The UKNCC Collections provide information in the catalogues and on the databases accessible on this web site. Below is some information on the growth of fungi and algae which may be useful. For information specific to the strains provided by the collections see catalogues or database. For further information contact the collection holding organisms you are interested in. During 1999 the UKNCC is producing a uses and properties manual which will provide an off the shelf publication providing with such information and more. An announcement will be made on the web site when it is available.

Growing Algae

Growing Fungi

GROWING ALGAE

There is an extensive literature on culture techniques and maintenance conditions (e.g. Stein, 1973; McLellan et al., 1991; Becker 1994). Medium composition depends both on the requirements of the algae (e.g. diatoms require the inclusion of a silica source; many marine algae require vitamins) and the preferences of the researcher. Information on medium suitability and full recipes are listed in the catalogues of all the major protistan culture collections (e.g. CCAP Catalogue: Tompkins et al., 1995). In general, master stock-cultures maintained by routine serial subculture are grown under sub-optimal temperature and light regimes [< 20oC and < 50 mmol photon m-2s-1 on shorter than normal daylight (12 h or less light : 12h or more dark)]. This type of regime maximises the interval between subcultures and thus minimises handling/transfers of the strain. Additionally, the use of medium containing organic carbon for maintaining axenic strains capable of heterotrophic or mixotrophic growth and solidified, rather than liquid, medium may be employed to maximise the period between transfer to fresh medium.

Details of medium requirements for individual strain requirements are listed on the UKNCC strain database and the CCAP catalogue and web-site (Tompkins et al., 1995; http://www.ife.ac.uk/ccap)

The CCAP also publish short booklets on this topic:
Culturing Algae - A Guide for Schools and Colleges
A Beginners Guide to the Collection, Isolation, Cultivation and Identification of Freshwater Protozoa

These may be purchased directly from the CCAP.

References

Becker, E. W. "Cambridge Studies in Biotechnology, Vol. 10. Microalgae: Biotechnology and Microbiology" Cambridge University Press, Cambridge, 1994.

McLellan, M.R., Cowling, A.J., Turner, M.F. and Day, J.G., 1991, Maintenance of algae and protozoa, in Kirsop, B. and Doyle, A. (Eds.) Maintenance of microorganisms and cultured cells, pp. 183-208, London: Academic Press Ltd.

Stein, J. (Ed.), 1973, Handbook of Phycological Methods: Culture methods and growth measurements. Cambridge: Cambridge University Press.

Tompkins, J., DeVille, M.M., Day, J.G. and Turner, M.F. (Eds.), 1995, Culture Collection of Algae and Protozoa Catalogue of Strains, Ambleside: Culture Collection of Algae and Protozoa

GENERAL HINTS ON GROWING FUNGI

Generally, fungi grow best on media that are formulated from the natural materials from which they were isolated. CABI Bioscience utilizes extracts from soil and plant materials such as leaves, stems or seeds placed on solid agar. Optimization of growth conditions is important. Avoidance of selection of variants from within the population, strain deterioration and contamination are important when growing strains for use and essential when maintaining cultures in this way for the long-term. The major factors affecting growth are medium, temperature, light, aeration, pH and water activity.

• Medium
• Temperature
• Light
• Aeration
• pH
• Water activity
• Near ultraviolet light (black light)
• Subculturing
• Mite infestation prevention
• References
• Recommended growth conditions for some fungi

