Madder’s Family

Madder has several relatives that are also rich in useful reds. These plants are native here in Denmark, and have been used as red dyes a very long time back.

Believe it or not, the year is drawing to a close. So, I want to try to summarize all the many dyeing experiments I did over the year.

This summer, I searched for madder’s relatives, to find as many as possible. Madder, Rubia tinctoria, belongs to the madder family (Rubiaceae) in which you also find the bedstraws (the genus Galium).

Galium species do not contain as large amounts of red dye as cultivated madder does, but several of the species grow wild here in Denmark, and their historical use is well known.

Madder plant growing in my dye garden.

The first Galium species to present itself was cleavers (Galium aparine). It’s everywhere! Anybody who has ever walked outside surely know this plant. Or at least its seeds. They are extremely good at clinging to clothing and dog fur. The whole plant is covered with clingy hooks – the very same that cultivated madder has.

My attempt to dig up cleaver roots quickly came to an end. The roots have the thickness of sewing thread, so a lot of digging is required. But the roots are said to contain dye, so I’m keeping them on my list of maybes.

Cleavers up close. You can see the characteristic clingy hooks on the seeds. The very same that madder is covered with.

Lady’s bedstraw (Galium verum) is the plant mentioned by most natural dyeing books. I tried growing it in the garden this year, seeding it outside in the spring, but nothing grew.

Whenever you’re looking for a specific plant or mushroom, but haven’t found it yet, it’s simply invisible. But, once you find it, you start seeing it everywhere. The relationship between Lady’s bedstraw and myself developed exactly like that over the summer. Once I found it, it was everywhere! For example this coastal grassland:

Coastal grasslands with very sandy and infertile soil, perfect for Lady’s bedstraw. I took this picture in a region of Denmark called Mols.

 

Lady’s bedstraw truly thrives in the nutrient-poor, sandy soil, along with yarrow and St. John’s wort.

Lady’s bedstraw growing in a big cluster.

Unfortunately, several walks with a shovel only yielded a very small handful of Lady’s bedstraw roots – so little that my scale didn’t register. Like with cleavers, the roots are extremely fine, and they tangle up with roots of grass etc. In combination with stony, sandy soil, the digging job gets hard. To get your hands on a larger pile of these roots, I suspect you have to grow them in a well-prepared sandy soil without obstacles. Anyway, I tried dyeing with my small handful of roots, but it gave almost no color.

But then, on a forest walk, this plant turned up – hedge bedstraw (Galium mollugo):

Flowering hedge bedstraw photographed in July.

Hedge bedstraw is also mentioned by different books as a dye plant, so I brought out the shovel once more. Again, it was difficult. The forest soil is obviously full of tree roots that make digging quite impossible. But I managed to get a couple of handfuls of roots, mainly because hedge bedstraw roots are not that thin. I dug up the roots on July 9th. The next day, after cleaning, the slightly dried roots weighed 30 g.

My pile of hedge bedstraw roots, with reds clearly showing under the out bark.

I soaked the roots in cold water overnight, then dyed my usual alum mordanted 12-gram skeins of Fernris to test the dye. I removed the overnight water because Jenny Dean does, but I should have concluded from my madder experiments that it is not necessary to do so. The water used to soak the roots overnight simply contains a small amount of dye, with the same properties as the dye you extract when you heat the roots in water (the small 6-gram skein laying across the others in the picture below was dyed with the discarded water).

Then, I dyed alum mordanted 12-gram skeins in a 1st and 2nd dyebath, in exactly the same way as if it had been madder: heating up to 60 degrees C, then leaving the yarn in the dyebath until the next day. The first bath gave a convincing red-orange, which would not have been a surprise had it been madder I was dyeing with. The dye is less abundant in hedge bedstraw than in madder, but the difference is actually smaller than anticipated. Here, I used 30 g of roots on 12 g of yarn, with madder, you would get this shade with less than 100% weight of fiber.

