ECONOMIC POTENTIAL: DIVERSIFIED AGRICULTURE MUSHROOM CROP

 

Low Technology Mushroom Cultivation using Agricultural
Wastes as Substrates
Mushrooms are an attractive crop to cultivate in developing countries for many reasons. One of the most
compelling points would be that they are grown on agricultural wastes. It enables us to acquire substrate
materials at low prices or even for free and to conserve our environment by recycling wastes.
Mushrooms can be grown on virtually all types of available wastes.


SUBSTRATE EXAMPLES

Figure 1. Banana and its leaves

 

 

Figure 2. Bean pods

 

 

Figure 3. Maize husks and cobs

 

 

 

Figure 4. Mixture of corn waste

Figure 5. Wheat Straw

Figure 6. Dried cottonseed hulls

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Figure 7. Hard wood chips

 

 

 

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Figure 8. Commercial waste cotton piles

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Figure 9. Various wild grasses

 

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Figure 10. Straw

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Figure 11. Sawdust

 

 

 

According to a recent survey, approximately 200 different wastes are available as mushroom substrates. So, every grower producing mushrooms can make their own best substrate choice from virtually any form of agricultural waste. We must not be at all surprised that the evaluation of all these kinds of different wastes leads us to a renewed appreciation for what is called a waste. Growing mushrooms gives so much satisfaction and produces so much food and income that further use of this practice can result in a great complete contentment of families and villages!

 

Mushroom Cultivation


Mushroom growing house (MGH) xxxx

 

The recommended MGH (shown below) is an 8 x 6m brick and mortar structure, with cement floor, gable thatched roof with jute bags ceiling. The building has screened (wire gauze) windows on all four sides, with 2 windows on one of the longer sides and a door between the windows on the other longer side. Double doors are recommended, with the outer door being a screen door. In certain past instances, resources (money) were provided to acquire bricks, cement, window and door frames and pay the builders. The community was to provide all the other resources necessary to complete the mushroom growing houses. A MGH ready for use was to have the floor covered with river sand up to a depth of 2cm. On occasion, the gabled roof was modified to a flat roof, because the recommended roof type was expensive to put up.

 

 

 

 

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Recommended MGH w/flat roof

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Mushroom growing house made of grass

 

 

At virtually all past sites, the MGH was modified in various ways. At some sites, the structure was made of grass, with a lining of plastic sheeting inside. Generally, the size of the MGH was reduced to 4 x 3m, with a flat roof. At another site, the walls of the mushroom house were made of pole and dagga, with a flat thatched roof. On occasion pre-existing structures were modified into mushroom growing houses. In some cases, traditional kitchens (round huts) were renovated and modified into mushroom houses. In other cases, existing structures in the homestead were partitioned and modified into mushroom houses.

 

 

 

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Traditional Kitchen

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Partitioned Structure

 

Additional work involved in the construction of the MGH included laying of sand for the floor, collection and processing of the substrate and spawning, and watering of the mushroom house at least once a day. Where resources were very limited, the family modified disused structures in the homestead. These included round huts and partitioned structures in the already existing buildings. Resources were then channeled into acquiring the spawn and plastic tubing, which were needed to grow the mushroom. Uses of naturally available materials were demonstrated in MGHs that were constructed utilizing thatch grass, pole and dagga. These materials are readily available, especially after the rainy season, and are normally free. One just has to have the labor to cut and collect.


How to Build a Clean Room in a Simple Way


Upon completion of the mushroom growing house, it is important to construct a cost-efficient, yet effective clean room for handling mushroom cultures. Polythene sheets and sawn timber can be fixed together to form the basis of a sterile clean room. The clean room utilizes a sterile airflow from a basic bench mounted HEPA filter/fan assembly, or laminar flow bench. The size of the clean room is directly correlated to the cubic airflow capacity of the laminar flow bench.


Introduction


What is a clean room?
A hygienically clean, sterile enclosed air space where mushroom mycelium can be isolated from its normal environment in which it must compete with a host of other organisms to survive.

How does it work?
All air in the clean room is ‘cleaned’ by passing it through a High Efficiency Particulate (HEPA) filter to remove the vast majority of airborne particles (living and dead), including: skin, pollen, dust, mould, bacteria, and fungal spores.

Why do mushroom growers need a clean room?
Mushroom spawn is expensive and can be difficult to find. It also does not travel over long distances well. A grower who produces his/her mushroom spawn gains a greater knowledge and understanding of the mushroom growing process.

