News

News
 
Prevention of mites in cultures by Webmaster Online protocols 2016-02-06 18:32:27
 

A.H.S. Onions

Formerly of: CAB International Mycological Institute,
Culture Collection & Industrial Service Division,
Ferry Lane, KEW,Surrey TW9 3AF, United Kingdom

World Federation for Culture Collections
Technical information sheet No 1

Published by: UNESCO/WFCC Education Committee 1989

Introduction

Mites can be a problem in fungal culture collections. These small animals of the genera Tyroglyphus and Tarsonemus occur naturally in soil and almost any organic material and fungi are particularly susceptible to their attack. They can be seen by the naked eye as tiny white dots almost at the limit of vision, often about 0.25 mm in length. As they tend to bury themselves in the mycelium, they multiply rapidly and the first indication may be the deteriorated (moth-eaten) look of the cultures, with wandering trails of mini-colonies on uninoculated agar.

Mites can be brought into the laboratory on fresh plant material, decaying mouldy products, shoes, bodies of insects or even cultures from other laboratories. They thrive in moist warm conditions, so may often first appear in Europe in early summer or in new laboratories with new equipment, damp walls and new wood. Not only do they eat the cultures but they also carry fungal spores and bacteria on their hairy bodies as they move from one culture to another.

Prevention

Prevention is preferable to having to control an outbreak. This can be summed up as general hygiene and screening of all cultures and material coming into the laboratory and destroying or isolating ail infested material. Even so, separate handling, storage and quarantine of fresh material is desirable, as slightly infested cultures can develop heavy infestation if undetected at first examination. If it is necessary to handle infested material, the culture collection should be well isolated.

Control

Different workers have varying views on control. Some who are handling a quick turnover of infected material may not even regard a few mites as serious, but in a culture collection they spell disaster. A combination of prevention and action seems preferable. These can best be classed in several categories: (1) Hygiene, (2) Fumigation, (3) Mechanical and chemical barriers and (4) Protected storage.

1. Hygiene

Hygiene coupled with quarantine is perhaps the best protection.

1.1 All work surfaces must be kept clean.

1.2 Cultures should be protected from airborne contamination.

1.3 Mites can be carried on workers' hands and clothing.

1.4 Cramped laboratory conditions and crowded arrangement of cultures increase the risk of infestation.

1.5 Work surfaces and benches should be washed regularly with acaricide. The acaricide is left for sufficient time to act (overnight) and washed off, preferably with alcohol. As some acaricides are toxic to man, protective gloves should be worn.

The acaricide used at the CMI is Actellic (ICI, Agrochemicals, PLC). Other acaricides available for agricultural and grain storage purposes are Murfit, Reldan and Dursban (Murphy Chemicals Ltd) and Satisfar (Sandoz Ltd

As mites appear to become resistant, the acaricide should be changed from time to time.

1.6 Infected cultures should be removed immediately and sterilized if possible. All cultures in the immediate area should be checked and isolated.

2. Fumigation

Cultures may be stored in cupboards or boxes with acaricides either as preventative or short-term treatment. Camphor and paradichlorbenzene (PDB) have been used for this, but are now regarded as toxic. PDB also has some effect on fungal growth. Drops of Kelthane and Crypo on culture plugs (Smith, 1967) were effective. Current safety practice would suggest that fumigation is no longer desirable.

3. Mechanical and chemical barriers

Many physical methods of prevention of infestation and spread have been tried.

3.1 Cultures are placed on a platform or tray surrounded by water, oil, petroleum jelly or other sticky material. Handling of cultures becomes unpleasant and protection is only from crawling mites.

3.2 Culture bottles or plates may be sealed, but it is necessary to allow growing cultures free respiration, so a means of sealing which is permeable to air is desirable.

Snyder and Hansen (1946) sealed bottles below screw caps or above the cotton wool plugs (well pushed down) with sterile cigarette papers using copper sulphate glue (20 g gelatin, 2 g copper sulphate, 100 ml water). The pores of the paper allowed respiration but prevented movement of mites, thus protecting clean cultures and isolating infested ones. Care is necessary to ensure the seal is effective.

Smith (1971) recommends the use of disposable plastic bottles with plastic caps which, when screwed down, still allow respiration but exclude mites.

Smith (1978) described a screw-lid closure with a hole sealed with Metricel.

Sealing Petri dishes and bottles with various modern plastic tapes often reduces spread but, by means of cracks or wrinkles, mites can eventually penetrate cultures stored for a long time.

Tight cotton wool plugs present a considerable barrier but are not completely effective, though a mite that has passed through cotton wool is often much cleaner. Some workers treat plugs with mercuric chloride solution. This kills the mites but is poisonous and dangerous to handle even if a red dye is included to indicate its presence. It is also toxic to fungi.

4. Protected storage

Many methods used for long-term storage of cultures in culture collections prevent infestation.

4.1 Mites do not infest cultures stored under mineral oil.

4.2 Cold storage at 4-8C greatly reduces movement of mites but does not kill them, so they continue to multiply when the cultures are removed from the refrigerator.

4.3 Deep-freeze storage at approximately -20C usually kills any mites present. Storage in a deep freeze for three or four days can be used prior to cleaning to treat infested material that is too valuable to discard.

4.4 Freeze-dried ampoules are sealed and totally protected.

4.5 Storage at ultra-low temperatures, for example in liquid nitrogen, gives total protection.

4.6 Cultures stored in silica gel are in vials or bottles with screwed-down caps, so are totally sealed.

REFERENCES

Smith, R.S. (1971). Maintenance of fungal cultures in pre-sterilized disposable screw-cap plastic tubes. Mycologia 63, 1218-1221.

Smith, R.S. (1978). A new lid closure for fungal culture vessels giving complete protection against mite infestation and microbiological contamination. Mycologia 70, 499-508.

Snyder, W.C. & Hansen, H.N. (1946). Control of mites by cigarette paper barriers. Mycologia 38, 455-562.

     
 
Liquid-drying of micro-organism using a simple apparatus by Webmaster Online protocols 2016-02-06 18:32:45
 

Liquid-drying of micro-organism using a simple apparatus

Khursheed A. Malik, Ph.D.

DSM-Deutsche Sammlung von MIkroorganismen und Zellkulturen GmbH,
Mascheroder Weg 1B, D-3300 Braunschweig, Federal Republic of Germany

 

World Federation for Culture Collections
Technical information sheet No 8

Published by: UNESCO/WFCC - Education Committee 1990

Introduction

Liquid-drying (L-Drying) involves vacuum-drying of samples from the liquid state without freezing. It is one of the sophisticated techniques used for the long-term preservation of microorganisms.

Several microorganisms, which are sensitive to freezing or freeze- drying, can successfully be preserved by liquid drying. Liquid-drying has several advantages over freeze-drying and has been effectively used for preserving large collections of fragile microorganisms in various culture collections. However, specialized equipment is required for L-drying. Recently a simple and effective liquid-drying method has been described for the successful preservation of sensitive microorganisms (including various anaerobes) which fail to survive freezing or freeze-drying (Malik, 1990).

Based on this process a liquid-drying method using a simple apparatus is described. The method is simple and the equipment described here can be easily constructed in most laboratories.

