Cultivation tests on spent coffee grounds

Around end of last year I have collected for some days the spent coffee grounds from my workplace. I sterilized it in pressure cooker in glass jars and then inoculated it with spawn either bought from Mushroombox.co.uk or what I collected in the wild.

I have read a lot about recycling coffee in this way, but I wanted to see with my own eyes.

I have tried several species, mainly oysters: Pleurotus citrinopileatus (yellow oyster), Pleurotus ostreatus (grey oyster), Pleurotus eryngii (King oyster), Lentinula edodes (shiitake), Hypsizygus ulmarius (elm mushroom), Pleurotus djamor (pink oyster), Hericium erinaceus (lion's mane). With the latter two I had no success, but I will give it another try later on.

All the rest managed to form pinheads (small pre-shrooms) and they mostly developed to elongated fruit bodies. The elongation was due to the fact that the basic setup I used was not meant to control levels of O2 and CO2 and to provide regular fresh air exchange, all crucial at time of fruit body development.
After the moving is complete back to Hungary I hope to renew the experiments on more species and with a better suited setup.

Spawn run (while the shroom conquers the substrate) took something between 30-60 days, which is pretty long for so little quantity, but they were basically forgotten there in the dark of my shelves on room temperature and without any additional substrate material, coffe grounds are pretty dense to be simply overrun.

After this incubation period the jars looked like this:

I opened them up and placed into nursery boxes with a simple humidity and temperature meter:

 The boxes were covered, and the top opening was covered with a garden foil:

I tried to keep the humidty above 85-90%, by keeping the bottom of the box constantly moist and removed the cover for air exchange in the mornings and in the evenings. These were all but sufficient for the proper mushroom formation, but at least I got some results.

The grey and yellow oyster and the shiitake did well on the substrate:

Pleurotus ostreatus

Pleurotus ostreatus

Lentinula edodes

Pleurotus citrinopileatus

The P. eryngii took its time compared to the others: until the king oyster decided to form some shy fruit bodies, the yellow oyster gave 3 waves of crop...

Pleurotus eryngii

 My wild collection of Pleurotus was extremely desorientated. No matter what I did, it kept forming these impossible shaped fruit bodies. I think it really missed the fresh air movement that it got used to in nature:

wild Pleurotus strain

I am not very familiar with the elm mushroom. It formed plenty of pins, out of which only some developed into bigger fruit bodies, strongly elongated and deformed:

Hypsizygus ulmarius

 I made some checks after disposal of the test glasses. I found that run within the substrate was only partial in most cases, as visible on the photos below:

It is mostly due to the density of the coffee. Most larger scale projects mix at least wood shavings to the coffee, I might give it a try next time.

Spring check up on wild shrooms II

I have checked upon the Aesculus hippocastanum trees (horse-chestnut) next to the road, which nurtured Flammulina velutiped during the cold winter months in their long verticular cracks.

The shrooms were still there, both old and young. I collected 350-350g from each (old and young) for the dyepot, leaving still enough on the trees. I also planned to take sample in the strain collection, but due to the moving (from Netherlands back to home) that was not feasible any longer, but I anyway had an earlier collection from these trees and they looked okay.

And this is how they look after some frost period:

Spring check up on wild shrooms I

I have restarted recently my regular cycling trips in the neighbouring forests and pastures after the winter period. The shrooms are still sleeping, apart from some newgrown Trametes hirsuta and versicolor (brown variety) I havent found yet new crop.

First pic: T. hirsuta, other three: T.versicolor brown variety.

I have also found some leftovers from the autumn: Scleroderma citrinum, Piptoporus betulinus, and some gorgeous Calvatia gigantea, that was the best surprise of the day, even the week.

Scleroderma citrinum

Piptoporus betulinus

Calvatia gigantea

The Calvatias, some 4-5 of them were sitting in a row after each other next to the small hump of the road. On the other side of the hump a local farmer collected his nitrogen rich wastes, which made the C.gigantea's appareance logical. This also meant that they were there all along autumn, I just didnt get to see them due to the dense vegetation, though I was regularly checking on this hump for Boletes. I could not resist to take one of these beautiful babies with me for my strain collection and for the dyepot, I am so looking forward to the results!

Dyeing with dried Boletus edulis

This is one of the final results of the dyeing fever that took on me two weekends before.
I took dried Boletus edulis and tested several methods on half skeins of Regina Classic 002 wool after scouring and soaking for overnight. All skeins were dyed in the same dyebath in a consecutive manner.