After the second bath, which also worked well, I was evidently feeling on top of things, and threw in a 50 g skein. There was not much dye left, but to extract everything that was there, I left the bath with yarn in a jar outside. That was in mid-July.

A couple of times, I heated the entire jar over a water bath to give the process a helping hand, but the rest of the time, it was just standing there. I turned over the yarn to get an even dye, and for a while, it also fermented. Both time and fermentation should help release the dye. Also, I imagine that a skein of yarn in the bath will soak up the dye as it is released, permitting more to come out (alizarin has a rather low solubility in water). In any case, my large skein stayed in the jar for 6 weeks, and turned out a pleasing coral color. And, the dye bath ran clear, so there was probably nothing left in the roots.

Dyeing with roots of hedge bedstraw (Galium mollugo). 1st bath (left), 2nd bath (middle), and the large skein on the right is fermentation of the 3rd bath. The small skein across was dyed in the water used to soak the roots overnight.

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Spring Cleaning

In the summer, when all the plants stand tall, I usually collect good bundles of tansy, yarrow, and other wild dye plants. And they have to go before the next harvest.

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My dyestuff stores from last year contained big bundles of mugwort and tansy, a smaller amount of yarrow, a box full of dry velvet pax, and dry pomegranate shells (among other things).

Spring has shown itself from its worst side this year, but I’ve managed to get outside with my little stove on an extension cord, working to bring down the amount of stored dyestuffs.

First, velvet pax. I found quite a nice harvest of this mushroom last year, more than half of what i found was from driving through a small forest, spotting the mushrooms, and hitting the brake!

I had 190 g of dried mushrooms. On 100 g of wool, that gave a good green (middle skein in photo below) and the afterbath a green-beige (right). I could not capture the color in the photo, but I was pleasantly surprised how well the dried mushrooms retain the color potential, including the green tones. In conclusion, velvet pax is a very good dye mushroom, fresh or dry.

There’s a beige skein on the left in the photo below. That’s 100 g of yarn, dyed with enough dried mugwort to fill a large dye pot completely. I even gave it an iron afterbath. Thinking back, this is actually the second time i get dull beige from dry mugwort, and the conclusion is that it does not dry well. The fresh plant, on the other hand, gives a nice yellow-green.

From left: dried mugwort and iron, dried velvet pax, 1. and 2. bath.

Next up, pomegranate shells. I had saved a very modest amount of shells, from just two fruits, weighing 85 g dry. I followed Jenny Dean’s “Wild Colour” and put the shells in a plastic bag and pounded them with a hammer. To test the new (to me) dyestuff, I wound two 12-gram skeins of Fenris (100% wool) and a small 5-gram skein of Bestla (silk-merino).

The pomegranate shells gave nice yellows on wool and silk. I modified one of the wool skeins with iron, and that gave a darker, greener tone, that actually looks a lot like the color from velvet pax.

Next time people eat pomegranates around here, the shells will be saved. They give a nice color, and they are available during winter, where little else is there in terms of fresh colors.

Pomegranate shells on silk-merino (back) and wool (middle), and modified with iron (front).

Several large bundles of yarrow, tansy, and mugwort turned into the yellow-beige first dye for a new round of matrix dyed yarn for Baby Vindauga kits. The second yellow os weld, and the skeins are overdyed with indigo as usual to produce the 9 different blues and greens.

Matrix dyed wool in blue and green.

And once I got started, a matrix in purple and blue, using cochineal and indigo, also appeared.

Matrix dyed wool in purple and blue.

The matrix skeins turned into contrast colors for new Baby Vindauga Kits, you can see them at my Etsy shop:

Purple-blue Baby Vindauga Kit.
Green-blue Baby Vindauga Kit.

Lichen Windfall

Lichen windfall is perfect for natural dyeing, since it does no harm to pick up the fallen ones, they will no longer grow. One of the most common and easy-to-recognize lichens in windfall is Ramalina fastigiata.

