A Detailed Guide to Edible Mushroom Spawn Storage and Top-Dressing Methods

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 grown vigorously and covered 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, transferring it to a fresh tube once 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 straw mushroom spawn, the spawn of other edible mushrooms can all be stored by this method.

2. Liquid paraffin storage method. Take chemically pure liquid paraffin (required to be free of water and not moldy), put it in a triangular flask, add a cotton plug and wrap with paper, sterilize at 1 kg/sq cm pressure for 1 hour, then place it in a 40°C constant-temperature incubator for several days to evaporate the water in it, until the paraffin oil is completely transparent. Transfer the treated paraffin oil onto a blank slant and culture at 28°C-30°C for 2-3 days; use it only after confirming that no contaminating microorganisms have grown. Then, using aseptic technique, inject the liquid paraffin into the slant test tube to be stored. The injection amount should preferably be 1-1.5 cm above the slant medium. Insert a rubber stopper, seal the mouth with solid paraffin, and store it upright in a low-temperature dry place. The storage period is more than 1 year, and at low temperature the preservation time can be extended further.

3. Sand-soil tube storage method. Take river sand, soak and wash it in water several times, pass it through a 60-mesh sieve to remove coarse particles, then soak it in 10% hydrochloric acid for 2-4 hours to remove organic matter, then rinse with water until the pH value reaches neutral, and dry it for later use. At the same time, take poor soil or garden soil, soak it in water to make it neutral, remove the supernatant after settling, dry it, grind it fine, and pass it through a 100-mesh sieve. Mix the treated sand and soil in a ratio of (2-4):1, use a magnet to draw out the iron material, then dispense into small test tubes or ampoules, 0.5-2 grams per tube, plug with cotton, 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 it passes. Take the slant spawn that has already formed spores, inject 3-5 ml of sterile water under aseptic conditions, 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 tube 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 avoid spore germination. Seal the prepared sand-soil tube with paraffin; it can be preserved for 2-10 years at low temperature.

4. Filter paper strip storage method. Take white filter paper (for collecting dark-colored spores) or black filter paper (for collecting white spores), cut it into small strips of 4 cm x 0.8 cm, lay them flat in a petri 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%, and then store at low temperature.

5. Natural substrate storage method. 1) Wheat grain storage method. Take clean wheat free of shriveled grains and impurities, soak it 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 them 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 them into test tubes, 2-3 grams per tube, then clean the test tubes, plug with cotton, sterilize (1.5 kg/sq cm, 2 hours), and set aside after passing the sterility check. After the test tube substrate cools, inoculate, culture at an appropriate temperature, and after the mycelium has covered the substrate, coat the cotton plug with paraffin and store at low temperature. Transfer once about every 2 years. 2) Bran koji preservation method. Take fresh bran and pass it through a 60-mesh sieve to remove coarse particles. Mix the bran and tap water at 1:1, put it into small test tubes, filling each tube to about 1/3 of its height, add a cotton plug, wrap with paper, and autoclave (1.5 kg/sq cm, 30 minutes). Set aside after passing the sterility check. Transfer vigorous spawn grown on slant medium into sterile bran koji tubes. During transfer, be careful to mash 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 has covered the bran. 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% ml of distilled water, stir evenly, dispense into test tubes, 5-10 ml per tube, sterilize (1 kg/sq cm, 30 minutes), and set aside after passing the sterility check. Inoculate the spawn to be preserved into potato glucose 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 a test tube of physiological saline that has passed inspection, insert a sterile rubber stopper, coat and seal with paraffin, and store at room temperature or low temperature.

7. Freeze vacuum drying method. Suspend the already cultured, abundantly grown cells or spores in a bacterial suspension made from sterilized serum, egg white, or skim milk. Under aseptic operation, dispense the suspension into sterilized glass ampoules, 0.3-0.5 ml per tube, then connect with a freeze-drying apparatus using a pressure-resistant rubber tube. Place the ampoules in a freezing bath and freeze rapidly at -30°C to -40°C, then vacuum-dry in the frozen state, and seal the ampoules by melting under vacuum. Store at -20°C low temperature; generally it can be preserved for more than 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. Next, prepare ampoules; add 0.8 ml of a 10% (volume ratio) glycerol distilled water solution as a cryoprotectant to each bottle, plug with cotton and sterilize (1 kg/sq cm, 5 minutes). After the sterility check, 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 cool slowly at a rate of 1°C per minute, so that the preserved material gradually and evenly freezes, until -35°C; after that, the freezing rate no longer needs to be controlled. After the ampoules are frozen, immediately put 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.