Material and Methods

Preparation of thin discs of carrier material

Ampoules of neutral glass (45 x 10 mm) are filled with 0,1 ml of 20 % (w/v) skim milk (Bacto, Difco 0032) containing 10% (w/v) neutral activated charcoal and one of the effective protective agents such as 5 % (w/v) meso-inositol, 5% (w/v) glutamate, 5 % (w/v) raffinose or 10% (w/v) honey. The activated charcoal used is of medicinal grade ( available from Caelo, 4010 Hilden, FRG) but any other bacteriological activated charcoal of comparable quality can also be used. The ampoules are loosely plugged with non-absorbent cotton wool and sterilized at 115 ’C for 13 minutes. These are frozen at about -30 ’C for a few hours and then freeze-dried in bulk for about 6 hours using a standard freezedrying technique (Malik,1988 or for freeze-drying with a simple apparatus see TIS No.7. Malik, 1990).

Preparation of protective agents

Solutions of most effective protective agents like meso-inositol (5 % w/v), honey (10 % w/v), sodium glutamate (5 % w/v), raffinose (5 % w/v) are prepared in distilled water, filter sterilized and stored at 4’ C. For oxygen sensitive microorganisms 10% (w/v) neutral activated charcoal with 10% (w/v) meso-inositol is prepared in a screw cap bottle (Malik, 1990). The solution is boiled, bubbled with nitrogen gas and the bottle is closed tightly and autoclaved at 115 ’C for 13 minutes.

Preparation of cell suspension for L-dryinq

A thick cell suspension (at least 10(8) cells per ml) is prepared in an appropriate protective medium. In the case of liquid cultures, the cells are harvested by aseptic centrifugation for 30 minutes at 4000 xg in screw-cap bottles and the pellet is suspended in a protective medium to yield a heavy cell suspension. The thick cell suspensions under anaerobic conditions are obtained as described previously [ see TIS No.4, Malik,1989 ].

Filling of ampoules with cell suspensions

The ready ampoules (containing a thin disc of carrier material) are equilibrated at 2†-25’ C for few minutes. To each ampoule about 0,025 ml (1 drop with a Pasteur pipette) of cell suspension are added aseptically onto the thin-disc with care so as not to touch the sides of the ampoules. The ampoules are quickly placed in metallic lids and transferred into a metallic jar maintained at 20 ’C in a water bath (see Fig.1 A).

After 20 to 30 minutes of equilibration these are subjected to drying under vacuum. For strains very sensitive to oxygen, the ready ampoules are first placed for few hours in anaerobic Bio-bags (Type A, Marion Scientific Corporation, Kansas City, Ml) or Anaerocult P bags (E. Merck, Postfach 4119, D-6100 Darmstadt). These are carefully taken out one by one and after the addition of reduced cell suspension the ampoules are immediately transferred again to such fresh anaerobic bags, sealed and kept for about one hour at room temperature for equilibration and removal of oxygen. Thereafter, these are subjected to drying under vacuum.

The liquid-dryinq procedure

The outline of the L-drying procedure and the major steps involved are shown in Fig.1. For the double vial preparation, liquid-drying is done in two stages involving primary-drying and secondary-drying.

The primary drying is achieved in two stages. The cold trap is chilled to about -35’C and is connected to the vacuum pump and the metallic evacuation jar (maintained at 20’C in a water bath see Fig.1 A). A vacuum controller is attached between the cold trap and the vacuum pump to control the vacuum.

The cold trap tube is a U-shaped thick glass or preferably a metal tube of about 2-3 cm diam and about 30 cm length. It is filled with blue/dry silica gel and is placed in a metallic beaker as shown in Fig.1 A. It is chilled to about -35’C. A cooling mixture of ethylene glycol: water (1: 1) is placed for few hours in a deep-freezer and is cooled down to about -30’C (preferably to -40’C). This is poured into the cold trap to give a maximum depth. The cooling mixture is also sealed in deep-freeze plastic bags and is cooled down or frozen in a deep-freezer or over liquid- nitrogen. This super cooled or frozen coolant in the bags is added to the cold trap and the bags are changed periodically (preferably after every 20 25 min) throughout the drying process in order to maintain the temperature at a minimum level. Commercially available anti-freeze liquids used in car radiators as coolants are also satisfactory as an alternative for the low-temperature bath and cold trap. A double jacketed straight tube (exterior about 6x 30 cm and interior about 3x25 cm with outlet and inlet tubes of about 2x5 cm, as shown in Fig.1A) can also be used as a cold trap. If available, dry ice is used for freezing the ampoules and is added to the cold trap periodically throughout the experiment.

The vacuum is switched on and the temperature of the water bath is maintained at 20’C. First step drying (Drying-l) is continued for about 30 minutes at about 5-10 mbar after which the vacuum controller is readjusted for 1,0 to 0,1 mbar vacuum and second step drying (Drying-ll) is conducted for about one hour.

At the end, the vacuum is replaced with nitrogen gas (especially for strict anaerobes and for the ampules which are not to be sealed under vacuum). For secondary drying and sealing, the ampoules are transferred to soft glass tubes (130 x 15 mm) containing silica gel and cotton plugs. The outer tubes (outer vials) are then constricted by hand or by using Edward Ampoule Constrictor), attached to the manifold and mounted on an evacuation jar. This operation is illustrated in Fig.1 B. The vacuum is switched on and secondary drying is conducted for 1-2 hours (at 0,1 to 0,001 mbar). The pink silica gel in the outer tubes will turn again to blue at this stage.

The constricted outer tubes are carefully sealed, by hand or by using a Flaminaire blow torch, one by one maintaining vacuum (Fig.1 C). For more details see TIS No.7.

Revival of cultures from liquid-dried ampoules

Improvement in viability can be achieved through the use of different culture media. Thus during reactivation of preserved microorganisms it is recommended to use the most favourable media and growth conditions. During several years of experience I have observed that in the case of sensitive microorganisms when the preserved (dried or cryogenically stored) cultures are revived, the counts on agar media are usually lower than in liquid media and similarly agar media of higher surface tension (such as nutrient agar) usually results in lower viable counts than mineral media of relatively lower surface tension.

The contents of the liquid-dried ampoule are thus dissolved in sterile (prereduced in the case of anaerobes) liquid growth media and added or injected (for anaerobes) into 15-20 ml of growth medium. Freshly inoculated phototrophic cultures are placed for a few hours in the dark at appropriate incubation temperatures and later under normal growth conditions in the light. The dried cultures are incubated at a relatively lower temperature than the optimum growth temperature. A few cultures may exhibit a prolonged lag period and thus are incubated for relatively longer periods. Normal growth usually appears after a second transfer into fresh medium. The use of activated charcoal in suspending media for reactivation is recommended due to its various advantages (Malik.1990).

Estimation of viability and stability

Survival recoveries are checked before L-drying, immediately after L-drying and after storage. For the estimation of viability counts serial dilutions are prepared in appropriate liquid media. From each serial dilution 0,1 ml volumes are plated on agar media plates. The number of colonies are counted from the plates and average colony forming units per sample are calculated. The revived cultures are also observed for mutation, change in colony morphology or other characters. For cultures which were difficult to grow in or on agar, only liquid dilutions series are made. In such cases the number of cells is determined using the most probable number method (MPN).

Long-term storage

Stability of L-dried cultures during storage is very important. A high level of residual moisture content or exposure to oxygen have detrimental effects on the dried product. Liquid-dried material is hygroscopic and its exposure to moisture during storage can destabilise the product.

The higher the storage temperature, the faster a product will degrade. Thus, the storage of L-dried cultures at lower temperatures will extend their shelf life. The unsealed L-dried ampoules can safely be stored for several years at about -30’C. It has been observed by the author that similar viability counts were obtained after 4-5 years of storage when unsealed freeze-dried culture were maintained at -30’C as compared to the freeze-dried cultures which were sealed under vacuum and were stored at 9’C (Malik, 1976).