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When walking outside on rainy, windy days, I very often find lots of lichens scattered on the ground under trees. Lichens that the wind has torn down from branches. Sometimes, on the day after a big storm, I’ve come home from walks with all my pockets plus random trash bags filled with windfall. Wonderful windfall with that amazing scent that only lichens have.

Collecting windfall does no harm, since these lichens are not able to continue growing anyway. It’s the best (some would say only) way to obtain lichens for dyeing. When I come home with such a treasure, I usually spread it out on a plastic tray to dry (to prevent mold).

Lichen windfall drying at home. It looks like a big piece of Evernia pruniastri on the left, Ramalina fastigiata on the right, and probably a Parmelia species on the bottom.

But before dyeing with lichen windfall, it’s necessary to sort the lichens and determine the species, since you will need to use the boiling water method (BWM) with some species, and the ammonia method with others:

Boiling water method – it is what it sounds like. Simmer the lichen in water and cool off. Add the yarn to the dye bath and heat it for an hour without boiling.

Ammonia method – the difficult one. Steep the lichen in 1% ammonia (originally, stale urine was used) for several weeks or months, opening and shaking the jar daily to aerate. The red liquid in the jar is the dye bath.

In both methods, no mordant is required, since lichen dyes are substantive (they bind directly to wool without the help of a mordant).

Lichens steeping in 1% ammonia.

In order to type lichens, I recently bought myself a copy of “Lichens, An Illustrated Guide to the British and Irish Species” by Frank S. Dobson. It contains a detailed introduction to lichens, and a detailed key with photos and descriptions.

With my copy of Dobson, I’m planning to take a closer look at the types of lichens that are commonly found in the windfall here in my corner of Denmark. That is, how to recognize them, how to dye with them, and which colors to expect.

I’m beginning with a very common type of lichen, which may very well be the easiest one to recognize: Ramalina fastigiata. Often, large tufts of this will fall, and they are completely covered in small outgrowths that look like tiny suction cups. The outgrowths are apothecia, the fruiting bodies of the lichen. They make spores for sexual reproduction. When the spores germinate in a new location, they meet with a new alga to become a new individual lichen. But the dyer doesn’t have to worry about all that, being able to recognize apothecia is the important part.

A piece of Ramalina fastigiata, completely covered in apothecia. Tufts like this can measure up to about 5 cm (2 inches).

Karen D. Casselman mentions the Ramalina species on the list of ammonia methods lichens in her book, “Lichen Dyes, The New Source Book”.

I’ve previously tested the ammonia method on Ramalina fastigiata and achieved a light rose color (pictures here).

But Casselman also mentions the and Ramalina species in her list of boiling water method lichens, so I decided to test that method on Ramalina fastigiata. I used equal amounts of wool yarn and lichen, and achieved no color at all (no pictures!). The conclusion: Ramalina fastigiata is strictly an ammonia method lichen.

An Earthball Study

Earthballs contain a yellow-brown dye, but also a large and annoying amount of tiny, black spores. So I set out to find out if the spores contain any dye or if they could just be discarded.

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Common earthball, Scleroderma citrinum.

A couple of years ago, I dyed a lot of yarn with earthballs. The color turned out a nice yellowish brown, but the yarn was simply full of spores that continued to drizzle out, both when winding the yarn into skeins and when knitting with it.

The drizzling pores were obviously annoying, but I also started wondering if the spores are even safe to breathe? It’s usually said that earthballs are “moderately toxic”.

In their book “Färgsvampar & svampfärgning” (Dye mushrooms and dyeing),  Lundmark & Marklund label earthballs “good” dye mushrooms, so it would be a pity to give up on earthballs just because of the spore problem. Lundmark & Marklund mention that earthballs contain the dyes badion A, norbadion A, and sclerocitrin.

Sclerocitrin is also described in the research paper “Unusual Pulvinic Acid Dimers from the Common Fungi Scleroderma citrinum (Common Earthball) and Chalciporous piperatus (Peppery Bolete), Angewandte Chemie International Edition, 2004, 43, 1883-1886 by Winner et al. They show that the “brilliant yellow” dye sclerocitrin is found in “remarkable amounts” in earthballs. As the title says, sclerocitrin is also found in peppery boletes. I haven’t looked for it, but a mental note has been made.

