Low cost/ low tech mushroom cultivation on agricultural waste products

The project I am working on currently was an idea of a friend (Tatek) and colleague at the University of Valladolid. In Cooperation with the Ethiopian environment and forest research institute, which focusses on the exploration and use of NTFP (Non Timer Forest Products), we are looking for possibilities of cultivating edible mushrooms on agricultural waste products. In future, this project shall give oppurtunities to the farmers of ethiopia to integrate this method as a part of circular economy. This project is not directly scientific, but rather focusses on mastering different techniques of mushroom cultivation, in order to develope adequate procedures adapted to the localities and infrastructure of the rural areas in Ethiopia.

state of the art

To give a short insight into our current status, I will shortly explain our key approaches and obstacles:
Since we have a university lab for this purpose, we are working very equipped on most steps of the expansion of mycelial mass. We are currently using Agar-to-grain and grain-to-substrate methods. We have an autoclave, a clean bench, incubaters and a fridge (+cutting tools, bunsen burner etc.).

We are currently working with the species Pleurotus ostreatus, Pleurotus eryngii, Flammulina velutipes, Trametes versicolor, Chondrostereum purpureum, Agaricus bisporus, Agaricus brunnescens and Lentinula edodes. The focus is on edible species over medicinal ones.

For Pleurotus eryngii, Flammulina velutipes, Agaricus bisporus and Agaricus brunnescens we used isolation techniques (with hydrogen peroxide) on specimen bought at the supermarket. The Isolated Agaricus strains seem to suffer from senescence or contamination (I will add pictures).

We are using polypropylen bags and reused preserving jars for the spawn and final substrates. Our fruiting chamber is simply a growing tent.

So far we were succesful of fructificating P. ostreatus and Lentinula edodes for 1-2 flushes. Our contamintation rates are also quite low (of course due to professional equippment).

Spawn and substrates of the other species (exept Agaricus strains) are in progress.

Important challenges to work on

Since we are using several species, substrates and techniques simultaneously, I will specially share with you all the parts of this project that are most challenging, to avoid overloading this thread. If you have any questions, you are very welcome to ask!

Main challenges are:

The fructification without the use of electricity
As most of you most likely already expercienced, there are several factors regarding the environment of a growing chamber, to fullfil the demands of our fungal friends. The maintainance of adequate relative air humidity, air exchanges (oxygen) and light necessary for the primordial and fruiting phase without the use of electricity (in worst case with low electricity needs) are one of our main objectives. So far we are using the growing tent and an aerosol for humidity. It is known that aerosoles are favoring bacterial contamination and the danger of suffocation of primordial formations due to excessive application. Furthermore, we are winding up seeing the water landing on the walls and running down to the floor and our blocks drying out after the first flush. The fruiting bodies of P. ostreatus also seemed pale, probably as a result of insufficient light. If you are aware of techniques that could help us solve this problem, please let us know! :slight_smile:

The sterilization of substrates without autoclave or pressure cooker

This step of the procedure has not been developed yet (we focused on the fructification), but should be elaborated soon (parallel to our other trials). Techniques to try are a three day sequence of boiling the substrate for 30 min each days, giving the spores inside the substrate time to germinate, before heating (and hopefully destroying them) the following day. Another interesting technique uses a several day yeast fermentation to remove unwanted microorganisms. I will share updates on the progress of these methods!
If you are aware of further sterilization techniques, we are happy to hear about it

PS: a picture of our cloned Agaricus strain that looks very sick. It looks moreless like a fluffy mycelium (not sure if visible on picture) with a dark bottom part (almost like a contaminant), growing extremely slow. The hyphae started shooting out of the extracted flesh, which supports our hope that it is not a contaminant. But since no one of us has seen proper Agaricus mycelium before, we are not sure if it is supposed to look like that. Does anyone here know more? :slight_smile:

I will share some more results in the following weeks, keeping you updated!

We are happy about any kind of feedback, question or criticism!


Hi @timst! So good to see some action in here!

To the fructification part: How about soaking them in water (maybe laced with some H2O2) after each flush? That should reduce the need for moisture in the air. Also spraying with spray bottles is possible but tiresome, since it has to be done every few hours.