Selected references

Annear, D.l. (1956). The preservation of bacteria by drying in peptone plugs. Journal of Hygiene 54: 487.

Banno, T. and Sakane, T. (1979). Viability of various bacteria after L-drying. IFO Res. Comm. 9, 3545.

Hieda, K. (1981). Induction of genetic changes in Saccharomyces cerevisiae by partial drying in air of constant relative humidity and by UV. Mutation Res. 84: 17- 27

Malik, K.A. (1976). Preservation of Knallgas bacteria. In Proceedings of Fifth International Fermentation Symposium (H.Dellway, Ed.) p. 180. Westkreuz Druckerei and Veriag, Bonn and Berlin.

Malik, K.A. (1985) . Modern Methods of Gene Conservation. A Laboratory Manual. PASTIC Press Pakistan Science and Technology Information Centre, Islamabad, Pakistan.

Malik, K.A. (1987). The role of culture collections in the stability and preservation of microorganisms (J.Amen and P. Tesson, Eds.) pp 118-150. Societe Francaise de Microbiologie. Paris

Malik, K.A. (1988). A new freeze-drying method for the preservation of nitrogen-fixing and other fragile bacteria. J. Microbiol. Methods 8: 259-271

Malik, K.A. (1989). Cryopreservation of bacteria with special reference to anaerobes. Publication No.4. UNESCO/WFCC Technical Information Sheets (TIS). DSM, Braunschweig

Malik, K.A. (1990). Freeze-drying of microorganisms using a simple apparatus. Publication No.7. UNESCO/WFCC Technical Information Sheets (TIS). DSM, Braunschweig

Malik, K.A. (1990). Use of activated charcoal for the preservation of anaerobic phototrophic and other sensitive bacteria by freeze-drying. J. Microbiol Methods

Malik, K.A. (1990). A simplified liquid-drying method for the preservation of microorganisms sensitive to freezing and freeze-drying. J. Microbiol Methods

Sakane, T, I. Banno and T. Iijima (1983). Compounds protecting L-dried cultures from mutation. IFO Res. Comm. 11: 14-24

     
   

J. A. Stalpers, A. De Hoog, And IJ. Vlug

Centraalbureau voor Schimmelcultures

Mycologia
79(1), 1987, pp. 82-89.

Copyright 1987, by The New York Botanical Garden, Bronx, NY 10458

ABSTRACT

A method for cryopreservation of fungi in straws is described in detail; for large scale application special equipment had to be developed for which technical diagrams are provided.

Key Words: preservation. fungi.

The storage of fungi in liquid nitrogen has generally been recognized as the most reliable method for long term preservation. For culture collections which are recognized as patent deposits it has become a necessity because of the obligations for the maintenance of patented strains under the Budapest Treaty.

The disadvantages of cryopreservation including the risks connected with the necessity of a constant supply of liquid nitrogen (LN), the expensive, space-consuming and sometimes dangerous use of glass vials and the lack of good alarm and automatic filling systems, have been strongly reduced by recent developments. The use of polypropylene straws instead of glass vials (Elliott, 1976; Elliott and Challen, 1979) is both cheap (for the whole CBS collection 360,000 straws (12-fold) are necessary which cost about 140 US $) and safe (when a glass vial is improperly sealed and LN has leaked in, thawing may cause an explosion resulting in flying glass, but a straw only splits) and the latest version of the Union Carbide alarm and filling system is relatively reliable.

Because of the small size of the straws the consumption of LN drops drastically when compared with an equal number of glass vials. The estimated annual LN consumption for one vivostat (filled) is about 5000 liters; the whole CBS collection, which can be stored 8-fold in three vivostats, would require 15,000 liters. Of course the consumption depends a great deal on how many times the vivostats are opened and how much is added.

As all the equipment for handling the straws had to be developed, it is considered useful to describe the method in detail and to provide technical diagrams.

Preparation of straws

After testing many specimens of several trade marks of straws with diameters of 3 and 4 mm, Elliott and Challen's (1979) conclusion was confirmed: the straws with the best overall score with respect to resistance to autoclaving, heat-sealing and storage in LN were 4 mm polypropylene Sweetheart Winfield drinking straws, produced by Sweetheart International Ltd. (straw code 0175050). Straws with diam of 3 mm with the same qualities were not found and the 3 mm size proved too small for easy handling when agar plugs are used. However, the size may be useful when broth cultures are used and thus the equipment was designed for both 3 and 4 mm straws.

Straws of about 200 mm long are fixed in a mold (52 x 32 x 47 mm) and cut into pieces of 47 mm using an ordinary cutting device (Graef, type EH-170 T), from which a part of the protective shield had to be removed (FIG. 1, arrow). For each strain 12 straws are prepared. The straw holder (FIG. 3, DIAGRAM 1, DIAGRAM 2) is placed in an adjusting frame (FIG. 2b, DIAGRAM 3) and the straws are lined up (FIG. 3). The pin is put through both clamping pieces and screw-tightened for fixing. When straws of 3 mm are used, an adapting tube with a diameter of 4 mm and a wall thickness of 1 mm is placed on the straws and the pin is stuck through the tube and the clamping pieces. The straws (maximum three strawholders at a time) are then heat-sealed with an Automaster sealer (AM 400, Audion Electron, Amsterdam), which has a seal of 5 mm wide (FIG. 4). Each straw is hand-labeled with a CBS accession number using an alcohol-resistant microwriter (black, Staedtler Lumocolor 318). All 12 straws are then put into a bacteriological screw cap bottle (28 ml, 82.5 x 28 mm, A. Gallenkamp & Co. Ltd., London) and sterilized in an autoclave at 120 C for 20 min.

Handling of straws

All manipulations are carried out in a Biohazard recirculating laminar flow cabinet (CLF 406, Clean Air, Woerden, The Netherlands) using sterilized instruments. After lining up the straws, the straw holder is placed on a support (FIG. 2a, DIAGRAM 4) at angles of 30 or 45 degrees dependent on the preference of the technical assistant. The straws are then filled half full with a cryoprotectant. The standard is 10% aqueous glycerol, but some fungi require 10% dimethylsulfoxide (DMSO in water) (Hwang et al., 1976) or an aqueous solution containing 8% glucose and 10% DMSO (Smith. 1983). From agar cultures in Petri dishes 5-8 plugs of 2.8 mm diam (for 3 mm straws 1.8 mm diam) are punched out with a cork borer with pin (DIAGRAM 5) and transferred into the straw. When all straws are filled the series is heat-sealed.

Compared with the first sealing, the sealing time will be longer as a consequence of autoclaving and the possibility of some moisture being present at the inner side of the straw at the place of sealing. All straws are checked for leakage and if necessary resealed. Then labels of 7 x 5 mm with the CBS accession number and provided with an ultra low temperature resistant rubber glue (Nederlandsche Speciaal Drukkerijen, Delft, The Netherlands) are glued on the upper seal. Finally, the straws are placed in a drawer, cooled with a rate of -1 C/min to -40 C in a plasma freezer (Forma Scientific, U.S.A., model 8070) and stored in liquid nitrogen.

For each strain 12 straws are prepared; 8 are stored in their final unit, one is stored separately for a viability check within a week and the others are stored in the first row of the same drawer in which the first 8 can be found. Of these, one is to be checked after a year and the remaining straws are extras.

Thawing takes place in a water bath for five minutes at 30-35 C (Oomycetes: 20 C). Then the straws are rinsed in ethanol 96%, opened and plated on suitable media.