Overview
In order for a clean room to effectively prevent contamination of mushroom cultures, it must be constructed with minimal mould growing, organic biodegradable materials. High sterile airflow is achieved by placing a large cubic fan capacity in a small area; any airborne contaminants that enter the
clean room are quickly removed by the laminar flow bench. The clean room can be used to inoculate agar plates with mushroom mycelium from culture slants or with tissue samples taken from wild specimens. It can be used to conduct agar-to-agar, agar to grain, grain-to-grain, sawdust to grain, and
sawdust-to-sawdust transfers, using a standard domestic pressure cooker to sterilize the various mediums. The design is intended as a place to teach, learn and experiment with mushroom sterile culture. It is not capable of withstanding the demands of commercial mushroom spawn production.

 

Constructing the clean room


Materials required
- 50 × 2.5m roll clear polythene sheeting (120 gauge, 8 mil, 0.5mm thickness)
- 2 × 1” (50 × 25mm) sawn timber-preferably treated
- 2.5” (60mm) length counter sunk wood screws, tacks or staples
- Silicone caulking sealant
The basic premise is that clear polythene lined wooden frames are made to a standard size. The frames are then joined together to make an enclosed space that is entirely lined by the mould resistant, sterile polythene. Four frames in total, one containing a door constructed in a similar manner. The room is best sited on a concrete base. Other materials that can be used for a sterile floor are linoleum (lino), or butyl rubber sheet.

Constructing the frames
Construct two side frames using 50× 25mm sawn timber to the following dimensions:
2m long × 2m high with a bracing bar positioned at 1m. The two upright lengths of timber should be positioned with the narrow edge facing outwards.
The polythene is cut to length 2.5 × length of the frame. Then folded back and tacked round the rear of one side, thus creating a clear panel. A bead of sealant can be run down the timber to prevent the polythene from ‘flapping’. The two end frames are made to 2 × 1m in size. An entry must be made in
one of the end frames (Fig. 1, 2, 3). The door is best constructed along the same design as the frames. Make sure the door fits well with as few gaps as possible. Household window insulation foam makes a good seal round the doorway.

 

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Figure 1, 2, 3. End frames of clean room

 

 

Joining the frames together


Figure 4. Frame connection

 

To make an airtight seal between the frames, first run a 10mm wide bead of silicone sealant: 10mm from the edge of each frame, all the way round the edge of each frame. The frames should then be positioned at right angles to each other and secured with screws or nails (Fig. 4). After securing the four frames, apply another bead of silicon along all the joins, including the floor. Cut a 2.5 × 1.5m piece of sheeting and secure this over the top of the structure in the same way as the frames were made (a bracing bar may be necessary to add strength).

Equipping the clean room
When creating a sterile environment in the clean room a basic laminar flow bench is required. To be effective at stopping contaminants to mushroom culture the air in the clean room must be passed over a HEPA filter (Fig. 5, 6) once every 60-80 seconds. This is achieved by matching a fan with an output range of 400-500 cubic feet per minute (CFM) with a HEPA filter that is no more than 6 inches (15cm) in depth. Off the face of the filter, an airflow range of 125-150 CFM will give the desired filtering capacity. The cubic air capacity of the clean room is 141 cubic feet. Work surfaces should be made from scratch resistant non-degradable materials such as steel, and positioned at waist height.

 

 

 

Figure 5, 6. Simply made HEPA filter

 

Operation, cleaning and maintenance
The laminar flow bench should be left in continual operation. This will keep the room dry and free from mould growth and other contamination. Cleaning is best carried out one hour before use. A 5% solution of domestic bleach (sodium hydroxide) is used to clean all surfaces. All operatives should wear newly laundered clothes when entering the clean room. General maintenance should be carried out as wear and tear occurs. Any holes in the polythene should be immediately sealed with adhesive tape, and the silicon sealant should be checked regularly (monthly) and remove and replace as necessary. It is recommended that the HEPA filter be changed after 12 months of operation.

Production of Oyster Mushroom Grain Spawn Introduction Mushroom growing using home grown spawn is a process of cellular expansion. Mushroom mycelium is initially grown on a nutrified agar media. This is then used to make grain spawn. The grain spawn is subsequently used to make the final fruiting substrate.