Regarding the sterilization, I believe the boiling over three consecutive days is a great idea. Tydallisation works quite well actually. But a yeast fermentation would have the huge advantage of reducing the energy input. Another method for bulk substrates could be soaking in a solution of cal and water to increase the pH (this is only possible with some species, Pleurotus for example).

I don’t know about agaricus honestly but this looks like normal mycelium to me. The colour wouldn’t spread in the shape of the other mycelium like that if it were contamination in my opinion. It’s possible it grows slowly.

Keep it up!

Hey @phylanx !

Thanks a lot for your ideas. On the next fructification we will try soaking the block in water and H2O2 and see what happens! I´ll keep you updated about it.

For the sterilization method using cal and water, is it important to soak the bulk substrate in the solution or is it possible to add cal right away when mixing the substrate? And how much cal should be added approximately to avoid ending up with a brick? :smiley:

And thanks for your evaluation of our Agaricus strain. The book “Growing gourmet and Medicinal mushrooms” gives a clue about the senescence of store bought Agaricus strains. Apparently the industry is trying to protect their valuable strains from being “stolen”…

I´ll keep you updated about the progress!


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Update on the progress:

We are unsure about the appearance of our Flavulina velutipes culture. With age it seems to get a little bit grainy on the surface (looking a bit like white conidia), which has been described similar in the literature. Before we try to let it fructificate and spoil our growing chamber with potential conidiospores, I want to ask if anyone here has experience with this culture and it´s mycelial characteristics?

Here is a picture of the Flammulina velutipes culture on millet. In the upper part you can see the grain-like structures. We also checked one sample under the microscope, with the basic approach to look for clamp connections, which would at least tell us if we are having a Basidiomycete here and higher the chances of it being the correct species.

Our lack of experience on that came with loads of uncertainty, but you can see yourself. My somewhat biased eye can make out some septae and something looking a bit like clamp connections before them. I would be happy about every advice regarding this topic :slight_smile:

Furthermore, we put Pleurotus ostreatus and Chondrostereum purpureum inside our fructification chamber. So far, the only reaction observed was a coloration around the holes that we cut into the plastic:

This is our Pleurotus block of oak substrate mixed with approximately 40% coffee ground. The colonization happened extremely fast, though there might have been a lack of oxygen on the bottom part. This picture was taken 5 days after the holes were cut. Does anyway know why the openings turned dark and yellowish? Is it a reaction to oxygen, or maybe excessive watering?

Thanks in advance!

I’m not sure I can help you out with your questions, but I can tell you that your grain jar is too full! You want to fill it only to a maximum of ~2/3rds in order to have space to shake it. As you can see, your culture is growing down from the top and with more space you could shake it up at around 20% colonization and then it will colonize faster.

The change in color around the wholes is worth monitoring. Are these wholes pointed to the side? They shouldnt be in contact with the floor. 50% coffee makes contamination more probable. I wouldnt go beyond 20%, especially with your low-tech approach.

Ah, the velutipes can look like that if I remember correctly. If you want to be sure, let it colonize to 100% and leave it a few days longer. If it’s a mold, it will make conidia that are typically colored (black or green being most common). I don’t think it’s that. Another possibility is cobweb mold, which would be really bad and could be, but I can’t really help you how to be sure it’s not that as I don’t have much experience with that one.

Good luck!

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Hey @phylanx,

thanks for the advice! We wanted to go for a bottle fruiting and maybe got a bit greedy on using all the space possible for substrates! We´ll try to give it some more room on the next round.

The block is laying on a box turned upside down, the holes are not touching the floor! Do you know why coffee is increasing the probability of contaminations? Is it due to the high nutritional values?
And do you think it would be a good idea to open the hole bag exposing higher mycelial surface area to the fresh air? Or should we be more patient?

Thanks a lot!

Hey there, I just stumbled over this.

Coffee has lots of nitrogen, I believe that’s the main reason why it increases contamination risk. I imagine it has some phosphor and trace minerals as well, not sure though.
I wouldn’t open the bag. There is a technique where you make slits in the bag and cover them with a certain type of tape which lets air pass through. I don’t think it’s an option for your setting, you will have to try out different things and get some ideas on how to do this low-tech approach in your area and conditions.