The method has been applied to about 2000 strains and the results are summarized in TABLE I. This table is slightly optimistic, especially with regard to the Oomycetes, where the method has been modified after the results of the original method were disappointing (more than 50% failures). As the new method is not yet in use for a long time, and the number of unfinished tests is thus relatively large, the percentage of success will probably decrease to an estimate of about 70%. Moreover the strains were generally recently isolated (up to two years in culture, except for about 500 Basidiomycetes which were up to 12 years in culture). Even if these positive effect are eliminated, our results are still comparable with or better than those obtained with glass vials.


Figs. 1-3


Fig. 4, 5


Diagram 1


Diagram 2


Diagram 3


Diagram 4


Diagram 5


Table I


LITERATURE CITED

Elliott, T. J. 1976. Alternative ampoule for storing fungal cultures in liquid nitrogen. Trans. Brit. My colt Soc. 67: 545-546.

----, and M. P. Challen. 1979. The storage of mushroom strains in liquid nitrogen. Ann. Rep. Glasshouse Crops Res. Inst. 1979: 194-204.

Hwang, S.-W., W. F. Kwolek, and W. C. Haynes. 1976. Investigation of ultra low temperature for fungal cultures III. Viability and growth rate of mycelial cultures following cryogenic storage. Mycologia 68: 377-387.

Smith, D. 1983. Cryoprotectants and the cryopreservation of fungi. Trans. Brit. Mycol. Soc. 80: 360363.

Accepted for publication July 22, 1986.

     
 
Freeze-drying of micro-organism using a simple apparatus by Webmaster Online protocols 2016-02-06 18:33:13
 

Khursheed A. Malik, Ph.D.

DSM-Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH,
Mascheroder Weg 1B, D-3300 Braunschweig, Federal Republic of Germany

World Federation for Culture Collections
Technical information sheet No. 7

Published by: UNESCO / WFCC - Education Committee 1990

Introduction

The advantages of freeze-drying are obvious. It is a convenient method for the preservation and long-term storage of a wide variety of microorganisms. However, special precautions are needed for the preservation of microorganisms sensitive to desiccation, light, oxygen, osmotic pressure, surface tension and other factors. During several years of experimentation I have developed various new methods and have optimised freeze-drying conditions to achieve successful Iyophilization of a large collection of difficult and fragile microorganisms.

Normally specialized equipment is required to create the conditions-conducive to the freeze-drying process. The costs of such specialised equipment required for freeze-drying can be substantial, and thus the process may appear to be expensive in spite of its many advantages. A freeze-drying process is described here using a simple apparatus.

This method is based on a new freeze-drying method which has recently been described (Malik, 1988). Some effective protective agents (Malik, 1976 & 1988) for example skim milk and meso-inositol or honey or glutamate or raffinose, are used to suspend cells to be freeze-dried in order to protect these against known freezing and drying injuries.

Several anaerobic bacteria which are sensitive to aerobic freeze-drying, can successfully be preserved using activated charcoal (5 % w/v) in the suspending media along with the above protective agents (Malik, 1990).

With this simple method various yeasts, sporulating fungi, and bacteria can successfully be preserved. Many delicate microorganisms such as nitrogen-fixing bacteria like Azospirillum, Azotobacteraceae, Rhizobium, Xanthobacters, Spirillaceae, Vibrios, and others like Alcaligenes, Ancylobacter, Flavobacterium, Pseudomonads and few Rhodospirillaceae resulted in a fairly good viability and stability after Iyophilization and during storage. However, relatively low viability results in the case of various yeasts such as Leucosporidium, Sporobolomyces, Rhodosporidium, Zygosaccharomyces and fragile bacteria like Aquaspirillum, Spirosoma, Flectobacillus, and others, as compared to the Iyophilization done under standard conditions (Malik, 1988 & 1990). The method is simple and the equipment described here can be easily constructed in most laboratories.

Material and Methods

Preparation of freeze-dried skim milk ampoules

Ampoules of neutral glass (45 x 10 mm) are washed in a detergent, then rinsed in distilled water and are air dried. For double vial preparation the ampoules are labelled and filled with 0,5 ml of 20 % (w/v) skim milk (Bacto-Skim milk Difco 0032) containing 5 % meso-inositol or 5 % raffinose or 10% honey. The ampoules are loosely plugged with non-absorbent cotton wool and sterilized at 115' C for 13 minutes. These are frozen at about -30 lo -40' C for a few hours and are freeze-dried for about 6 hours, as described under the freeze-drying procedure (primary freeze-drying).

Large batches of ampoules should be avoided but this depends upon the capacity of the freeze-drying system ( vacuum pump, condensing temperature of the cold trap, etc.).

Preparation of protective agents

Solutions of most effective protective agents like meso-inositol (5 % w/v), honey (10 % w/v), sodium glutamate (5 % w/v), raffinose (5 % w/v) are prepared in distilled water, filter sterilized and stored at 4' C. For use these are selected according to the storage temperature available for storage of freezedried ampoules. For more details see Mali, 1988.

Preparation of cell suspension for freeze-drying

The cultures are grown on appropriate media until the late logarithmic phase of growth. A thick cell suspension (at least 10(8) cells per ml) is prepared in an appropriate protective medium. In the case of liquid cultures the cells are harvested by aseptic centrifugation for 30 minutes at 4000 xg in screw-cap bottles and the pellet is suspended in a protective medium to yield a heavy cell suspension. The ready ceil suspensions are kept in an ice bath before filling the ampoules.

Filling of ampoules and slow freezing of cell suspension

The already freeze-dried skim milk ampoules are cooled to about -30' C for 1-2 hours. To each ampoule about 0,03 ml (1 drop with a Pasteur pipette) of ice cold cell suspension is added aseptically on to the skim milk plug with care so as not to touch the sides of the ampoules. The ampoules are quickly placed again for 1-2 hours in a deep-freeze and are frozen at approximately 1-2'C/min to about -30' C. This is easily achieved if ice cold samples are placed in commercially available deep-freezers.

The freeze-drying procedure

Freeze-drying involves the removal of water from frozen cell suspension by sublimation under reduced pressure. The outline of the freeze-drying procedure and the major steps involved are shown in Fig. 1.

The cold trap tube (a U-shaped thick glass or preferably a metal tube of about 2-3 cm diam and about 30 cm length), is connected with the vacuum hose (preferably the U-tube is filled with blue/ dry silica gel) and is placed in a metallic beaker as shown in Fig.1 A. It is chilled to about -35'C. A cooling mixture of ethylene glycol : water (1: 1) is placed for few hours in a deep-freezer and is cooled down to about -30'C (preferably to -40'C). This is poured into the cold trap to give a maximum depth. The cooling mixture is also sealed in deep-freeze plastic bags and is cooled down or frozen in a deep-freezer or over liquid-nitrogen. This super cooled or frozen coolant in the bags is added to the cold trap and the bags are changed periodically (preferably after every 20 -25 min) throughout the Iyophilization run in order to maintain the temperature at a minimum level. Commercially available anti-freeze liquids used in car radiators as coolants are also satisfactory as an alternate for the low-temperature bath and cold trap. A double jacketed straight tube (exterior about 6x 30 cm and interior about 3x25 cm with outlet and inlet tubes of about 2x5 cm, as shown in Fig.1A) can also be used as a cold trap.

If available, dry ice is used for freezing the ampoules, to cool down the cold bath and is added to the cold trap periodically throughout the experiment.

