Pest and Disease Control for Edible Fungi and Spawn Storage Techniques

1. Low-temperature periodic transfer storage method. Inoculate the spawn to be stored onto a suitable slant medium and culture at an appropriate temperature. When the mycelium has vigorously grown to cover the slant, remove it and store it in a dry place at 3°C-5°C low temperature or in a 4°C refrigerator or freezer. Transfer to a fresh tube every 4-6 months; the specific interval should be determined according to the characteristics of the spawn. During storage, note that the ambient temperature must not be too high, to prevent mold from entering the tube through the cotton plug. Therefore, if a cotton plug is used, it can be wrapped with clean sulfate paper or kraft paper, which reduces the chance of contamination and also prevents the medium from drying out. Except for the spawn of straw mushrooms, the spawn of other edible fungi can all be stored by this method.

2. Liquid paraffin storage method. Take chemically pure liquid paraffin (requiring no moisture content and no mold) and place it in a triangular flask, plug with a cotton plug and wrap with paper, sterilize at 1 kg/sq cm pressure for 1 hour, then place in a 40°C constant-temperature incubator for several days to evaporate the moisture therein, until the paraffin oil is completely transparent. Transfer the prepared paraffin oil onto a blank slant and culture at 28°C-30°C for 2-3 days; use only after confirming no contaminant microorganisms have grown. Then, using aseptic technique, inject the liquid paraffin into the slant tubes to be stored. The injection volume should preferably be 1-1.5 cm above the slant medium. Insert a rubber stopper, seal the mouth with solid paraffin, and store upright in a low-temperature dry place. The storage period is over 1 year, and at low temperature the preservation time can be further extended.

3. Sand-soil tube storage method. Take river sand, soak and wash it in water several times, pass through a 60-mesh sieve to remove coarse particles, then soak in 10% hydrochloric acid for 2-4 hours to remove organic matter, then rinse with water until the pH reaches neutral, and dry for later use. At the same time, take poor soil or garden soil, soak it in water to make it neutral, after settling remove the supernatant, dry and grind fine, and pass through a 100-mesh sieve. Mix the prepared sand and soil in a ratio of (2-4):1, use a magnet to draw out the iron material, then distribute into small test tubes or ampoules, with 0.5-2 grams per tube, plug with cotton plugs, wrap with paper and sterilize (1.5 kg/sq cm, 1 hour), then dry-heat sterilize (160°C, 2-3 hours) 1-2 times, carry out a sterility test, and use after passing. Take the slant spawn that has already formed spores, and under aseptic conditions inject 3-5 milliliters of sterile water, scrape the mycelium to make a spore suspension, then use a sterile pipette to draw the bacterial liquid and drip it into the sand-soil tube until the sand-soil is soaked through. Place the inoculated sand-soil tubes into a vacuum desiccator containing a desiccant, connect a vacuum pump and evacuate for several hours, until the sand-soil is dry. The vacuum drying operation must be completed within 48 hours after the spores are inoculated, to prevent the spores from germinating. Seal the prepared sand-soil tubes with paraffin; they can be preserved at low temperature for 2-10 years.

4. Filter paper piece storage method. Take white filter paper (for collecting dark-colored spores) or black filter paper (for collecting white spores), cut into small strips of 4 cm × 0.8 cm, lay them flat in a culture dish, wrap with paper and sterilize (1 kg/sq cm, 30 minutes). Use the hook suspension method to collect spores, letting the spores fall onto the filter paper strips. Place the filter paper strips carrying the spores into storage test tubes, then place the storage test tubes into a desiccator for 1-2 days to remove the moisture from the filter paper, bringing the moisture content of the filter paper to about 2%, then store at low temperature.

5. Natural substrate storage method. ① Wheat grain storage method. Take wheat without shriveled grains or impurities, wash it clean, soak for 12-15 hours, add water and boil for 15 minutes, continue hot soaking for 15 minutes, so that the wheat grains swell but do not burst, drain the water, spread out to air-dry, and bring the moisture content of the wheat grains to about 25%. Mix calcium carbonate and gypsum into the cooked wheat grains (ratio of wheat grains, calcium carbonate, and gypsum is 10 kg:133 g:33 g). After mixing evenly, put into test tubes, 2-3 grams per tube, then clean the test tubes, plug with cotton plugs, sterilize (1.5 kg/sq cm, 2 hours), and use after passing a sterility test. After the tube substrate cools, inoculate, culture at an appropriate temperature, and after the mycelium grows over the substrate, coat and seal the cotton plug with paraffin, and place at low temperature for storage. Transfer once every 2 years or so. ② Bran koji preservation method. Take fresh bran and pass through a 60-mesh sieve to remove coarse particles. Mix the bran and tap water at 1:1, put into small test tubes, filling each tube to about 1/3 of its height, add cotton plugs and wrap with paper, autoclave (1.5 kg/sq cm, 30 minutes), and use after passing a sterility test. Transfer robust spawn growing on slant medium into sterile bran koji tubes. When transferring, be careful to stir the bran in the small test tubes as evenly as possible, so that it is in a loose state, culture at an appropriate temperature until the mycelium grows over the bran, then place the small bran koji tubes in a desiccator and store at low temperature or an appropriate temperature.

6. Physiological saline storage method. Take 0.7-0.9 grams of pure sodium chloride, put it into 100% milliliters of distilled water, stir evenly and distribute into test tubes, 5-10 milliliters per tube, sterilize (1 kg/sq cm, 30 minutes), and use after passing a sterility test. Inoculate the spawn to be preserved into potato dextrose liquid medium and culture with shaking at an appropriate temperature for 5-7 days. Using aseptic technique, draw a small amount of the cultured spawn and inject it into the test tubes of physiological saline that have passed inspection, insert sterile rubber stoppers, seal with paraffin, and store at room temperature or low temperature.

7. Freeze vacuum drying method. Suspend the already cultured, abundantly grown mycelium or spores in a bacterial suspension made from sterilized serum, egg white, or skim milk. Using aseptic technique, distribute the suspension into sterilized glass ampoules, 0.3-0.5 milliliters per tube, then connect to a freeze-drying apparatus with a pressure-resistant rubber tube, place the ampoules in a freezing tank and rapidly freeze at -30°C to -40°C, then vacuum-dry in the frozen state, and seal the ampoules by melting under vacuum, and preserve at -20°C low temperature. Generally, they can be preserved for over 10 years, but the cost is relatively high.

8. Liquid nitrogen ultra-low temperature preservation method. First, prepare the spawn to be preserved into a bacterial suspension for later use. Second, prepare ampoules; add 0.8 milliliters of a 10% (volume ratio) glycerol distilled water solution as a cryoprotectant to each bottle, plug with a cotton plug and sterilize (1 kg/sq cm, 5 minutes). After a sterility test, inoculate the spawn to be stored, seal the bottle mouth by flame melting, and check for air leakage. Place the sealed ampoules in a freezer and slowly lower the temperature at a rate of 1°C per minute, so that the preserved material gradually and evenly freezes, until -35°C; thereafter the freezing rate need not be controlled. After the ampoules are frozen, immediately place them into a liquid nitrogen tank and preserve at -150°C to -196°C low temperature. This method is used by only a few research institutes.