 

 

 

Preparing agar media
The starting point is a mushroom culture (usually in a test tube). This will either be from a clone that you have taken yourself from a mushroom specimen, or purchased from a culture laboratory. Until you have gained experience in cloning and assessing wild mushroom specimens, it is best to use a proven, productive strain from a laboratory.

Growing out your culture
Mushroom mycelium needs nutrition to grow. In other words, it needs something to feed on. Agar agar (a seaweed) contains almost no nutrition, but acts as a gelling agent when mixed with water, so that the mycelium has a flat, solid surface to grow across. A combination of agar agar, water and one or more nutritional substances gives a satisfactory method for growing out healthy mycelium.

 

Figure 7, 8 Tissue culture from a mushroom specimen and transfer to agar culture

 

There are various grades of agar: food-grade for cooking and higher grades for the purpose of culturing. If possible, it is best to use one of the higher grades, as these will contain fewer impurities and probably have better gelling properties than the food-grade. It comes in a dry powder form. There are many different sources of nutrition that can be used in agar media, but probably the most commonly used formula is Malt Extract Agar (MEA). The MEA formula that we use is as follows:


1 liter water
20 grams agar
30 grams barley malt extract*
2 grams nutritional yeast
It is actually called MEYA because of the addition of the yeast. In addition, commercially prepared nutrified agar media can be purchased from mushroom cultivation suppliers; this needs only water adding to it.

Preparing and pouring media
You will need scales and a liter flask that can be used in a pressure cooker. A domestic pressure cooker is ok, but will not form a vacuum when cooled (i.e. it will draw in contaminated air), and therefore should be placed in the clean room environment for cooling. If possible, a proper pressure sterilizer with a pressure gauge is best to use. For pouring the sterilized media, you will also need a supply of isopropyl alcohol and disposable paper towels.

1. Weigh out your ingredients, add the correct amount of water (preferably non-chlorinated, sterile) and mix together in the flask (Fig. 9).

 

2. Use non-absorbent cotton or aluminum foil to stop up the flask. Do not plug it tightly with a stopper, because pressure built up inside the flask while sterilizing will cause it to fly off and the media to boil too furiously (Fig. 10).

 

 

Figure 9. Add distilled water so that the potato starch extract reaches 1L

Figure 10. Aluminum foil-stopped jar

Figure 11. Put the jars in the autoclave and sterilize.

Figure 12. Pour the solution into petri dishes

 

 

Figure 13. Early signs of contamination.

Figure 14. Petri-dish contaminated with green mold.

 

3. Fill your pressure cooker/sterilizer with the correct amount of water (this will vary according to what model you are using-refer to manufacturer’s instructions).

 

4. Place the flask in the cooker. If you are using autoclavable glass petri dishes, these should be sterilized in the cooker at the same time. If you are using disposable dishes, these are usually sterilized already, and the packet should only be opened in the clean room.


5. Place cooker on heat source, bring it up to the correct temperature/pressure-250°F /15 psi and sterilize for 45 minutes. Try to keep the temperature/pressure constant, otherwise the media will not work as well (Fig. 11).


6. Let the pressure reduce to zero-If using a domestic pressure cooker, place in the clean room while it is cooling down.

 

7. In preparation for pouring your media, shower and put on freshly washed clothes. The aim is to reduce as many contaminants as possible, particularly bacteria and lower fungi that stick to you and your clothes and could contaminate your dishes. If possible, put on clothes that have not been exposed to the outside air since being washed. Hair carries a lot of contamination as well so wash it and tie it back (if long), or use a hair net/clean hair covering.

 

8. As agar media solidifies before pouring, clean the work surface in front of your laminar flow cabinet with isopropyl alcohol. Also, clean your own hands with the alcohol and continue to do this at intervals, until you have finished. Pour the dishes that are closest to the filter surface first. The clean air coming off the filter will pick up bacteria, skin fragments etc that are on you, and blow them away from the dishes. Do not put your hands between the filter surface and the open dish, or the air will blow any contaminants from your hands on to the surface of the media.

 

9. Lift the dish lid with one hand and pour the media with the other. Do not let your fingers hang over the rim of the dish or its lid, because skin fragments will cling and cause contamination. While the lid is raised, angle its underside towards the filter so that it only has clean air blowing on to it. At all stages, try not to breathe over the open dishes, as breath is laden with bacteria (Fig. 12).