For the double vial preparation freeze-drying is done in two stages involving primary freeze-drying and secondary drying. When the condenser (cold trap) temperature has reached about -35' C the frozen (to about -30' C) ampoules are transferred quickly to the chilled metallic evacuation jar, which is dipped to about 2 cm depth in the cold bath containing coolant at about -35'C (Fig.1 A). The vacuum is switched on and the temperature of the cold bath is allowed to elevate. If available, a vacuum meter or controller can be attached between the vacuum pump and the evacuation jar to control the vacuum. Primary-drying is continued for about 3 hours to achieve maximum desiccation ( at about 1 to 0,1 torr or mbar). At the end, the vacuum is replaced with nitrogen gas (especially for ampoules which are not subjected to secondary drying and are not to be sealed under vacuum). At the end of the experiment the water collected in the cold trap is drained out or the silica gel is replaced.

Constriction of ampoules, secondary drying and sealing

In a double vial system the ampoules (containing primary freeze-dried cell material) are sealed under vacuum in a soft glass tube. After primary freeze-drying of the ampoules the projecting ends of the cotton plugs are trimmed in level to the end of the ampoules and these are transferred into soft-glass large tubes (130 x 15 mm) containing blue silica gel and cotton plugs. For insulation small amount of glass wool is also pushed down along with the ampoule (inner vial ) to the bottom of the large tube (outer vial). This outer tube is then constricted, on a low flame by hand or by using Edward Ampoule Constrictor, 20-25 mm above the glass wool to avoid burning of the cotton plug of the inner vial. The constricted tubes are then attached to the manifold and mounted on a metallic evacuation jar. This operation is illustrated in Fig.1 B . The vacuum is switched on and secondary drying is conducted for 12 hours (at 0,1 to 0,001 torr). The pink silica gel in the outer tubes will turn again to blue at this stage.

The constricted outer tubes are carefully sealed, by hand or by using a Flaminaire blow torch, one by one maintaining vacuum (Fig.1 C). To avoid cracking of glass a flame containing oxygen should never be applied to such neutral glass that has not previously been warmed on a normal flame

Revival of cultures from freeze-dried ampoules

During reactivation of preserved microorganisms it is recommended to use the most favourable media and growth conditions. During several years of experience, I have observed that in the case of sensitive microorganisms when the preserved (Iyophilized or cryogenically stored) cultures are revived, the counts on agar media are usually lower than in liquid media and similarly agar media of higher surface tension (such as nutrient agar) usually results in lower viable counts than mineral media of relatively lower surface tension. The contents of the freeze-dried ampoule are thus dissolved in sterile liquid growth media and incubated at a relatively lower temperature than the optimum growth temperatures. A few cultures may exhibit a prolonged lag period and thus are incubated for relatively longer periods. Normal growth usually appears after a second transfer into fresh medium. In a few cases growth is inhibited by the high concentration of protective mixture used during Iyophilization.

During reactivation the presence of activated charcoal in the suspending media results in higher survival recoveries, and the reactivated cultures grown in the presence of activated charcoal prove much more stable and could be maintained relatively longer as living cultures (Malik,1990).

The use of activated charcoal as an adsorbent of harmful radicals in suspending media for the reactivation of anaerobes is also recommended

Estimation of viability and stability

Survival recoveries are chocked before freeze-drying, immediately after freeze-drying and after storage. For the estimation of viability counts serial dilutions are prepared in liquid media. From each serial dilution 0,1 ml volumes are plated on appropriate growth agar media plates. The number of colonies are counted from the plates and average colony forming units per sample are calculated. The revived cultures are also observed for mutation, change in colony morphology or other characters.

Long-term storage

Stability of freeze-dried cultures during storage is very important. A high level of residual moisture content or exposure to oxygen have detrimental effects on the freeze-dried product. Freeze-dried material is hygroscopic and its exposure to moisture during storage can destabilise the product.

The higher the storage temperature, the faster a product will degrade. Thus, the storage of freeze-dried cultures at lower temperatures will extend their shelf life. The unsealed freeze-dried ampoules can safely be stored for several years at about -30'C. It has been observed by the author that similar viability counts were obtained after 4-5 years of storage when unsealed freeze-dried culture were maintained at -30'C as compared to the freeze-dried cultures which were sealed under vacuum and were stored at 9'C (Malik, 1976).

Selected references

Ellis, J.J. and J.A. Roberson. 1968. Viability of fungus cultures preserved by Iyophilization. Mycologia 60: 399-405

Heckly, R.J. (1985). Principles of preserving bacteria by freeze-drying. Developments in Industrial Microbiology 26: 379-395

Mackenzle, A.P. (1977). Comparative Studies on the freeze-drying survival of various bacteria: Gram type, suspending medium and freezing rate. Develop. biol. Standard, Vol.36: 263-277 (S. Karger, Basel).

Malik, K.A. (1976) Preservation of Knallgas bacteria. In Proceedings of Vth Intern. Fermentation Symposium (H.Dellway, Ed.) p. 180 Westkreuz Druckerei and Verlag, Bonn and Berlin.

Malik, K.A. (1987). The role of culture collections in the stability and preservation of microorganisms (J.Amen and P. Tesson, Eds.) pp 118-150. Societe Francaise de Microbiologie. Paris

Malik, K.A. (1988). A new freeze-drying method for the preservation of nitrogen-fixing and other fragile bacteria. J. Microbiol. Methods 8: 259-271

Malik, K.A. (1988). Long-term preservation of some Rhodospirillaceae by freeze-drying. J. Microbiol. Methods 8: 273-280

Malik, K.A. (1990). Use of activated charcoal for the preservation of anaerobic phototrophic and other sensitive bacteria by freeze-drying. J. Microbiol. Methods

     
 
Cryopreservation of yeasts in polypropylene straws by Webmaster Online protocols 2016-02-06 18:33:27
 

J. Henry and B. Kirsop

National Collection of Yeast Cultures AFRC Institute of Food Research
Norwich Laboratory Colney Lane Norwich NR4 7UA United Kingdom

World Federation for Culture Collections
Technical information sheet No 3

Published by: UNESCO / WFCC - Education Committee 1989

Introduction

It has been shown (1, 2) that the survival levels of yeasts stored in liquid nitrogen at -196C are high and superior to those obtained with other preservation methods. Good strain stability following cryopreservation is also recorded.

Storage at temperatures -130C below which molecular activity does not occur, can be achieved by storage in the liquid phase of nitrogen or in mechanical refrigerators operating at these temperatures (3) and are to be preferred to storage at high temperatures. The method described here is for storage in liquid nitrogen.

Cultures may be stored in polypropylene cryotubes, glass vials or polypropylene straws. This method uses straws and has the following advantages:

  1. Economies of space (5-6 straws can be stored in each cryotube).
  2. Prevention of seepage of liquid nitrogen into the samples; liquid nitrogen may penetrate the washers of polypropylene cryotubes, but cannot penetrate the sealed straws.
  3. Polypropylene drinking straws are cheap and readily obtainable throughout the world.
  4. Polypropylene is very much safer to use than glass vials.
  5. The straws are available in different colours, allowing colour-coding of samples.
  6. The culture can be recovered for use by removal of a single straw, the rest of the samples remaining frozen.

Method

1. Preparation of straws

Coloured polypropylene straws (available from most catering distributors and many shops) are cut into 2.5 cm lengths. One end of a straw is sealed by holding firmly in a pair of unridged forceps 1 mm inwards so that the projecting end is 1 cm from the flame of a fish-tail bunsen burner. The polypropylene melts almost immediately and forms a strong seal that sets firm within a second or two.

The straws are placed in a glass petri dish and autoclaved at 121C for 15 min.

2. Inoculum

The culture to be frozen is grown in YM broth (Difco Laboratories Ltd. 0711-01) at 25C for 72 h, if possible on a reciprocal shaker. Each straw requires 0.1 ml of a suspension containing between 10(6) and 10(7) cells/ml. Cell concentration has been found to have little effect on the percentage of cells surviving.