I really love their welcoming warm, shiny colors. I am bringing it tomorrow to my workplace where a collegue will hopefully be willing to knit a nice hat out of them ( I cant knit :( ).
update: a hat is ready :)!

For the mordanting I used standard alum and a home made iron/copper mordant bath: I placed some 8-10 pieces of eurocents ranging from 1 till 20 plus 4-5 yellowish screws, and placed yarns into it for cold soaking for overnight. It took a while until the cover dissolved from the screws and the first yarns with the overnight cold mordanting method were more copper mordanted, while the ones I simmered thereafter were more affected by the iron screws, and acted more iron-mordanted.

From left to right:

• 1.st bath: unmordanted: warm golden yellow.

• 1.st afterbath: cold mordanted with alum for overnight. In the photo is not apparent but this is a stronger and brighter color as the unmordanted version: strong warm golden yellow.

• 2nd afterbath: I poured some 2-3 tablespoon of ammonia (it is a lot compared to the dye pot which is 2-3 liters, I manage to boost pH to 10 normally with 1-2 teespoons for same volume of liquid from pH 6-7) and placed iron/copper cold mordanted wool in it. It is grey.

• 3rd afterbath: I placed iron/copper simmered (30-60min) wool in it. It turned out be a warm greyish-brown. I think either the ammonia was mostly soaked up by the previous yarn or my iron/copper dyepot slowly turned into the direction of iron as the screws started to release their iron content, or there is a difference in cold and hot mordanting in this way - or a combination of any of these three :).

Start-up with dyeing - Part III. Sampling

I have continued to test wild mushrooms for color in the way described in the previous post:

1. After collecting the mushrooms, prepared a dyebath from them by simmering for ca 30-40 minutes, and then placed the liquid in the fridge for about 10C. And they stayed there unfortunately for couple of month. Those with a pH around 6.3 looked and smelled okay, those above pH7 had either bacterial or mold contam and smelled most of the time.


2. I have scoured the yarn (wool, Regina, color 002) by simmering for half an hour or so and then mordanted a third of it in alum, and another third in iron. It was the same mordant bath as in the previous post --> the yarns are still "over"mordanted, by using too much of the mordant (iron especially, so iron results I just disregard for these experiments).


3. After mordanting I prepared the test swatches (containing an unmordanted, an alum and an iron mordanted piece) and placed them by three into the hand warm dyebath, and simmered them for 60-120 minutes, depends. Unfortunately couple of time I didnt take enough care and the bath began boiling. I dont have a water temperature meter, but I plan to get one for the next experiments.


4. After the dyebath I washed and placed to dry one of the test swatches. Another one I put into warm water, with pH 4.5 (modified by white vinegar) and the third one in pH 10, modified by ammonia. These I simmered for another half an hour.

Below are the results (again, photomashine still sucks).

From left to right:

• 3 unmordanted samples, with acid, normal and alkaline pH, then
• 3 alum mordanted and thereafter
• 3 iron mordanted, pH row the same.

Oh, and yes I used to have photos of from the time I collected the fresh mushrooms for identification purposes, but my laptop got recently stolen (from the backpack of the car, in front of a police station...) and there it went all my photos from earlier times, including the most uptodate version of my strain database :((((. I am still crying when I am thinking of it. Learning: backup backup backup your files.

Calvatia excipuliformis

I used the already ripen (and thus dark) sporemass of one specimen.

It gave a rather pale goldenish in acidic range, and pale green on higher pH levels. No significant difference between unmordanted and alum mordanted yarns, but I havent tested light and washfastness yet.

It seems as an interesting candidate for dying, provided you are not craving for some very striking colors.

They say it is saprotroph, so I might give a try to its cultivation later on (especially that it is edible when young)

  

Panaeolus antillarium

I found this one growing on the top of a heap of sheep poo, on an anotherwise nice pasture. I am pretty sure it is Panaeolus and there is some likeliness that it is antillarium... I collected very cautiosly some 5-6 of them on their long stems. The gills were pretty dark already, they were however not very old mushrooms.

On the photos you might see some difference, but as I am now looking at the test swatches next to me, they actually all look the same pale beige. It is a dye dud suspect, in fact.

I might try later with older specimens.