 

10. When each dish has been poured, leave to cool and solidify. Don’t hang around in the clean room for longer than you have to, as you are just potentially spreading contamination (Fig. 13, 14)!

Inoculating petri dishes
You will need an alcohol lamp for sterilizing the scalpel. If laboratory film (e.g. parafilm) is available for sealing the dishes, there is less chance of contamination. However, if the dishes are left in front of the laminar flow filters until they are colonized with mycelium, film is not essential.

1. Heat the scalpel tip in the alcohol lamp until it is glowing red (Fig. 15). Cool it by dipping it into the centre of the dish that you are about to inoculate.

 

2. Take your culture (having already loosened the lid) and open it without letting your fingers hang inside (Fig. 16). With the scalpel, tease out a wedge of mycelium, lift the lid of the dish and quickly transfer the wedge to the agar, immediately replacing the lid. This is more difficult than it sounds-the mycelial wedges have a huge tendency to cling desperately to the sides of the test tube, while refusing to be picked up by your scalpel! So don’t worry if it’s a bit of a mess to begin with, it just takes practice. The mycelial wedge is placed in the centre of the dish so that growth can radiate away from it.

 

Figure 15. Sterilizing transfer tools and the mouth.

Figure 16. Remove a loopful of agar culture from the of test tube by alcohol lamp test tube.

 

3. If you are using laboratory film, immediately seal up the dish that you have just inoculated. If not,
place it away from the immediate area where you are working, but still in front of the filter.

 

4. Repeat the above steps until there is no mycelium left in the test tube. A standard test tube culture can
usually inoculate 2-3 dishes.

 

5. The dishes should be incubated at 24°C, or as close as possible. Oyster mushroom mycelium is usually quick to start growing; you should see fuzziness, the first sign of growth, within 2-3 days. The mycelium is usually white, and most strains should colonize the dish within 10 days (Fig. 17). If the uncolonized parts of the dish develop areas of different coloring, this most likely indicates some form of contamination, perhaps blue-green mould or yellow slime stuff. In this event, the affected dish should be discarded. If throwing it away is not an option, you can try to leave the contaminant behind, by transferring squares of healthy mushroom mycelium from the affected dish to a new, clean dish. But be warned that moulds in particular produce millions of spores, and any disturbance can cause these to become air-borne and contaminate your healthy mycelium and clean dish, or in a worst-case scenario, your whole clean-room. If this last happens, the only option is to empty the whole room and clean every inch. Clean rooms with a high air flow are less susceptible to this, as any spores are quickly drawn into the filters before they have a chance to settle.

 

 

 

Figure 17. Fuzzy growth on agar culture

Figure 18. Fully colonized pertri dish

 

When your dishes are fully colonized (Fig. 18), they can be used for 3 purposes:
- To inoculate further dishes of agar media. Follow the above procedures for making the media and pouring the dishes, but instead of using a test tube culture, cut squares of roughly 1cm out of a healthy colonized dish and transfer one square to each new dish. 1 petri dish can easily inoculate at least 15 more dishes.

- To inoculate test tubes of agar media so that you have back ups of your strain

- To inoculate sterilized grain, resulting in grain spawn

Preparing Grain Spawn


Grain
Many different types of grain can be used. We use rye grain, which is a relatively soft grain that cooks easily without becoming too sticky or forming clumps.

Spawn containers
Containers can be jars (glass or plastic) or plastic bags. It is essential that they are capable of withstanding pressure sterilization. It is easier to thoroughly mix mycelial wedges from your dishes through jars. Jars should be fairly wide-mouthed, for ease of inoculation, and have tight-fitting lids with an 8mm hole drilled through them. If filter discs are available, these should be fitted over the holes, on the underside of the lids. If not, you can cut out a layer of cardboard the same diameter as the lid, and fit this into the lid. Filter discs should be soaked for at least 1 hour in a 5% household bleach solution; cardboard discs should also be soaked, but for not as long – otherwise they will disintegrate! Soaking helps to dislodge hidden contaminants, which are then killed by the sterilization process. Bags, if used, should be custom-made with filter patches (Fig. 19). If these are not available, it is best to use jars of some sort. Containers without a filter do not work very well, for reasons explained below, and it is difficult to improvise an effective filter for a plastic bag. Also, bags need to be sealed in some way after inoculation - if you have access to a proper heat sealer this is no problem, otherwise something else such as packing tape will have to be used, which is less effective. The purpose of the filter discs/patches is to allow a low level of gaseous exchange. Mushroom mycelium needs a supply of fresh oxygen while it is growing, otherwise it will quickly become anaerobic (without oxygen) and contaminate, regardless of how clean the environment and materials. The filters allow oxygen to enter, without contaminants being drawn in. If filtration is a big problem for you, the only other option (that we can think of!) is to put a small hole in the top of whichever container you are using, and place it over a heat source. Hopefully, convection will keep gases moving upwards and carry potential contaminants away, whilst giving the mycelium access to oxygen.

