C H A P T E R X I 1

Isolation, Growth and Preservation of Bacteriophages

EVE BILLING*

Department of Microbiology, University of Reading, England

I. Introduction . 11. Isolation

A. Growth conditions . B. Platingmethods . C. D. Isolation from natural sources . E. F. Purification .

A. Plate method B. Broth method C. Temperate phages .

Separation and inactivation of bacteria

Isolation from lysogenic and carrier strains

111. Preparation of High-titre Stocks .

IV. Preservation . References .

. 315

. 317

. 317

. 318

. 320

. 321

. 323

. 324

. 325

. 325

. 326

. 326

. 328

. 328

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 phage- host systems. For the most part they involve simple operations and require no complex apparatus or equipment. Those who seek more specific informa- tion 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.

316 EVE BILLING

TABLE I Isolation methods for bacteriophages

Bacteriophage Reference

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

Rhizobium spp. Spirochaeta rosea Haemolytic Streptococcus Streptomyces venezuelae Xanthomanas malvacearum

phytopathogens

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 (1 960) Schneider et al. (1964)

Cross & Garrett (1963) Schwinghamer (1965) Lewin (1 960) Kjems (1960) Kolstad & Bradley (1964) Hayward (1 964)

Okabe and Goto (1963) describe sources of phages for other plant patho- genic bacteria and further methods for their isolation. For information on selection of phages for typing purposes, see Parker, this Series, Vol. 7.

In 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

XII. BACTERIOPHAGES 317

phages may often provc easy, specific enrichmcnt for phages with a particular morphology, nuclcic acid content, or host rangc is scldom a practical proposi- tion. The isolation of temperate phages depends on the availability of suitable lysogenic and indicator strains and presents somewhat different problems (see Section 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

318 EVE BILLING

l.Oyo is sufficient; for soft agar 0.6-0*7% of agar is the usual recommen- dation. 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 neces- sarily 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 condi- tions. 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

I XII. BACTERIOPHAGES 319

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 dilu- tions 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 tempera- tures 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.

320 EVE BILLING

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. The 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 impor- tant 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 (0-45 pm) 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 suspen- sion 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.

XIX. BACTERIOPHAGES 321

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 in- itially 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 1 ml 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.

322 EVE BILLING

Phages which form large plaques can present certain difficulties in quantita- tive 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 propa- gated (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 consider- able 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 Entero- bacteriaceae 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

XII. BACTERIOPHAGES 323

at 30,000 rpm in a Spinco Model L centrifuge. They found RNA phages in four drop 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 appropriate type. 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 agar- layer 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 concern- ing 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

324 EVE BILLING

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 influenced by the conditions of growth.

In 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 allow the 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-

XII. BACTERIOPHAGES 325

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 phage- host 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 har- vesting, 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 treat- ment may be applied earlier either by adding to the suspension or by inver- ting dishes (glass not plastic) for 15 min over 0.5 ml of chloroform in the

326 EVE BILLING

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 lyso- genized instead of lysed. By altering the growth conditions, it may be pos- sible to reduce the frequency of lysogenization, e.g., by raising the tempera- ture 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).

XII. BACTERIOPHAGES 327

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 phosphate- buffered 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 phage- host 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

The stability of a phage will depend on the suspending fluid. In 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 suspen- sions 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 screw- capped 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 some- times 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. (1 964). 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. (1 966). J. Bact., 91, 1305-1 3 13. Clark, W. A. (1962). Appl. Mictobiol., 10, 466-471.

XII. BACTERIOPHAGES 329

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.,

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,

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.,

Jacob, F., and Wollman, E. L. (1959). In “The Viruses” (Burnet, F. M. and Stanley,

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.,

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

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.

45,329-338.

H., Eds). Academic Press, New York and London.

12,329-370.

W. M., Eds). Academic Press, New York and London.

216,343-346.

Bacteriology”. Cambridge University Press, London and New York.

Zinder, N. D., Valentine, R. C., Roger, M.,.and Stoeckenius, W. (1963). Virology, 20,638-640.