enrichment cultures of, 18 Thiobacillus, enrichment cultures of, 18,19 isolation and cultivation of, 18-19, 33-34 T . denitri’cans, 33 T . thio-o3cidans, 33 T. thioparus, 34 Thiobacteriaceae, 5 , 7, 18-19 Thiocapsa, 5 , 8 , 19-20 T . roseopersicina, 8 Thiocyanate, thiobacilli oxidizing, 19 Thiocystis, 5 , 8, 19 T. violacea, 8 Thiodictyon, 5,8,19-20 T. elegans, 8 Thioglycolate, as reducing agent for media, 127 Thioglycollic acid, as reducing agent for anaerobiosis,89 Thiopedia, 5 , 8, 19-20 T . rosea, 8 Thioploca, 5 , 6, 16-18 T. ingrica, 6 Thiopolycoccus, 5 , 8, 19-20 T . ruben, 8 Thiorhodaceae, elective medium for, 58 growth media for, 68-69 illustrations of, 8, 9 isolation and culture of, 19-20, 57-59 light intensities for, 74 medium for, 37 physical conditions for growth of, 73-74 representatives of, 5 stock culture maintenance, 60 Thiospira, 5,7, 18-19 T . agilis, 7

T. dextogyra, 7 T. tenuis, 7 T . Winogradsky, 7 Thiospirillum, 5 , 9, 19-20 T .jenense, 9, 35, 36 T . Rosenbergiy, 9 Thiothece, 5,8,19 T . gelatinosa, 8 Thiothrix, 5,6, 16, 18 T . annulata, 6 T . nivea, 6 T. tenuis,6 Thiovulurn, 5,7,18-I9 T. majus, 7 T . minus, 7 T . Mulleri, 7 Threonine, utilization by anaerobes, 155 Threonine deaminase, test for anaerobes, 154,155 Tissue cultures, mycoplasmal contamination of, 236-237 Tolypothrix, medium for, 288 T. tenuis, 5 3 heterotrophic growth of, 63-64 illumination and pigments of, 75 temperature for growth of, 73 Total cell count, mycoplasmal growth measurement by, 249 Toxigenic clostridia, key to, 109 Toxigenicity, demonstration of, 102 Toxins, of Cl. welchii, 96, 97 Trace-elements, for blue-green algae, 64 Trace metals, for algae, 272, 273, 296298 Trace nutrients, algae and, 271, 272 Tree bark, myxobacteria from, 192-193 Tube-roller, mechanical, 130 Tungsten lamps, for algal illumination, 280 Turbidity, growth of mycoplasma by measurement of, 245-246 Turbidostat culture, of algae, 303, 304

U Ubiquinone, in Rhodomicrobium sp., 65 Ultraviolet light, algal culture purification using, 56 phase host lysis by, 324, 327 Ulva, media for, 291, 292

3 69

SUBJECT INDEX

Uni-algal cultures, 56, 273 micromanipulative procedures for, 54 purification of, 54-57 Urea, mycoplasmal metabolism of, 221,

229,248

W Waksman and Starkey’s medium, 33 Water-bath shakers, 165, 168 Water-baths, for therrnophile culture,

165-166,168

V Van Niel’s medium, 20, 37 Vancomycin, resistance of Bacteroides to, 159 Veillonella alcalescens, 137 V . egasogenes, 137 Veillonellas, in rumen, 143 Versine, for glassware treatment, 285 Viable cell count, mycoplasmal growth measurement, for, 248-249 rumen bacteria, of, 136-1 38 Vibrio costicolus, 181 V.parahamolyticus, 182 Viscap sealing cap, 20 Vitamin B12, algae, for, 272 Thiorhodaceae, for, 69 Vitamins, see also under individual names, algae, for, 296 photosynthetic bacteria, for, 68 Vitreoscilla sp., cultivation of, 64 VL blood agar, for Gram-negative anaerobes, 153 Volvocales acetate algae, 270

Water sampling, from lakes, 3 Winogradsky columns, 3, 19, 20, 21, 59 Wolossynskia, 290 Wounds, clostridia from, 104-107 Wright’s capsule, for anaerobic jars, 81,

82,84

X Xanthomonas spp., phages of, 316, 321 X. malvacearum, phage of, 316 Xanthaphyceae, sulphur-utilizing, 270 Xanthophyta, medium for, 288 Xylan, media for rumen bacteria, 143

Y Yeast extract medium, 226

Z

Zooxanthellae,291,292

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