Scleroderma citrinum

It was ca. half a handful of ripen specimen with greyish-brown sporemass.

It give a strong light brown, especially with alum and without post-dyeing simmering in water with modified pH. As will be seen for a next post, I managed to keep dyeing with this dyebath, so S.citrinum is a good candidate.

 Too bad it is mycorrhizal and cant be cultivated on its own. These ones grew around Castanea sativa (sweet chestnut) mixed with pine and silver birch, in moss, in larger groups.

Some of them hosted a parasitic Bolete, with the creative name Boletus parasiticus. (those parasited Scleroderma were not used for the experiment, though would have been interesting as their interior is at times differently colored than normal Scleroderma) 

Xerocomus armeniacus

With Xerocomus and Boletus species one must be very careful with identification.
This one grew under an oak tree next to the road (and exclusively under that one tree, nowehere else was I able to find this species), and it kept producing new fruitbodies every couple of weeks throughout the summer-autumn period. They were mostly present with blue Russulas, the Xerocomus being at the bottom of the small hump where the water flows and is more moist and the Russula up on the hump with about a meter.
I was always eager to find them, mostly not even picked just awed and photoed. They had really pretty velvety apricot-reddish caps, were not growing too big, while flesh and stem bright yellow, both staining blue when touched. Based on this my best guess became X. armeniacus. Unfortunately they were prone to be attacked very early by a mold, that was even quicker than maggots...

I made two separate dyebath following the same principles from two separate collections from the same place (specimen not too old either case). If you check below, it is stunning how same the results are.

Alum gave the best colors (pale green). Interestingly, strongest colors came on the test swatches simmered in pH 4.5 water after the actual dyebath, both on unmordanted and on alum mordanted swatches.

Test nr.1.

Test nr.2.

Xerocomus communis

These lads were growing at a number of places, in the forest as well as in the middle of the city on alleys. I think they were closest to X. communis, as had small size (up to 5-10 cm), cap yellowish with cracks that are often reddish, red dots in the base of the stem and very very often full with maggots, and is blueing upon touch.

Used specimen were not too old, not too young average shrooms, and they gave strong light green on alum after acidic afterbath and strong light brown on higher pH ranges with or without afterbath. And best of all I could keep on using the dyebath thereafter for a number of yarns :).

As all Xerocomus species, it is mycorrhizal, so no hope for a small home production for a while.

Start-up with dyeing - Part II. Sampling

The very first test swatches contained also cotton and acryl pieces of yarn, but apart from soaking up the precious dyewater, they werent doing anything at all (at that time I didnt know yet that cotton needs a pretreatment with washing soda), so I reduced a sampling to wool, unmordanted, mordanted with alum and iron, respectively. The mordants I bought from Ebay from ForestFibres, who also sell undyed yarns. But for the experiments I used white dyed wool from Adriafil (Regina), which I could buy next corner. I compared results of scoured (prewashed in soap) and unscoured yarns, but didnt see significant difference. Anyway to make sure to get rid off the chemicals used for commercial yarns I decided to scour the yarns for normal dyeing.

Here is a test swatch I used:

It is now apparent to me (especially if I look at how brown the iron mordanted swatch is) that I awfully overused alum and iron, I used over a tablespoon of each into the mordanting water of the samples, while later on I saw that even a teaspoon might be enough for to mordant whole yarns.

In the below pic, 3 different types of mushroom dye water is at work on the samples:

Hapalopilus rutilans (or nidulans)

And my personal favourite, the PINK dyewater from Hapalopilus rutilans:

In fact, the little piece of H.rutilans that I found was so unsignificant looking that I was struggling for a long time to identify it.
After passing a piece of it  (then called UNID4 in my strain collection) on growth medium to keep the strain in the hope that one day I will learn what this mushroom is, I kept the other specimen that I found in the fridge for maybe 1-2 months, before deciding that the matter is hopeless. So just out of curiousity I created dyewater out of it, and to be honest I wasnt expecting anything else than some weak brownish liquid. I had to sit down when I was faced with this intense purple coming out of that one little piece of shroom. It is really a pity that it was exhausted before I could have used it for real yarns.
I looked it up in Miriam Rice's books, and I realized that it only can be Hapalopilus rutilans. I cross checked with pics in the internet and my shroom books, and it was correct! This is how dyeing helped me in mushroom identification :).
Below are the fotos. You must know that I have a small, cheap and old photo mashine, which has real troubles giving back the actual shades of colors. Totally not suited for photographing dyed yarns.
So imagine more pink on the pictures, please.
After dyeing I bathed the samples for 30 min or so in acid (pH 4.5 by white sushi vinegar) and in alkaline (pH 10 by ammonia) water to see what pH changes might result in.
In case of H. rutilans not much happened, but in other cases some differences are apparent.
I am now convinced that one will get different colors if the dyebath itself is lets say pH 10 or if the dyebath is pH5-6, and after dyeing you bath/simmer it in clear water with pH10. It is an interesting topic and I will certainly run some test on it in the future.
But here are the photos:
pH 10 on the left, pH 4.5 on the middle and unchanged pH 6.3 on the right
It was interesting to see that bathing in pH10 water makes most probably the dye more water soluble and deattaches it from the yarn as after the bath the samples were visibly paler than the other two pH ranges (pH 6.3 unbathed and pH 4.5 bathed in water)

Xerocomus communis

Some photos of still wet sample yarns. If I check back the dry samples, it must have been Xerocomus communis.The very brown ones are iron mordanted on each one, the middle is with alum and the ones on the right are unmordanted. Number 2 and 1 had an afterbath in pH 10 and 4.5 water, while nr 3 had unchanged 7.2 pH.

Paxillus involotus and Hypholoma fasciculare

Next photo is of the drying swatches of Paxillus involotus and Hypholoma fasciculare. I compared results of scoured and unscoured wool, again unmordanted, alum and iron mordanted and pH 4.5 and pH 10 afterbathes. In case of these mushrooms I didnt see a significant difference in the results about scouring and pH afterbathes. Mordanting, however, does have an effect.

The unchanged pH is above 7, as both liquids were affected by bacterial contamination (they were also smelly :( ).

I have Hypholoma fasciculare in my strain collection, I expect it should be possible to cultivate the same way as Pleurotus or Shiitake species. I will give it a go and then re-run the tests with not bacterial contaminated dye water :).

Paxillus I will give a go as well, as folks suspect it might not be always mycorrhizal but able to exist on wood debris as well. (see MushroomExpert)

/ to be continued /

Start-up with dyeing - Part I.

During the late autumn period, after I got hooked up on Miriam Rice's mushroom dyeing books I prepared dye bathes from all sort of mushrooms that crossed my path in the forest in a volume bigger than my fingertip. Unfortunately I havent found too much from each species (ca. a handful max in most cases), so as it later turned out, my dye bathes were not awfully packed with colors.

Nevertheless, as I was looking at my rather washed out colors after the first slight disappointment I was thinking what amazing colors will I be able to achieve once I will have sufficient amount of mushroom available. Most sources suggest at least 1:1 ratio of mushroom and wool in weight, in my case wool heavily overwhelmed mushrooms.

As I also learnt by experience, different mordants and different dyes (and their combination) respond differently to different treatments. For some mushrooms, cold soaking for a day or so gives a pretty good result, while for others one must simmer it for some time, otherwise dye molecules wont attach to the fibre no matter how long it has been soaked. For shorter simmering one certainly gets brighter colors, but I tyically had to simmer for 2-3 hrs as the amount of dye molecules were so low in the bath, and so I mostly have duller tones.

Fresh mushrooms were used each time for the dyebath, with 3 exceptions of dried samples. Liquids were then stored in the fridge at 10C for some months. No need to mention, that most of them got pretty smelly either from bacteria or from fungi and their pH were elevated from around 6.3 (pH measured for non smelling liquids) to 7-8. I expect that extended storage may also affect coloring capability. Next time I will rather freeze them and prepare dyebath fresh. According to some freezing anyway has a good effect on the amount of available dye.

I used a commercially spun merino wool yarn (Regina Classic from italian manufacturer Adriafil, base color 002, lot 004 each) in white color for all dyeings, as I havent manage to locate hand spun untreated yarn in the vicinity and for a reasonable price. Apparently it is the official wool used on the "Speed Knitting" competitions, so should do as a start :).