3. Preparation of cryoprotectant

A 10% glycerol solution is prepared, filter-sterilized, and stored in sterile screw-cap bottles.

4. Inoculation of straws

Equal quantities of inoculum and cryoprotectant are mixed aseptically in a sterile bottle. A single straw is removed with forceps from the petri dish and filled with inoculum using a Pasteur pipette. When filling it is necessary to place the end of the pipette close to the sealed end of the straw and to fill to within 3 mm of the open end. The filled straw is then sealed at the open end as described in step 1. above.

Six straws are placed in each plastic, screw-capped 2 ml ampoule (Nunc, Gibco-Europe Ltd).

5. Freezing

Primary freezing. It is very important to wear protective clothing when using liquid nitrogen refrigerators or handling frozen specimens. Ampoules are frozen to -30C by placing in a refrigerated room. If a room at -30C is not available, a refrigerated cabinet or cooling bath Camlab Ltd) may be used. If aluminium canes (Union Carbide) are to be used for the secondary freezing, ampoules may be placed on the canes at this stage. If secondary freezing is to take place in storage drawers (Union Carbide), the ampoules may be well spaced in wire racks for the primary freezing. The cooling rate is not critical in this method, but is probably in the region of 5C/min., depending on the size of samples and containers. Cells are held at -30C for 2 h to allow dehydration to take place.

Secondary freezing. The ampoules are transferred to the canisters or storage drawers of the refrigerator (Union Carbide) and immersed in the liquid nitrogen, care being taken to prevent the samples from thawing. If the distance between primary and secondary freezing containers is great, samples should be transported in a chilled Dewar or in any other suitable container.

6. Revival

Using sterile forceps, a straw is removed from the ampoule and placed immediately into a screw-cap bottle containing water at 35 C. The bottle is shaken to facilitate rapid thawing. Several straws may be thawed simultaneously in this way.

7. Viable counts

Before opening the straws, the cells are redispersed by squeezing the straws carefully several times. The straws are then wiped with 95% alcohol and one end is cut off using sterile scissors. The contents are removed using a Pasteur pipette. It may be necessary to disperse cells further by repeated pipetting at this stage.

Two drops of the suspension (0.06 ml) are transferred to 0.54 ml sterile water to make a 101 dilution. Further dilutions, plating and counting are carried out using standard methods (4).

8. Storage

Ampoules are stored in the liquid nitrogen refrigerator (Union Carbide), care being taken to maintain the liquid nitrogen at such a level that ampoules are completely submerged.

Notes

1. A number of other cryoprotectants have been used successfully both by the NCYC and other workers. Some have been used for a fairly wide range of yeasts, others with a few strains only. Substances used successfully include glycerol (20%, 10%, 5%), dimethyl sulphoxide (DMSO, 10%), glycerol plus DMSO, ethanol (10%), methanol (10%), YM broth and hydroxyethyl starch (10%, 5%).

2. The NCYC has found that primary freezing and dehydration at -20C, -30C or -40C for 1, 2 or 3 h is equally successful for the two test strains of Saccharomyces cerevisiae used to develop the method. The intermediate protocol (-30C for 2 h) has been adopted and proved successful for a wide range of species.

3. The NCYC has found that, in general, higher levels of survival are obtained from aerobically grown cultures than from those grown with limited access to oxygen.

4. It is important that the outside of the straws is kept dry during filling, as wet straws do not seal well.

5. When filling several straws with the same inoculum, it is convenient to lay each straw as it is filled against a glass rod in a sterile petri dish until all straws are filled. The straws are then sealed and put into ampoules. This is more convenient than filling and sealing each straw separately.

6. If the outside of the straws are dry, they do not adhere to each other when placed in ampoules. Removal of straws from ampoules is facilitated if straws vary slightly in length.

7. The different coloured straws can be used to colour-code yeast strains to aid retrieval from the refrigerators, and it is clearly sensible to store only one yeast strain in each ampoule.

Survival levels and shelf life

Survival levels between 50 and 100% are frequently obtained for a wide range of yeast species, using the standard method. Higher survival levels for individual strains could probably be obtained by careful adjustments to the protocol. The NCYC has detected no drop in viability in test strains over a period of 26 months and others record good survival for periods of up to 4 years. In view of the high initial survival rates, shelf life can be expected to be good.

References

Hubaleck, Z. & Kockova-Kratochvilova, A., 1978. Antonie van Leeuwenhoek 44, 229-241.

Kirsop, B.E. & Henry, J.E., 1984. Cryoletters 5, 191-200.

White, W. & Wharton, K.L. 1985 (Jan-Feb). International Laboratory, 58-69.

Miles, A.A. & Misra, S.S., 1958. J. Hyg Camb. 38, 732-749.

     
 
Cryopreservation of fungi by Webmaster Online protocols 2016-02-06 18:33:41
 

P. Hoffmann, Ph.D.

DSM-Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH
Mascheroder Weg 1 B, D-3300 Braunschweig, Federal Republic of Germany

World Federation for Culture Collections
Technical information sheet No 5

Published by: UNESCO / WFCC - Education Committee 1989

Introduction

Cryopreservation in liquid nitrogen (LN) is a reliable method for long-term storage of microorganisms. Different protocols have been published (Kirsop & Snell, 1984; Smith & Onions, 1983; Elliott, 1976; Stalpers et al., 1987), varying with equipment, special needs, preference of materials or the type of organism. A simple method is described here, which is applicable to a large variety of microorganisms such as mycelial fungi, molds, and yeasts.

Materials and Methods

Equipment

Different types of storage tanks which accommodate 1.0 to 1.8 ml cryotubes (e.g. Intermed NUNC, Denmark) may be employed. The "canister & cane system" (e.g. COSMOS L40, Messer Griessheim, F.R. Germany) (Fig. 1 I,K), if available, is preferred because it allows the removal of only one "cane", holding several ampoules at a time (Fig. 1 I). The storage capacity of tanks employing the "drawer system" is greater (e.g. BT 55, Air Liquide, France), however, LN evaporation rates are higher and removal of one ampoule requires lifting of a whole set of drawers out of the storage container.

PVC (polyvinylchloride) or polypropylene "straws" with a diameter of 2-3 mm are cut to a length of approx. 25 mm on a paper cutting machine. One end of the PVC straws is immersed into acetone for a few seconds and is heat-sealed at a temperature of approximately 280ÈC; other materials may be sealed in a gas flame. The sealed straws are sterilized by autoclaving (15 min, 121ÈC). PVC straws not treated with acetone usually reopen during sterilization.

For sealing of the straws a commercial household sealing machine with adjustable temperature may be used. Forceps with specially designed tips will greatly facilitate handling of the straws

A block accommodating the frozen cryotubes is recommended, if opening of the tubes outside of the container is necessary and thawing of the remaining straws is to be avoided. The block is either made of aluminium, brass or copper and is surrounded by a styrofoam carrying case. It will keep the temperature of the cryotubes below -120ÈC for about 15 min.

Preparation of organisms

A schematic outline of the procedure is given in Fig. 1. Yeasts are grown in liquid culture (Fig. 1B) or on a suitable solid medium (Fig. 1A) to a colony size of approximately 2 mm diam. Two colonies of the strain are removed from the agar with a loop, carefully suspended in 1.5 ml sterile glycerol (10 % w/v in water) (Fig. 1C) and filled into the sterile straws (sealed at one end) with a disposable syringe or Pasteur pipette (Fig. 1D). The straws may then be sealed completely and transferred aseptically to a sterile cryotube (Fig. 1H).