Media

The growth requirements for fungi may vary from strain to strain, although cultures of the same species and genera tend to grow best on similar media. The source of isolates can give an indication of suitable growth conditions; thus isolates from jam can be expected to grow well on high-sugar media, species from leaves may sporulate best in light, those from marine situations may require salt, and those from hot deserts and the tropics, high growth temperatures.Cultures are usually best grown on agar slopes in test-tubes or culture bottles. The majority of fungi can be maintained on a relatively small range of media. However, some fungi deteriorate when kept on the same medium for prolonged periods, so different media should be alternated from time to time.Most laboratories prefer not to keep a large stock of different media and the majority of isolates can be maintained on a relatively small range depending on the specialization of the collection, e.g. medical fungi grow well on Sabouraud's medium. Experience at CABI, formerly IMI, is that cultures grow more satisfactorily on media freshly prepared in the laboratory, especially natural media such as vegetable decoctions. These are usually easy and relatively cheap to prepare and preparation can be carried out with limited facilities. Small quantities can be sterilized using a domestic pressure cooker and, if necessary, the pH can be adjusted using drops of hydrochloric acid or potassium hydroxide and measured using pH papers. However, proprietary media are often useful and can be very important in replicating work of others. Some media for special purposes such as assay work will require very careful preparation.A wide range of media are used by different workers. Authors tend to have their own favourite media. Raper & Thom (1949) use Czapek's Agar, Steep Agar and Malt Extract Agar for the growth of penicillia and aspergilli, while Pitt (1980) in his monograph on penicillia recommends Czapek Yeast Autolysate (CYA) and Malt Extract Agar (MEA).Preferences for growth on particular media are normally developed over many years and are the result of experience. The standardization of media formulae is necessary for most work. Media will affect colony morphology and colour, whether particular structures are formed and may affect the retention of properties. Examples of particular preferences are given below and in Smith & Onions (1994).

• Mucorales do well on Malt Agar (MA) and will not grow in Czapek Agar (CZ) as they lack enzymes to digest sucrose.
• Many fungi thrive on Potato Dextrose Agar (PDA), but this can be too rich, encouraging the growth of mycelium with ultimate loss of sporulation, so a period on Potato Carrot Agar (PCA), a starvation medium, may encourage sporulation.
• Fusarium species grow well on Potato Sucrose Agar (PSA).
• Wood inhabiting fungi and dematiaceous fungi often spore better on Cornmeal Agar (CMA) and Oat Agar (OA) both of which have less easily digestible carbohydrate.
• Cellulose destroying fungi and spoilage fungi, such as Trichoderma, Chaetomium and Stachybotrys retain their ability to produce cellulase when grown on a weak medium such as TWA or PCA with a piece of sterile filter paper, wheat straw or lupin stem placed on the agar surface.
• All sorts of vegetable decoctions are possible and apart from the advantages of standardization it is reasonable to use what is readily available, e.g. yam media might be preferable to potato media in the tropics.
• Entomophthora species can be grown in culture on several media but are reported to do best on an egg yolk medium.
• The introduction of pieces of tissue, such as rice, grains, leaves, wheatstraw or dung, often produces good sporulation. The use of hair for some dermatophytes has proved very successful (Al-Doory, 1968). Animal hair or feathers should be de-fatted in organic solvents first to ensure good growth.

Temperature

The majority of filamentous fungi are mesophilic, growing at temperatures within the range of 10-35°C, with optimum temperatures between 15 and 30°C. Some species (e.g. Aspergillus fumigatus, Talaromyces avellaneus) are thermotolerant and will grow at higher temperatures, although they are still capable of growth within the range 20-25°C. A small number (e.g. Chaetomium thermophilum, Penicillium dupontii, Thermoascus aurantiacus) are thermophilic and will grow and sporulate at 45°C or higher, but fail to grow below 20°C. A few fungi (e.g. Hypocrea psychrophila) are psychrophilic and are unable to grow above 20°C, while many others (e.g. a wide range of Fusarium and Penicillium species) are psychrotolerant and are able to grow both at freezing point and at mesophilic temperatures (see Smith & Onions 1994 for further information).

Light

Many species grow well in the dark, but others prefer daylight and some sporulate better under near ultraviolet light (see below). Most leaf- and stem-inhabiting fungi are light sensitive and require light stimulation for sporulation. At CABI BIOSCIENCE most cultures are grown in glass-fronted incubators to allow in the light or in illuminated incubators. In some cases a period of darkness is beneficial as some fungi are diurnal and require the transition from periods of light to dark to sporulate.