The following species were tested, and they were willing to dye the fibers to a bigger or smaller extent. (Unfortunately some of them were fully exhausted already by the test swatches, and could not be used later on the half a skeins that I dyed.):

Piptoporus betulinus --> lignivorous polypore (dried) --> test swatch + half skein
Hapalopilus rutilans --> lignivorous polypore (fresh) --> test swatch
Hypholoma fasciculare --> lignivorous (fresh) --> test swatch + half skein
Calvatia excipuliformis --> saprobic (fresh, ripe) --> test swatch
Panaeolus (antillarium ?) --> saprobic (fresh) --> test swatch
Xerocomus sp. --> mycorrhizal (fresh) --> test swatch + half skein
Boletus edulis -->  mycorrhizal (dried from supermarket) --> half skein
Scleroderma citrinum --> mycorrhizal (fresh) --> test swatch + half skein
Paxillus involotus --> mycorrhizal (fresh) --> test swatch + half skein
Craterellus cornucopioides --> mycorrhizal (fresh from the market) --> test swatch

The next ones proved to be completely or mostly dye duds, not resulting in any significant dye with the so far tried methods:
Clitocybe odorata
Coprinus comatus
Coltricia perennis
Mycena pura
Pleurotus djamor
Trametes versicolor brown variety

/ see Start-up with Dyeing - Part II. for more /

Home made mushroom cardboard, tray and decors

There are quite a few good guides on the web (eg the one at Green Living Forum) explainig steps for preparing paper from mushrooms. It mostly includes chopping up the mushroom in a blender, while adding water, pouring it into a big bowl of water and with a specific technique collecting the material with a deckle, then pressing water out of it and leave to dry.

Coincidentally, similarly to Fergus on Green Living Forum I also started in mid January with Piptoporus betulinus, a wood eating polypore, found on birch. The 2 specimens I saved from the summer were already dry, but if you submerge to warm water for a short while it regains its structure and easy to chop up. One I simmered for an hour to test one of my first wool test swatches for its dye, the other was soaked only for the mentioned short while, but I did not find any difference in the end paper results.

I only had a pretty cheap onion blender, which was not willing to belnd the mushrooms in very small pieces, so I could not make a real thin sheet. I also did not manage to master the general method of collecting the puree from the water with a deckle, so at the end I pressed by finger the pieces on the deckle. My deckle is a simple tool used to prevent oil spillage over frying pans, but I hope to get once a real paper making deckle and see the difference. The best method seems to me for the newbies the one described at Green Living Forum, where you make a very finely chopped "cream" and spread it over evenly on the deckle.

Here is how mine interpretation of the process looked:

After that I used old but clean kitchen towels to soak up the excess moisture, and put them to dry above the radiator for the night. Miriam Rice's book advised to hang them up to dry among sheets of towels, but I didnt see how that would keep the sheets straight. I should have seen it better...

I also lined a plastic tray used as packaging for groceries with the mushroom and set it aside in the tray. As a third step filled up some silicone cooking forms with the remaining material.

To my greatest horror by morning the sheets where completely curled up on top of the radiator. I had to rewater them and place back to dry. I even ironed them, which was actually not that bad idea as it sounds, so far you iron them through a table cloth or so, as it compacted them and gave them a smoother surface. After that I placed them among sheets of papers into some big and extremely boring text book, and finally they dried straight.

THE PAPER

The end result were 4 small sheets of slightly flexible, ca.1mm wide, cardboard like sheets, which were pretty easy to cut to form with scissors.

As you may see it, the look resembles to corkboard and I think it could be indeed used for similar purposes. For use as cardboard alone might not be a good idea, as the sheets are rather brittle and would break easily. However, mixed with other, more flexible shrooms like Trametes versicolor would certainly give better results. Or one need to make a thicker sheet for a cardboard with increased strength.

THE PACKAGING

The tray was also okay, ca. 1.5-2 mm thick, though later deformed slightly in the middle. I think it could be used as packaging of certain specialty products where minimal liquid contact is expected (eg. whole pieces of organic apple, tray for beans, etc). Mass production is out of the picture, as fibrious mushrooms are typically polypores and they tend to develop rather slowly, but as a hobby activity may have a ground :).

The other point, why it would not be a viable setup, as there is already a better and quicker method to make mushroom packaging. Ecocradle, the trademark of the already mentioned Ecocradle company cuts out some innecesary corners from the process: they do not start to nurture fruiting bodies from the mycelial mass, but they rather use directly the substrate overgrown by mycelium as packaging material and cardboard. This means they can use more material and spare weeks if not months from start till end product. Some of their interesting concepts can be found here.