Sporulating fungi are grown on solid media until conidia develop. A heavy conidial suspension is prepared in glycerol (10% w/v) which is treated as in the case of yeasts.

Mycelial fungi are grown in media supplemented with 5 % (w/v) glycerol (Fig. 1E). Strains that do not tolerate the lower water activity caused by the cryoprotectant may be grown without glycerol and flooded with a 10% (w/v) glycerol solution shortly before processing. A sterile straw open at both ends is now used to punch the mycelium with the agar from near the margin of the colony (Fig. 1F). This procedure is repeated until the straw is filled completely (Fig. 1G). The straw is either left open at both ends and transferred aseptically to a sterile cryotube or it may be sealed.

Freezing

To obtain a freezing rate that is close to the theoretical optimum of 1-10ÈC per minute, the cryotubes are either transferred to a mechanical deep freezer at -70ÈC for two hours in a styrofoam box of 2 cm wall thickness or placed in the gas phase of a liquid nitrogen tank for about 40 min.

Thawing

For revival, one straw at a time is removed from the frozen cryotube; the sealed straws are transferred into a 50 ml glass beaker with warm water (30ÈC), open straws filled with mycelial fungi are thawed directly on agar slants at room temperature (22 to 25ÈC). Sealed straws may be surface sterilized by immersion into 70% ethanol (v/v), before they are opened with sharp, sterile scissors or pincers. The cell suspension is withdrawn with a fine Pasteur pipette. Incubation is done at appropriate temperatures until growth is visible.

Selected references for further reading

Elliott, T.J. 1976. Alternative ampoule for storing fungal cultures in liquid nitrogen. Trans. Brit. Mycol. Soc. 67, 545-546.

Kirsop, B., Snell, J.J.S. (eds.) 1984. Maintenance of Microorganisms. Academic Press, London,

Smith, D., Onions, A.H.S. 1980. The preservation and maintenance of living fungi. Commonwealth Mycological Institute, Kew, U.K.

Stalpers, J.A., de Hoog, A., Vlug, IJ. 1987. Improvement of the straw technique for the preservation of fungi in liquid nitrogen. Mycologia 79, 82-89.

     
 
Cryopreservation of bacteria with special reference to anaerobes by Webmaster Online protocols 2016-02-06 18:33:55
 

Khursheed A. Malik, Ph.D.

DSM-Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH,
Mascheroder Weg 1b, D-3300 Braunschweig, Federal Republic of Germany

World Federation for Culture Collections
Technical information sheet No 4

Published by: UNESC/WFCC - Education Committee 1989

Introduction

The methods used in the maintenance of stock cultures of microorganisms usually involve serial subculturing or other simple methods of preservation which are not only time consuming but can also result in genetic instability. Cryopreservation of microorganisms in liquid nitrogen at -196C is a very reliable method and is generally considered superior to other preservation methods. Bacteria preserved in liquid nitrogen normally show high survival rates and good strain stability during long-term storage. In liquid-nitrogen storage of microorganisms polypropylene cryotubes, glass vials, glass capillaries and polypropylene straws are generally used. In the method described here, screw-cap polypropylene cryotubes and mini-screw cap glass ampoules have been used. The latter are also suitable for the preservation of bacteria under anaerobic conditions. The method is simple, effective and economical with respect to the storage space and costs. In the preservation of anaerobes, generally no continuous stream of nitrogen gas, no anaerobic chamber or glove boxes are required. This method provides a suitable model for cryogenic storage of many fastidious and delicate bacteria.

General considerations for successful cryopreservation

Several factors can affect cell viability and stability during cryopreservation. During cryopreservation, dehydration of cells results and osmotic imbalance is created due to the changes in the concentration of salts and other metabolites. During the cooling process rupture of the cellular membranes can also occur by the formation of large ice crystals. Successful preservation can be achieved by the use of cryo-protective agents (such as dimethylsulfoxide, glycerol), maintaining a controlled rate of cooling (about 1C per minute to about -30'C) and an appropriate rewarming protocol (rapid thawing in a 37C water bath which takes about one minute for a glass ampoule and somewhat longer for a plastic vial ). In practice, a relatively slow cooling rate can be easily obtained by keeping ampoules/vials in mechanical deep-freezers for 1-2 hours or in the neck of the liquid nitrogen storage unit for some minutes and then lowering containers into it. It is, however, not good practice to plunge cultures directly into liquid nitrogen, as the liquid nitrogen may seep into any imperfectly closed or sealed capillaries, ampoules, or vials containing the bacterial suspensions. On removal from storage, nitrogen (inside an ampoule) will virtually instantly change to the gaseous phase causing an explosion. For safety reasons it is thus recommended that cultures should be stored in the gas phase of liquid nitrogen.

While preparing cells for cryopreservation several factors such as optimal growth conditions, physiological state of the cells (preferably from the late logarithmic to early stationary phase of growth), high cells density (106 to 10 cells per ml) should be considered as these can affect cell viability after cryopreservation. After mixing, cell suspensions should be kept for equilibration with the cryoprotective agent. For harvesting, liquid cultures are centrifuged. However, vigorous pipetting and high-speed centrifugation should be avoided and cells should be handled gently. Viability assays should be performed on all cultures before and after cryopreservation to assure long-term viability. To assure purity, identity of the preserved cultures should be verified and after freezing cultures should be recharacterized to assure their stability. Safety precautions should be observed when removing an ampoule from liquid nitrogen. The face shield, laboratory coat and insulated gloves should be worn as protection against liquid nitrogen splash and exploding ampoules. The level of the liquid nitrogen in the containers should be checked preferably on a daily basis and maintained to a constant level, as any drop in liquid nitrogen level below a critical volume can result in damage due to the warming of the samples.

Materials and Methods

Equipment

Screw-capped plastic cryovials of about 2 ml capacity (available from Nunc, Gibco Europe Ltd., or Nalge Company). These are supplied in sterile packings. These are not suitable for repeated use. Screw-cap glass ampoules (10x30 mm) of 2 ml capacity (available from Varian GmbH, Darmstadt FRG). These are generally used for autosamplers in gas chromatography and are provided with rubber septa and plastic screwcaps with holes for the injection of the samples. It is recommended to use ampoules with black screw-caps and oxygen-impermeable butyl rubber septa. The ampoules are washed, rinsed with distilled water, tightly closed and autoclaved. Before use these are labelled with the numbers of the strains to be preserved. Liquid nitrogen storage tanks with canisters, racks, canes, (supplied by Union Carbide; Air Liquide, France; Messer Griessheim, FRG; or else where) and precautionary arrangements like safety glasses, gloves etc.

Anaerobic facilities for the preparation of reduced media. Hungate tubes with septa (Bellco Glass Inc., 2047-16125). Butyl rubber overflow tubes (about 5 mm diameter) with Luer Lock adapters at both ends and long syringe needles (10-15 cm in length). Sterile gas tight hypodermic Luer Lock syringes. Cryoprotective agent glycerol and dimethylsulfoxide (DMSO), reagent grade. Glycerol (20% w/v in H20) may be sterilized by autoclaving for 15 minutes and is stored in screw-cap bottles in dark. DMSO (20% v/v in H20) is sterilized by filtration (using a Teflon syringe filter) or can be autoclaved undiluted at 114C for 10 minutes.