Aeration

Nearly all fungi are aerobic and, when grown in tubes or bottles, obtain sufficient oxygen through cotton wool plugs or loose bottle caps. Care should be taken to see that bottle caps are not screwed down tightly during the growth of cultures. A few aquatic Hyphomycetes require additional aeration, by bubbling air through liquid culture media for example, to enable normal growth and sporulation to occur.

pH

Filamentous fungi vary in pH requirements. Most common fungi grow well over the range pH 3 to 7, although some can grow at pH 2 and below (e.g. Moniliella acetoabutans, Aspergillus niger, Penicillium funiculosum).

Water activity

All organisms need water for growth, but the amount required varies widely. Although the majority of filamentous fungi require high levels of available water, a few are able to grow at low water activity (e.g. Eurotium species, Xeromyces bisporus). Some of those which occur on preserves or salt fish will only grow well on media containing high concentrations of sugar or salt. These fungi are referred to as xerophiles and halophiles respectively.

Near ultraviolet light (black light)

Fungi which require near ultraviolet (near UV and often referred to as black light, BL) light (wavelength 300-380 nm) for sporulation must be grown in plastic Petri dishes or plastic universal bottles for 3-4 days before irradiation. Glass is not suitable, as it is often opaque to ultraviolet light. Rich growth media should be avoided, as they may give rise to excessive growth of mycelium; nutritionally weak media such as potato carrot agar (PCA) are more suitable for inducing sporulation.At CABI BIOSCIENCE, three 1.22 m fluorescent tubes (a near ultraviolet light tube, Phillips TL 40 W/08, between two cool white tubes, Phillips MCFE 40 W/33) are placed 130 mm apart. A time switch gives a 12 h on/off cycle. The cultures are supported on a shelf 320 mm below the light source and are illuminated until sporulation is induced.

Subculturing

The simplest method of maintenance of living fungi is by serial transfer from staled to fresh solid or liquid media and storage in the most suitable conditions for the individual isolate. Many fungi can be maintained in this way for years by growth on suitable media. Successful maintenance is dependent upon transfer from well-developed parts of the culture, taking care to ensure that contaminants or genetic variants do not replace the original strain. Most fungi can be grown on Potato Carrot Agar (PCA) or Malt Agar (MA). However, others have specified growth requirements. Some dermatophytes survive best on hair (Al-Doory, 1968), some water moulds are best stored in water with the addition of plant material (Goldie-Smith, 1956), and other more sensitive water moulds may require aeration (Clark & Dick, 1974; Webster & Davey, 1976). Such methods are labour intensive and time consuming when large collections are involved.The main disadvantages of frequent transfer are:

• Danger of variation, loss of pathogenicity or other physiological or morphological characteristics.
• Danger of contamination by air-borne spores or mite carried infections.
• Requires constant specialist supervision to ensure that the fungus is not replaced by a contaminant or subcultured from an atypical sector.

• Collections can be kept viable for many years if supervised by a specialist.
• The method is cheap and requires no specialized equipment and for a small collection the time involved is not great.
• Retrieval is very easy.

The time period between transfers varies from fungus to fungus, for some every 2-4 weeks, the majority every 2-4 months, though others may survive for 12 months without transfer.Although Chu (1970) maintained several representatives of forest tree pathogens for 1 year at 5°C most organisms were best transferred after much shorter periods.