Earthballs have a dark or black spore mass inside, surrounded by a relatively thin outer wall. I decided on a small experiment in order to see if the spores contain any dye. If not, it would make sense to just leave them in the forest.

Halved earthballs with grey and black spores inside.

I used as small amount of earthballs for my experiment, gathered during the fall of 2016 and dried until use (2016 was not a good mushroom year, so not many earthballs were to be found).

Separating the spore mass from the mushroom’s outer wall was incredibly difficult. The parts were completely stuck together in the dry mushrooms, but in the end, I had 23 g of out walls and 10-11 g of spores. I soaked both overnight, the outer walls simply by adding water. The spores were stuck together in stone hard lumps that I separated by grinding them in my mortar. The spores repel water, I solved that by wetting them in denatured alcohol, then adding water.

The next day, I boiled the two dye baths and filtered the spore bath through a coffee filter. It took very long for the liquid to run through, that’s always the case when filtering a solution with many tiny particles. I then dyed a 10-gram alum mordanted test skein (Fenris 100% wool) in each bath, and got the result below – almost the same color from the two.

The top skein of yarn in the picture is dyed with the outer walls, the bottom one with the pores. I had hoped to find that the pores didn’t dye, but clearly that’s not the case. In principle, it’s not surprising, though, to find that sclerocitrin and the other pigments are distributed throughout the mushroom. The dark color of the spores is not caused by a pigment that acts as a dye.

In conclusion, all parts of the earthball contains dye, and discarding the pores would mean discarding a lot of good dye. So the best method for earthball dyeing would be using the entire mushroom, wetting the spores with alcohol, and then investing the time required to filter the entire dye bath before any wool is added.

Yarn dyed with different parts of earthballs. The top skein is dyed with the outer walls only, the bottom skein with pores only.

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Amazing Dyeing Failures 1

Failure in natural dyeing is commonly defined as not getting the result you expected. Beige, off white, baby yellow and other tones of grime are all examples of colors I have made no attempt to acheive, and yet, I have a big pile of skeins just like that. But there’s actually a lot to be learned from failures. Some give new ideas of what to try next. Others just tell you what not to do. Below, I’ll describe some of my failures – actually, I’ve failed so many times that this will only be the first installment, more to follow.

Alle de mislykkede og uønskede farver. Efter billedet blev taget overfarvede jeg med indigo.
Skeins of failure. They were all overdyed with indigo after taking the photo.

1: Bark Failure

Several books on dyeing will tell you that different types of barks are good dyestuffs. For example, Jenny Dean’s “Wild Color” mentions these barks and the color they should produce on alum mordanted wool: alder (brown-green), barberry (yellow), ash (bright yellow-green), apple (warm yellow), oak and willow (beige), and finally elm, birch, cherry, pear, and plum (pink).

For a while, the theme of my walks was bark; in the end, I found enough of these three to try them as dyestuffs:

  1. Birch (Betula) – I’ve used birch leaves several times for a sunny yellow, but not the bark. Some trees were cut down near our house, and I jumped at the chance. The trees had been left in a big pile, which I obviously had to climb to get to the good parts, and since I was of course wearing clogs, I fell down from that big pile in the end. With 60 g of birch bark in my pockets.
  2. Another day I hear some men working outside, shredding logs. On their day off, I casually walked by and managed to peel a good amount of bark off. The logs turned out to be alder (Alnus), the kind with the tiny cones. 70 g of bark.
  3. Last one is some bark from a forest walk. I jumped over a big, big ditch to get this. I’m pretty sure it’s beech (Fagus). My daughter jumped it too, so I had to save her afterwards. 94 g of bark.
Dagmar tæt på at falde i grøften
Dagmar, seen moping, came close to falling into a large ditch.