 

Figure 19. Fully colonized

 


Preparing the grain
1. Weigh out the amount of grain you require. The optimum moisture content of rye grain is roughly 50%, i.e. it will double in weight when cooked. 500g or less of dry grain per container is a good quantity to use when you are transferring mycelium from petri dishes to grain. Any more than this and it becomes difficult to mix the mycelial wedges through properly.

 

 

Figure 20. Sorghum seed

Figure 21. Grain pasteurization in a rice cooker

 

 

2. Grain will always contain hidden contaminants, no matter how fresh it is. Therefore, it needs to be pre-cooked (Fig. 21). Bring water to the boil in a large pan, then add the dry grain. Simmer for around 30 minutes. It is properly cooked when still firm, but soft enough to squash. Any longer than this and the grain kernels will swell so much that they will ‘explode’, which makes spawn more susceptible to contamination. Once you have done this a couple of times, it becomes very easy to determine the correct consistency.

 

 

Figure 22. Jar filled with the sorghum seed

3. Drain the grain. If you are making large quantities of grain spawn, it is fairly easy to improvise a draining container-we have used plastic dustbins with holes drilled in the bottom and sides, and also, sheets of metal mesh rolled into a bin shape and attached on to a base. As long as the water can drain off
rather than collecting at the bottom of the container and forming a big grain mush, there should not be a problem. Mixing the grain at intervals helps this evaporation process. If the grain starts to shrivel, it is drying out too much!


4. If you have access to calcium sulfate (gypsum), the addition of this will help to stop grain kernels clumping together after sterilization. Mix through at a rate of approximately 4g of gypsum to 1kg of dry grain.

 

5. Fill your chosen containers with the grain. If using jars, only fill to 3/4 of their capacity. If using bags, fill to no more than 2/3 of their capacity (Fig. 22). Animal feed scoops are useful for filling containers. You can either use scales to make sure each container is receiving the same amount of grain, or you can see volumetric scoops.


6. Close each container. If using bags fold the excess plastic over the bag. Load the containers into the pressure cookers. If you can space them apart to some extent, sterilization will be more even and efficient. If you have to pack them tightly, remember that it is more difficult for the steam to penetrate the centre of a cooker full of densely-packed containers, and sterilization time may have to be increased.

7. Fill the pressure cooker(s) with the correct amount of water (refer to manufacturer’s instructions) and bring up to sterilization point-250°F/15 psi. We use jars or bags that are filled with 1kg each of cooked grain, and sterilize these for 105 minutes. Smaller quantities of grain in jars will require shorter
sterilization times, e.g. 1L jars with 400g of cooked grain will probably only need 1 hour. If you are using bags and they are very tightly packed, they may need 3 hours. As this depends to some extent on your pressure cooker, you will have to experiment to find the optimum sterilization time.

 

8. When sterilization is complete, let the cooker return to zero pressure (if it does not form a vacuum on cooling, place in the clean room for this). Leave for at least 2 hours so that the jars/bags are cool enough to handle, then unload and place in front of the laminar flow cabinet. It is very important to make sure that they are sufficiently cool, as overly-hot grain will kill the mycelium you introduce. If in doubt,
wait until they are cold.

 

Figure 23. Sorghum seed-based spawn

Inoculating the grain
Each colonized petri dish can inoculate up to 20 cups of grain. The more you add to each container, the greater the speed of colonization. We like to use 1/2 to 1 whole dish to 1 jar containing 1kg grain, as this results in very fast colonization. The quicker the grain becomes fully colonized, the less chance there is of it becoming contaminated.


1. Clean the work surface in front of the laminar flow cabinet with alcohol. As always when working in the clean room, you should have showered and be wearing clean clothes.