Mite infestation prevention

Fungal cultures are susceptible to infestation with mites. These small animals, commonly of the genera Tyroglyphus and Tarsonemus, occur naturally in soil and on almost any organic material. They can be brought into the laboratory on fresh plant material, decaying mouldy products, on shoes, on the bodies of flying insects or even in cultures received from other laboratories. The damage mites cause is twofold. Firstly they eat the cultures, a heavy infestation can completely strip the colonies from an agar plate. Secondly they carry fungal spores and bacteria on and in their bodies and as they move from one culture to another the second culture can be cross inoculated and heavily infected with other fungi and bacteria.The mites commonly found associated with fungal cultures are about 0.25 mm in length. They can be seen by the naked eye only as a tiny white dot almost at the limit of vision, so infestation can easily go undetected. Given favourable conditions of high humidity and temperature they breed rapidly and spread quickly. Many cultures can be infested before they are noticed. Infested cultures have a deteriorated look and this is often the first indication of their presence.General hygiene and preventative precautions are better than having to control an outbreak. All incoming material should be examined when it enters the laboratory and a separate room for checking and processing dirty material is desirable. The sealing of incoming cultures, storage in a refrigerator or some form of screening and quarantine system can be helpful, as it is possible for cultures with only a light infestation at the time of receipt to develop a heavy infestation later.Methods of control used by different workers are various and a combination of precautions seems the answer.HygieneHygiene coupled with quarantine procedures is perhaps the best protection. All work surfaces must be kept clean and cultures protected from aerial and dust contamination for example by storage on protected shelves. The work benches and cupboards should be regularly washed with an acaricide, especially as soon as infestation is suspected. The procedure at CABI BIOSCIENCE is to wash down with an acaricide which is left for sufficient time to have an effect (15 min) and then cleaned off with alcohol. The benches are then repolished with a cloth if desired. The acaricide used at CABI BIOSCIENCE is Actellic 25EC (Fargro Ltd) which has not been found to be noticeably fungicidal. The acaricide must not be allowed to remain on the work surfaces as it is a skin irritant. Plastic gloves and a vapour filter mask should be worn when handling it. As mites appear to become resistant to some chemicals the acaricide should be changed from time to time. When mites are found the affected cultures should be removed immediately and if possible sterilized, and all cultures in the immediate area should be checked and isolated from the rest.

Fumigation

This method is used as a last resort or when moving into new premises and it should be carried out by a licensed specialized company. Where large numbers of important cultures or specimens are involved these items can be fumigated off site in specialized equipment. If this is necessary it is advised that specialist contractors are employed. Such fumigation generally involves the use of chemicals that are toxic to fungi, therefore unaffected cultures should be removed or protected.

Mechanical and chemical barriers

Many physical methods of prevention of infestation and spread of mites have been tried.The culture bottles, tubes or plates can be stood on a platform surrounded by water or oil, or on a surface inside a barrier of petroleum jelly or other sticky material. These methods may be a protection from crawling mites, but do not protect from mites carried by insects or on the hands and clothes of laboratory workers.Particularly useful for universal bottles and cotton wool plugged tubes is the cigarette paper method. In the case of tubes the plug can be pushed down and the tube sealed above the plug without fear of contamination. This method was first described by Snyder and Hansen (1946). The pores of the paper are of a size to allow free passage of air but are too small to allow mites to pass through. Care must be taken to see that a good seal is made, and that the paper is not damaged through handling. It has the advantage that it not only keeps mites out, but it also keeps them in - thus preventing spread of infestation. This method is preferred and used extensively at CABI BIOSCIENCE.

Method

1. Cut cigarette papers in half and sterilize in an oven at 180°C.
2. Stick a cigarette paper onto the universal bottle using copper sulphate gelatin glue.20 g gelatin: 2 g copper sulphate: 100 ml distilled water: 20 g gelatin are dissolved in 100 ml water and then the copper sulphate is added.
3. Burn the excess cigarette paper up to the outer edge of the tube or bottle.

Sealing of culture containers such as Petri dishes and universal bottles with sticky tape such as Sellotape or Scotch tape may reduce penetration but will not act as a complete barrier. Mites eventually find their way through cracks and wrinkles.