THE DECORATIVE ITEMS
As the last element of this small "project", the little cooking forms also dried, and the results were some pretty bears and hearts. I plan to make some jewelery items like small balls for necklaces and similar next time and will play a little with the particle size to see the changes in the surface.

The first post

I am quite a rooky (yet :)) in the vast kingdom of fungi (as well as blogging), but in the longer run I hope to be able to share bits and pieces of interesting information - and the results of the home made experiments. I aimed to organize information into Pages, but apparently that is not possible, so I will have to figure out something else to keep the blog tidy.

I am currently experimenting with cultivation of exotic mushrooms (mainly lignivore), which i ordered on Ebay or took samples from mushroom bodies bought in markets and shops, like velvet foot mushrooms, pioppinos, oysters, beech and straw mushrooms and co. I use spent coffe ground among others and the whole things is yet on an extremely small scale (like in 250ml jars...), and produce some very-very shy, long necked crop, due to lack of equipment and room. By may I will be moving to my own house with garden and a BIG cellar, who begs to be my lab + growth area, so I hope circumstances (and along with it results) will improve significantly.

During the summer/autumn period I have been collecting various wild mushrooms (i.e ALL that was not fast enough to run away ;) and which I could more or less identify), taken samples from them and built up a small strain collection. One of my personal favourites is the Agaricus augustus from the Anwerp zoo with its mesmerizing smell. (btw I hope it is still revivable - I havent got to pass it onto new medium since months...). If all goes well I will test-cultivate these folks (the non-mycorrhizal ones). Not the mention the glowing ones like Panellus or Omphalotus :D.

I noted also the strong smell of Cantharellus cibarius mycelium in the jars. Although this species produces mushroom bodies only when connected to its tree partner, and so commercial cultivation is not very viable yet, it really tickles me to see if the mycelium itself could be used as a spice or a main course, similarly to tempeh. Tempeh is fermented soybeans "conquered" by a type of mold (and it is delicious i can tell you). Of course production method would be different, but I imagine the end product's look similar to tempeh. I will come back to this point with more details later on.

My lure to mushroom cultivation is not only for food. Miriam Rice's books on mushroom dyeing took irrepairable damage on my course of life last Autumn in the good sense. Though myself was not yet able to produce convincingly strong colours on wool, it is mainly due to my inexperience and the minute amount of mushroom dye that I managed to save in the pre-winter period - and that I waited like 4 months before I used them up from the fridge. But I can hardly wait to lay hands on more mushrooms and as I have some dye goodies in my strain collection, like Hypholoma fasciculare or Hapalopilus rutilans, I have solid plans to produce my own mushrooms for dyeing. (Wish me good luck with it ;)).

Another aspect is the fibers in the fungi. It is possible to produce some very fine papers out of mushrooms, which I already ran some tests withm plan to upload the photos later on. There are also the ingenious Ecovative guys at http://www.mushroompackaging.com/ with their amazing products. They take a carefully selected agricultural waste material, get it run through by mycelium in a mould, disactivate growth of mycelium when run is complete - and there you have your packaging material. It is such a simple and beautiful idea! I am totally in love with it. They even use mushroom as insulation in buildings! Needless to say, that I can hardly wait for the day when these harmless, biodegradable, closed loop products overtake completely the place of styrofoam, etc.

One step further down the road would be to look for molecular building blocks, monomers, that could replace the current fossil dependant, non degrading polymers used in every aspect in our lives - at least in areas where use of these materials are not justified at all (eg. why use one way plastic cup that stays with us for hundreds of years as a minimum, to drink your coffe in 1-5 minutes...). I am also pretty convinced that it would be a better alternative compared to the use of agricultural crops, as would not endanger food resources. One possible candidate is chitosan, a chitin derivative, but I suspect there are more into fungi. I will keep you posted :).

- O, yes, one important point about mushrooms, that is known to wider audiences and was not mentioned so far: their medicial properties. I admit this is an interesting field, but a field laying quite far from my general interest. Nevertheless, I will do my absolute best to keep this subject also on the radar.
And as an absolute fun - and in case of proper amount of free time, I plan to create all sorts of imaginable and unimaginable decorational objects and jewelleries out of Fungi. I already saw some pretty inspirational blogs on this subject, and I hope to be able to unleash my creativity and come up with some pretty (and perhaps even useful and durable...) stuff.

Happy mushrooming for all!