2. Take a petri dish and remove sealing film (if used). Heat scalpel tip in the alcohol lamp until it is glowing red. Lift the dish’s lid and cut a criss-cross pattern of 9 or more wedges of mycelium. If you are not using laboratory film to seal the dishes, it is a good idea to leave a 5mm strip of mycelium running round the edge of the dish-this is in case contaminants have entered the dish from round the edge and fallen on to the outermost mycelium.

 

3. Replace the dish’s lid and reheat your scalpel.

 

4. Get your container(s) ready for inoculation-if jars then loosen the lids, if bags then open out the mouth. Be careful at all times not to let your fingers hang inside the container.

 

5. Remove the dish’s lid again, and, with the hot scalpel, pick up 1 or 2 wedges of mycelium. Quickly drop into the waiting container and repeat until the required amount of mycelium has been transferred.

 

6. Reheat the scalpel between each dish-if one of your dishes is contaminated, this will prevent the contamination from spreading across all your containers.

 

7. If you are using jars, then they need to be thoroughly shaken, so that the mycelium travels through the grain, resulting in even colonization. We find that a rolling and ‘see-sawing’ motion works well. Try not to let too many wedges stick to the sides of the jar; this again improves with practice.

 

8. If you are using bags, try to trap some clean air in each bag before sealing. Do this by holding the bag open with its mouth pointing toward the laminar filter, and then sealing. The trapped air helps the mycelium to colonize the bags quickly and healthily. Once sealed, each bag should be agitated so that the mycelial wedges move through the grain.


9. Incubate the containers at 24°C/75°F, or as close as possible. You should see some signs of growth within 3 days. After 7 days, shake the containers to distribute the colonized grain kernels evenly. The grain is fully colonized when it is completely white or off-white. This is the time when it is at its most
vigorous, so use it as soon as possible! If it is left to grow further, it will quickly over-incubate, forming a solid lump that is incredibly hard to break up (Fig. 23).


Your grain spawn can now be used for 2 purposes.
- To inoculate further containers of grain.
1 jar of grain can inoculate 10 times its own mass, so it is very easy to quickly turn small amounts of spawn into large amounts. As always, the more spawn you add, the faster the grain will colonize. (However, when inoculating a container of grain, beware of adding more than 20% of its mass, as growing mycelium produces heat and this will cause the grain to heat up too much and contaminate). Note that when using grain spawn from a jar, it must be broken up first. If it is a glass jar, do not bang it against your hand in case of breakage and injury-use something springy or semi-hard such as a thick padding of towels on a solid surface. Spawn in bags must also be broken up, but this is easily done by squeezing and shaking.
- To inoculate the final bulk substrate, from which mushrooms will eventually be fruited.

 

 

Figure 24. Grain spawn colonizing sawdust

Figure 25. Grain spawn ready to colonize wheat straw

 

 

Figure 27. Oyster mushroom on heat straw column

 

Growing conditions
Moisture and temperature are important factors that must be monitored in the MGH. River-sand on the floor of the MGH is used to maintain high humidity in the MGH. Clean water should be periodically sprayed on the mushroom growing bags, especially during fruiting. Farmers who have readily available water can flood the cement floor, without the sand. In the thatch grass MGH, the walls can be lined with plastic sheeting to prevent excessive water loss through the grass walls. With the soil rammed floors, the floor can be covered with plastic before river sand was added. This prevents excessive seepage into the ground. Where resources are limited, instead of buying a new hand sprayer for the MGH, new traditional hand brooms are recommended. These can be dipped in clean water and used to spray the bags. Black or white plastic tubing can be used as mushroom growing bags. Some farmers prefer the white tubing as this allow them to monitor activities in the mushroom bag. Others prefer the black pockets because they reduced the need to close up the MGH during the first 4 weeks when no light is required in the MGH.

Harvesting
Mushroom pinheads generally appear within 4-6 weeks after the substrate has been fully colonized by the mushroom mycelium. The bags are periodically sprayed with clean water to avoid drying of the mushroom pinheads. Within 3-5 days the mushroom fruiting bodies will be ready for harvesting.

Conclusion
Mushroom cultivation at all stages is cost-efficient and requires very little in the way of technology. The fact that agricultural waste makes the most ideal form of substrate means that it is readily available and easy to obtain. Mushroom growing operations can be established virtually anywhere and with the right initial supervision anyone can be taught this valuable skill. Mushroom cultivation is an ideal form low-cost, low-technology capital generation.