I used Jenny Dean’s general dyeing method for bark. She says that “barks are best soaked for several days or even weeks in cold water before processing. Then simmer them for one hour. Never boil bark, as this will release too much tannin”. So that’s what I did – left the three types of bark to soak for a couple of weeks. That was long enough that they started fermenting, and I can tell you that it didn’t smell that good.

But when I simmered 10 g test skeins of alum mordanted wools in the three bark dye baths, the color in the end was pale beige. I didn’t even bother taking pictures (because when you’ve seen one skein of pale beige wool, you really have seen them all), but you can see one sticking out between the pale pink skeins in the left side of the first picture above.

I have seen other dyers experiment with bark (for example, at my wool group’s dyeing day) and also get pale beige or off white. So right now, I’m not even convinced that it would ever work, and I probably won’t try it again unless someone can tell me what went wrong (please comment below if you know or if you’ve had good results dyeing with bark).

2: Slimy/Moldy Avocado Failures

There are established procedures for dyeing with avocados, but I’ve been experimenting with slightly different ways of doing it. I suppose to make the procedure easier and better, but of course ending up making it messy and complicated.

According to Carol Lee, avocado pits should not be allowed to dry before use because they will become so hard that they are impossible to chop. Instead, they should be frozen until use. I wanted to find a way to dry them anyway because my freezer is small.

So I chopped the pits and skins and then left them to dry. This worked well on a couple of occasions, but most times it did not because they became completely overgrown with mold before they had time to dry. Moldy materials may still work as dyes, but I think it is generally unwise to handle them repeatedly around the house, since many molds produce toxins that may be inhaled. So I went back to freezing the skins and pits.

dryavocado
Avocado pits and skin turn red as they dry, so it’s not that surprising that the dye bath they produce is also red.

Another experiment was to ferment the pits and shells for a looong time to see if they yielded more color that way. I used my dry material, soaked overnight, but I suspect the results would have been the same had I used frozen dyestuff.

I usually ferment avocado pits and skins by heating them up once in brine, then just leaving them. Normally for a few weeks or a month, this time for six months. And the dye bath did develop a deep red, but it also became extremely slimy.

Despite the sliminess, I tried dyeing a small test skein in this dye bath, but it didn’t yield good color. My guess is that the slime prevented good contact between yarn and dye. But I’m not convinced that a long fermentation couldn’t yield good color. I’ve been adviced to put avocado pits and skins in jars, close the jars, heat them up, and then ferment. Such jars should not go slimy. I’ll try that next time.

Beige med lidt rødlige striber
Beige with a red streak, that’s the look of yarn dyed with avocado slime.

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Late Summer Greens

This summer, I’ve dyed a nice pile of green wool using reed flowers and velvet pax – two dyestuffs that are a highlight of the dyer’s year. Reed flowers because they give such an electric green. You have to admit it’s a bit strange that these red flowers dye wool a wild green, but only if you get them into the dye pot absolutely fresh. If the flowers have opened or are not freshly picked, they will only give yellow. Velvet pax because its dusty greens are so lightfast. The two skeins in the back are dyed with velvet pax, the three in the front with reed flowers.

grøn green
Greens from reed flowers and velvet pax, the essence of late summer dyeing.

I’m becoming better at finding velvet pax. The first couple of years, I looked for it too late in the season. This year, I’ve found it growing several places, for example this archetypical plantation, where Dagmar is picking a big one. Just the kind of place that velvet pax likes to grow.

Dagmarplukker
Dagmar picking velvet pax (with the arm that’s not broken).

Velvet pax can be found in August, and this year, everything was early, so it was there at the beginning of August. And the mushrooms were huge – I found some that were 25 cm across.

sortfiltet
Characteristic brown tops of velvet pax, captured in a typical habitat.

Big, fat spiders are another joy of late summer. This one, which is possibly the fattest spider I’ve ever seen, lives outside our house. When I was sticking my camera right in its face, the neighbor’s big dogs started barking. Immediately, the spider lifted its front legs as if to attack. I chose to run away, so I only got a good shot from underneath the spider, where its pattern looks a bit like eyes. I think it’s a very light colored cross spider, since its body is pointy at the back. After reading that they can bite if provoked, I think my decision to flee was not a bad one.

edderkop
My pet spider.