Preparation of cell suspension for freezing

The aerobic cultures to be frozen are grown in appropriate media (under well aerated conditions ) and should be harvested preferably in active phase of growth. From agar slants the cultures are removed with a loop and gently suspended in sterile glycerol (10% w/v) or DMSO (5% v/v) to obtain a heavy cell suspension. For anaerobes this is done under a stream of sterile nitrogen gas. Thick suspensions (108 -10'' cells/ml) of liquid cultures are mixed in equal quantities with the double concentrated cryoprotective agents. For harvesting, the anaerobic cultures are centrifuged for 30 minutes at 4000 xg in the screw-cap bottles in which cultures are grown. The supernatant is removed anaerobically under a stream of nitrogen gas using an overflow butyl rubber tube of about 5 mm diameter with Luer Lock adapters at both ends and fitted with long syringe needles of 1015 cm length (see Fig. 1A). To obtain sterile nitrogen gas a sterile, cotton filled syringe is attached to a conduit connected to the N2 gas (99.99%) cylinder. The pellet is resuspended carefully in ice cold sterile DMSO solution (5% v/v in H2O). In the case of halophilic strains or cells which do not form a pellet a thick bacterial suspension (in growth medium) is mixed in the ratio 3:1 with ice cold sterile DMSO (20% v/v in H2O). For extreme halophilic strains optimum salt concentration should be maintained after mixing cell suspension with the DMSO. The cells are allowed to equilibrate with the cryoprotectant (15 minutes for DMSO, 30 minutes for glycerol) in an ice bath.

Filling of ampoules and freezing

While equilibrating, an aliquot of 1.0 to 1.5 ml of cell suspension is dispensed in to each plastic cryovial or glass ampoule. For anaerobes using a sterile gas-tight 5-10 ml syringe, the ampoules are evacuated for anaerobiosis and to facilitate filling (Fig. 1B). About 1 ml of thick cell suspension (equilibrated with the DMSO) is withdrawn with a 1 ml sterile oxygen free syringe (already flushed with nitrogen gas) and injected into each ampoule (Fig. 1C) Immediately after the glass ampoules or cryotubes are clamped onto labeled aluminium canes, placed at -30C for about one hour or for few minutes in the gas phase of liquid nitrogen. The canes are then placed in canisters, racks or drawers and frozen by direct immersion in liquid nitrogen or in the gas phase of liquid nitrogen (Fig. 1D).

Revival of cultures

The frozen ampoule is removed from liquid nitrogen. For partial thawing these are immediately immersed to the neck in the mini water bath at 37C (Fig. 1E) for a few seconds. After thawing the outer surface of the ampoules is dried by wiping and plastic vials are wiped with alcohol-soaked gauze prior to opening. For aerobic bacteria the screwcap glass vials can be opened and flame sterilized at the neck. The thawed contents of the ampoule/vial are immediately transferred to fresh growth medium to dilute the cryoprotectant, which other wise is lethal at higher temperatures. For anaerobes the septum of the glass ampoule is flame sterilized after putting a drop of alcohol and with a 1 ml oxygen free syringe a small volume (about 0.05 ml) of inoculum is withdrawn and injected into 5-10 ml liquid growth medium (Fig. 1F). The rest of the cell suspension is immediately frozen again (a self made wax block rack, chilled to -30C is used for transportation to the liquid nitrogen container see Fig. 1G) in liquid nitrogen for later use. In this way one ampoule can be used for several repeated retrievals or inoculations. The DMSO which is often toxic during growth is diluted 100-200 times in the culture medium to a non inhibitory concentration. The inoculated growth medium is incubated under appropriate growth conditions.

Estimation of viability counts

For aerobic bacteria, 0.5 ml of inocula is transferred to 4.5 ml of liquid growth medium and serial decimal dilutions are prepared. Plating and counting are done using standard methods. For the estimation of viable cell counts in anaerobic bacteria, 0.5 ml of inocula is transferred from the unfrozen (for cell counts before freezing) and from the thawed cell suspension (for cell counts after freezing) into prereduced 4.5 ml medium in screw-cap tubes (Hungate tubes with septa, Bellco Glass Inc., 2047-16125) and 6-8 serial decimal dilutions are prepared using oxygen free syringes and incubation is done under appropriate conditions. Agar roll tubes can be prepared for viable colony counts determination if such facilities are available. Colony counts on agar plates can be performed in an anaerobic glove box or anaerobic jars. Single plates can be incubated anaerobically in anaerobic Biobags (Type A, Marion Scientific Corporation, Kansas City, Ml, USA) or in Anaerocult bags (E. Merck, Postfach 4119, D-6100 Darmstadt). In the case of viable colony counts in agar roll tubes or on plates the number of colonies are counted from each dilution and average colony forming cells per sample are calculated.

For cultures which are difficult to grow in, or on agar, only liquid dilutions series are made. In this case the number of cells is determined using the most probable number method (MPN).

For comparison the viable cell counts before freezing and after freezing are recorded and percentage survival is calculated.

Selected references for further reading

Greiff,D., H. Melton and T.W. Rowe, 1975. On the sealing of gas-filled glass ampoules. Cryobiology 12: 1-14.

Kirsop, B. and J.J.S. Snell (eds.), 1984. Maintenance of Microorganisms: A Manual of Laboratory Methods. Academic Press, London.

Malik, K.A., 1977. Rapid surface colony counts determination with three new miniaturised techniques. Zbl. Bak. Hyg. I. Abt. A. 237: 415-423.

Malik, K.A., 1984 . A new method for liquid nitrogen storage of phototrophic bacteria under anaerobic conditions. Journal of Microbial Methods 2: 41-47

Malik, K.A., 1985. Modern Methods of Gene Conservation. A Laboratory Manual. PASTIC Press,lslamabad, Pakistan.

Simione, F.P., P.M. Daggett, M.S. MacGrath and M.T. Alexander, 1977. The use of plastic ampoules for freeze preservation of microorganisms. Cryobiology 14: 500-502

     
   

K. Painting and B. Kirsop

National Collection of Yeast Cultures, AFRC Institute of Food Research, Norwich Laboratory, Colney Lane, Norwich NR4 7UA, United Kingdom

World Federation for Culture Collections
Technical information sheet No 2

Published by: UNESCO / WFCC - Education Committee 1989

Introduction

It is sometimes necessary to obtain a quick estimate of the percentage of viable cells in a yeast sample. In traditional plate count methods results are not available until 3 or 4 days after inoculation A quick staining method can provide an estimate of viability in a few minutes.

Principle

Yeast cells that are viable contain an enzyme -that decolourises methylene blue, whereas dead cells do not. When cells from a yeast sample are suspended in the dye, it penetrates into all the cells, but is reduced only by the living cells. It is very simple, therefore, to distinguish between living and dead cells by examining them microscopically: dead cells are stained blue and living cells are unstained.

Preparation of methylene blue solution

Dissolve 0.01 g methylene blue in 10 ml distilled water. Add 2 g sodium citrate dihydrate and stir until dissolved. Filter through filter paper, making the volume up to 100 ml with distilled water.

Procedure

Mix equal volumes of yeast sample and methylene blue solution on a microscope slide. (It may be convenient to mix equal quantities of each, using a wire inoculation loop.) The cell concentration should be adjusted so that about 50 yeast cells are present in a microscope field, using a 40 x objective and 10 x or 12.5 x eyepieces.

Approximately 1000 cells should be examined and the percentage of unstained cells of the total recorded. This figure represents the percentage viability of the sample. When counting cells, buds that are less in size than half the parent cell are ignored.

Note: It should be remembered that this method measures the presence of an enzyme in a cell, rather than the ability of the cell to divide. It is possible that the enzyme is present in cells incapable of dividing and the method is thus less accurate than methods, such as plate count and slide count methods, which measure the ability of cells to produce daughter cells. It nevertheless provides a very useful rapid indication of the viability of yeasts samples.