Protected storage

The various methods of long term storage of cultures used in culture collections prevent infestation and spread of mites, but are of little use for day to day growth of cultures.Cold storage at 4-8°C definitely reduces the spread of mites, which are almost immobile at this temperature. However, on removal from the refrigerator the mites rapidly become active again. Storage of infested cultures in the deep freeze (< - 20°C) for at least 3 days gives better control. The cultures usually remain viable, whereas the mites are usually killed. The fungus will have to be re-isolated from the original culture as the contaminants introduced by the mites will eventually grow.Covering cultures with mineral oil increases the period of viability of cultures and mites will not penetrate the oil. If they do get into the cultures they will not be able to escape to infest other cultures. Again contamination of the culture may occur due to the growth of spores and bacteria carried by the mites.Cultures stored in silica gel are kept in sealed tubes or in bottles with the caps screwed down so penetration cannot occur. Freeze dried ampoules being completely sealed are impermeable to mites and they cannot penetrate ampoules stored at the ultra-low temperatures of liquid nitrogen.

Hazards encountered during mite prevention procedures

Industrial methylated spirits (IMS) This is highly flammable and of moderate toxicity. Do not wash benches down when there is a naked flame on the bench which you are cleaning. If a spillage occurs mop up with copious amounts of water and ventilate the site of the spillage.Actellic 25ECThis substance is of moderate toxicity and irritating to skin, all contact must be avoided. In the concentration at which it is used (3% v/v: 30 ml of stock to 1 l distilled water) it is much less toxic.Safety data for Actellic 25ECThe active ingredient, pirimphos-methyl, is an organophosphorus compound. Do not use if under medical advice not to work with such compounds. Wear synthetic rubber or PVC gloves when handling the concentrate. Wash splashes of concentrate from skin or eyes immediately. When applying actelic to laboratory surfaces wear suitable protective clothing, laboratory coat, gloves and organic vapour filter mask. When using do not eat, drink or smoke. Ensure adequate ventilation in confined spaces. Do not breath in. If necessary for personal comfort, wear a mask. Wash hands and exposed skin before meals and after work. Take off immediately heavily contaminated clothing. Store in the laboratory. Cover water storage tanks before application. Keep out of reach of children. Ventilate confined spaces thoroughly. Harmful to fish. Do not contaminate ponds, waterways and ditches with the chemical or used container. Keep in original container, tightly closed, in a safe place. Wash out container thoroughly and dispose of safely. Do not re-use container for any purpose.Carry out mite cleaning operations at the end of the working day wherever possible.

References

Al-Doory, Y. (1968) Survival of dermatophyte cultures maintained on hair. Mycologia 60, 720-723.

Chu, D. (1970) Forest pathology, storing of agar slants and cultures. Bi-monthly Research Notes 26, 48.

Clark, C. & Dick, M.W. (1974) Long term storage and viability of aquatic oomycetes. Transactions of the British Mycological Society 63, 611-612.

Clifford, B.C. (1973) Preservation of Puccinia hordei uredospores by lyophilisation of refrigeration and their subsequent germination and infectivity. Cereal Rusts Bulletin 1, 30-34.

Goldie-Smith, E.K. (1956) Maintenance of stock cultures of aquatic fungi. Journal of Elisha Mitchell Scientific Society 72, 158-166.

Pitt, J.I. (1980 ["1979"]) The Genus Penicillium and its teleomorphic state: Eupenillium and Talaromyces. London, New York: Academic Press.

Raper, K.B. & Thom, C. (1949) A Manual of the Penicillia. Williams and Wilkins: Baltimore.

Smith, D. & Onions A.H.S. (1994). The Preservation and Maintenance of Living Fungi. Second edition. IMI Technical Handbooks No. 2, pp 122. Wallingford, UK: CAB INTERNATIONAL.

Snyder W.C. & Hansen H.N. (1946) Control of culture mites by cigarette paper barriers. Mycologia 63, 455-462.

Webster, J. & Davey, R.A. (1976) Simple method for maintaining cultures of Blastocladiella emersonii. Transactions of the British Mycological Society 67, 543-544.

Recommended growth conditions for some fungi