Summer is also the time of year to test light-fastness. I tested a handful of colors on the windowsill from early July to mid August, and their light-fastness was quite different.

  1. Old polypores, the two top ones warm baths and the lower one a cold bath that brewed outside for some weeks. None of these yellow browns are very light-fast.
  2. Velvet pax, the color didn’t change. I’ve seen this light-fastness in previous test, so it really is that good!
  3. Orange Cortinarius mushrooms, I don’t know which species. Not that light-fast
  4. A matrix of madder and indigo, showing that saturated colors are much more light-fast than pastels
  5. Sorrel root, not very light-fast
  6. Birch leaves. Surprisingly light-fast
  7. Weld. Surprised by the fact that it’s less light-fast than number 6…
  8. Henna on alpaca. I’d say this is a medium light-fastness
  9. Calendula flowers. Surprisingly light-fast
Light testing summer 2016.

I’ve also dyed with tansy, which doesn’t give green, but “just” yellow on alum mordanted wool (no pictures of that). But when I admired the flowers, I suddenly wanted to check if they really do stick to Fibonacci numbers.

The Fibonacci series begins with two ones, and then the next numbers are found by adding the two previous ones:

1, 1, 2, 3, 5, 8, 13, 21, 34, 55, etc.

The last time I thought about Fibonacci numbers were for calculating the numbers of my Vindauga blanket where rectangles obey the golden ratio, approximated by the ratio between neighboring numbers in the Fibonacci series, eg. 55/34 = 1.61.

Below is a close-up of a tansy flower. And as promised, the numbers of rows of tiny buds are Fibonacci numbers – 13 clockwise rows and 21 counter-clockwise.

DSC_2985
Tansy flower obeying Fibonacci’s sequence.

Summer Rain

This summer passed in a big cloud of rain, which has been lovely for plants and mushrooms that came out early and in huge numbers. We went on lots of day trips, for example Skovsnogen Artspace:

skovsnogen
Skovsnogen artspace, a forest full of sculptures.

My mom has managed to finish a couple of knitting projects with yarn that I’ve dyed. An Elizabeth shawl designed by Dee O’Keefe in Einband that I’ve dyed with madder. This Icelandic wool is wonderful to knit with and to wear, but it also takes color beautifully. She also knit a pair of socks, the pattern is Laurel by Wendy D. Johnson, the yarn a sock yarn I’ve dyed purplish blue with indigo and a twist of cochineal.

wendyknitting
My Mom’s knitting successes, using yarn that I dyed with madder and indigo.

We went on a day trip to the hilly landscape at Rebild. The sheep are a perfect match for this landscape, and in the end, it is their grazing that maintains the heath (blueberries though, they don’t touch). I don’t remember ever seeing such steep hills anywhere else in Denmark – it tells you about the power of the melting waters from the end of the last ice age.

rebild_bakker
The hills of Rebild.

Rold forest is close by. There, we saw the unusual old beech trees, called “purker” in Danish. They have multiple contorted growths because they were cut down repeatedly for firewood. Fallen logs are left to rot, giving mushrooms and insects a much needed habitat.

roldskov
The ancient forest of Rold.

We also encountered biodiversity on the island of Livø. We went on a guided tour of the organic test farm, where experiments are made with growth practices for organic farming, as well as testing new crops such as quinoa and buckwheat.

It’s always a good thing to see a field of crops with lots of other plants in it, such as clover and cornflower. I’ve always loved cornflowers, but I do see them in a new light after reading about their color in “Handbook of Natural Colorants” by Berchtold & Mussak. The color comes from a supramolecular, self-assembled, complex of cyanidins, flavones, and metal ions (Mg2+ and Fe3+), and that’s why it cannot be extracted for dyeing. The complex comes apart, and the individual parts are not blue. This could be the case with other pretty colors that are impossible to extract? The amethyst deceiver failure comes to mind.

livø
On the island of Livø, off the coast of mainland Denmark.

I obviously couldn’t walk outside an entire summer without looking for lichens. I’ve added two books to my lichen library, one is a small and useful Danish pamphlet, “Laver i Tisvilde Hegn” by Hørnell, Jeppesen & Søchting. The other is the elaborate, somewhat academic “Lichens, An Illustrated Guide to the British and Irish Species” by Dobson.

I always find the most common lichens: Evernia prunastri, Ramalina fastigiataXanthoria parietina, and Hypogymnia physodes which I’ve already experimented with for for dyeing. So this summer, I’ve looked for Cladonia species.

I’ve often seen the funnel shaped lichen (top left in the image below) on the ground and on dead trees, and I believe it’s Cladonia fimbriata. I haven’t collected this lichen, since I’m not sure how to. One funnel at a time? Also, Casselman’s “Lichen Dyes, The New Source Book” does not mention this species.

Then there’s the reindeer lichens. Until recently, I thought they were mosses, but it’s never too late to learn something new. I found Cladonia portentosa (top right) in several places this summer, and my books do say that it is common, so I’ve collected a bit for dyeing.

I’ve only seen the bottom row lichens once each this summer, so I only took photos. Never pick a lichen if you don’t know if it’s rare. On the left, I believe, Cladonia rangiferina, and on the right, Cladonia coniocraea. Casselman does mention Cladonia rangiferina as a bwm (boiling water method) lichen that dyes shades of red to brown. Maybe it’s more common in other parts of the world.

cladonia
Different Cladonia lichens.

Home again, I’m beginning to prepare for the workshop on natural dyeing that I will teach the first weekend of October.

Walks in March

The weather here in Denmark has been all that bad in March. I suppose you’re supposed to make some comment along the lines of “in like a lion, out like a lamb”, but really, good and bad weather just depends on expectations. I’ve been sitting outside in the sun a couple of times already, knitting. And we’ve been on several good walks. There isn’t much new growing yet, so the most interesting living things right now are mushrooms and lichens growing on trees.

This is Evernia prunastri spreading over a tree trunk. I didn’t gather any of it this time, but I have dyed with it before. You have to use the ammonia method on this lichen, in which case it yields nice tones of pink.

everniaprunastri

We also walked to the edge of Gudenaa, the largest stream in Denmark (I wonder if you could even call it a small river?). Early in March, it had flooded quite a large area, but that is already coming down now.

gudenaa

We have borrowed a piece of garden not far from there, and I managed to plant the first seeds on February 28th, dyer’s greenweed. Before that, I kept the seeds in the freezer for two weeks because they need cold stratification to break dormancy. I didn’t try the freezer last year, and none of them sprouted last year. So I hope it works this time.

We also found ourselves walking in the forest, which was full of interesting things despite the time of year.

Here, a lot of cones that have been picked apart. We found them in a big pile under a tree. It’s the work of a squirrel, its signature being that there are still some bits of material sticking out from the cone’s stem. If a mouse had eaten these cones, it would have cleaned everything off.

egernkogler

We also found jelly ear / wood ear (Auricularia auricula-judae). It is edible, but they were a bit slimy, so we just left them.

judasøre

A perfect leaf skeleton (found by my daughter)

blad

And this log, patterned by the paths eaten by worms when it had bark. I wish I could knit this pattern…

træstamme

Finally, I found a lot of rusty gilled polypore, Gloeophyllum sepiarium, which grows on dead coniferous tree, it even grows well on treated logs, like these ones:

fyrrekorkhat1

These polypores are old, there’s actually lichens growing on them

fyrrekorkhat3

And here it’s growing in neat lines, guided by the cracks in the tree

fyrrekorkhat4

This is what the mushroom looks like seen from the bottom. It looks like gills, but this mushroom is a polypore with quite oblong pores.

fyrrekorkhat2

Rusty gilled polypore is supposed to contain a quite good brown dye, so I harvested a nice pile of them. More to come on that!

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Old Polypore

Dyer’s polypore is one of the very good dye mushrooms found here in Denmark (and many other places, including the rest of Europe and North America). It grows on dead wood, or parasitically on the roots of living trees. It grows in the same spot year after year, and grows a new fruiting body every year. That means you will often find dried-up mushrooms from the previous year close to the fresh growth of the year.

I’ve often found myself standing in a forest with a bunch of dry polypores from the year before, thinking that it was really too bad that they were wasted. So I decided to collect some, to test if they still contain any dye.

oldmushroom

I tried a single mushroom, weighing 24 g (it obviously would have weighed much more when it was fresh). I chopped it mushroom in small bits, and that partially powdered it.

I tried the dye bath on a 10 g test skein alum mordanted wool, and it turned a nice yellow-brown. So I used the bath a second time, and got a lighter shade. The old, dry mushroom clearly has a smaller dye potential than the fresh ones, but it does contain dye, so there’s no reason to leave it behind in the forest.

oldpolypore

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Mushroom Dyeing of 2015

allesvampefarver2

2015 is history, and it’s now 2016, but I think there’s just time to show you my mushroom dyeing of 2015, which brought a quite nice mushroom harvest.

Fall is my favorite time of year. Always has been. It’s the colors, the scents, and the long forest walks. We go to the same plantation in the northern part of Denmark every year, and this year was no exception. Part of the area has recently been turned into a test center for wind mills, but luckily, the windmills didn’t disturb the mushrooms! And they actually please the eye, the windmills, as they peek over the trees – especially when you consider their part in ensuring that Denmark will actually live up to its climate goal of 40% CO2 reduction in 2020.

windmills

My family already picked mushrooms before I was born, but always for eating, and always from a small, safe repertoire of about 5 species, with the main emphasis on the chanterelle, because it is very tasty and very easy to recognize.

We still hunt for edible mushrooms, and we are even training the next generation. See what an expert chanterelle hunter my 5-year old is:

dagmarkantareller

But these days I also hunt mushrooms for dyeing, and that makes it even more fun to walk in the forest – I always find something interesting! This is the yarn I’ve dyed with mushrooms this fall:

allesvampefarver

I’m really happy with this lot, and I’m thinking about a project where I could use all these colors together.

From right to left, they are dyed with common eartball (brown skeins, 900 g of mushrooms on 150 g of yarn), velvet pax (green-grey), Cortinarius semisanguineus (rose), some mixed Cortinarius ssp (tan).

I don’t know which mushroom the orange skein is dyed with. I didn’t take pictures of it, but I think it was a species of Cortinarius. Here’s the orange skein seen on a page of my big mushroom book with some species that it could possibly be, most of which are really poisonous. It’s hard to tell different types of Cortinarius apart, and some of them very poisonous, so always keep them apart from food mushrooms!

orangeslørhat

The light yellow skeins were dyed with common rustgill (Gymnopilus penetrans). It’s a very common mushroom, and after walking through an entire forest of them, I finally picked some. After trying it in the dyepot, I don’t think it’s a spectacular dye mushroom. There’s a number of ways to achieve this yellow color, and it’s not very abundant in this mushroom.

plettetflammehat

I also found a lot of sulphur tuft (Hypholoma fasciculare) which I find to be a mediocre dye mushroom, since it gives just another yellow, and not even a lot of it.

svovlhat

The last skein is best described as “off white”. I tried to dye it with amethyst deceiver although I sort of knew it wouldn’t work.

purpledeceiver

They look so pretty on the forest floor, but unfortunately, you’re best off just leaving them there. The purple color is indeed deceitful. It vanishes when you store the mushrooms for a couple of days, it even vanishes if it rains on them while they are still growing. This last fact tells you to give up right away. Predictably, even a large amount of mushrooms give no color on yarn, but I guess sometimes the true experimentalist has to verify the obvious.

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