Goldenrod Ball Galls

One of the things I love about dyeing with plants is that plants are amazing and awe-inspiring in so many other ways, too. First of all, they create their own food from energy from the sun, and provide all of us oxygen-breathers and plant-consumers with life and sustenance. For that alone I am so grateful. And that’s just the tip of the iceberg of amazing things about plants!

They are an integral part of complex inter-relationships that are not always obvious at my human eye level. I catch glimpses of some of these sometimes while I walk in the fields and woods, or when I garden. It makes me realize how much I don’t know about the intricate network of relationships between plants, animals, and microorganisms that are going on around me all the time.

Over the past several months I have noticed, and have had questions about, structures that I have found on fiber and dyeplants. I thought I’d share some of what I’ve been learning.

First, let’s consider goldenrod. The kind I have used for dyeing is Solidago canadensis, I believe. I didn’t end up running any dyebaths with goldenrod this summer. Nevertheless, all summer I kept a close eye on it while it budded, bloomed, and went to seed. In the late fall, the stalks had dried and become woody. The structure of the galls on the stems was clearly visible.

Here is a photo of galls at Wentworth Farm conservation area on November 24th. I was struck by how many of these galls there were in a relatively small space:

It makes me think of a futuristic city with high rise apartments accessible by flying rocket-cars:

Generally speaking, galls work like this: an insect lays an egg on a plant, and some kind of irritation or stimulation causes the plant cells to swell up around it. The swelling makes a cozy home for the baby bug while it grows and develops. Eventually, the adult bug emerges and continues its buggy lifecycle.

The galls I photographed are, I believe, goldenrod ball galls. I identified them using two excellent resources at my school‘s library, Naturally Curious by Mary Holland and Entomology by Ellen Doris. For on-line resources, the University of Wisconsin Extension Master Gardener page and this Nature North page were very helpful.

Here are some more details about goldenrod ball galls. Goldenrod ball galls are made by the goldenrod ball gall fly, Eurosta solidaginis. The adults are teeny little things, about 5 mm long, which makes me feel better about the fact that I’ve never noticed one.

In the late spring, the female fly lays eggs in the leaf bud at the tip of the stem. As soon as it hatches, the larvae drills into the bud and starts feeding. In response to the chewing, or perhaps the secretions of the larva, the goldenrod stem thickens. The larva eats the juicy and nutritious tissue inside the gall as it grows, and makes a little chamber for itself inside. It takes 3 or 4 weeks for the gall to fully form. The larvae molt a couple times through the summer and fall.

Each stage of larval development is called an “instar” which sounds sort of magical.

The third-stage larva is able to survive the winter by producing glycerol and sorbitol, which prevent its cells from damage by freezing. The third instar goes into a kind of hibernation called diapause all winter.

In the spring, the larva chews an escape tunnel through the fibers of the gall, stopping just before the outermost skin of the gall. Once it has fully metamorphosed, the adult fly doesn’t have any mouth parts. Insects plan ahead! The adult fly crawls out through the tunnel that it dug for itself earlier, then pops through the outer layer of the gall by inflating part of its head.

The adult flies only live for a couple weeks, without eating. During this time they mate and lay eggs. They do not travel far from where they are born, which is why there are often a large number of galls in a small area. The females use chemical sensors on their feet and antennae to make sure they are laying eggs on the right goldenrod species.

The exit hole is very small when the adult fly emerges. The ones that I found had holes of various sizes. Some of them had large holes that looked like they had been chipped away.

It turns out that downy woodpeckers and black-capped chickadees can detect the larvae inside the gall and dig them out to eat. The woodpeckers make a neat hole, but the chickadees have to chip away with their smaller beaks, so the holes they make are messier.

I’m guessing this one was made by a chickadee:

Inside-Outside Part Two

In this post I will describe more details about the dyebaths we made at the Inside-Outside Conference in Keene on October 21st. We ran four dyebaths with madder root, marigolds, weld, and orange cosmos.  As usual when I am running or leading an event, I didn’t get any photos. Hopefully the notes provided here will be useful even if they are lacking in visual information.

First of all, the fiber we were dyeing was woolen yarn. We dyed four skeins, each of which was 4 oz. I had pre-mordanted the skeins many weeks earlier with aluminum sulfate at a rate of 2 Tbsp. per 8 oz. (2 skeins could fit in a pot). The skeins had dried in the meanwhile, and had been soaked in water on the day of the workshop to “wet them out”, i.e. make sure they were thoroughly wet before dyeing.

Each dyebath heated for about 45 minutes to an hour. While they were heating, we were inside and they were outside, so we did not monitor the temperature. We strained the baths one at a time over a fifteen minute period of time. The skeins only sat in the dye baths briefly at first, long enough for participants to see some color emerging on the skeins. Then I had to pack everything up and bring it home. I transported the wet skeins in tubs and the dyebaths in empty gallon jugs. Once I got everything home, I put the skeins back into the pots with the dyebaths to soak. The next day, I heated the skeins in the dyebaths for a full hour and let them soak overnight again before pulling them out to dry and eventually wash.

Madder–The day before the conference I weighed out 4 oz. of madder roots, put them to soak in a pot of water, added about a teaspoon of calcium carbonate and a Tablespoon of soda ash (which made the bath about pH 8), heated the pot to about 160 degrees, maintained the heat for an hour, and then let the whole thing cool overnight. I’m not exactly sure where I picked up this term, but when I’m heating up plant material to make a bath I often refer to it as an “extraction”. So, this was the first extraction of the madder roots. I saved the strained liquid from that extraction, plus the original roots, and we made a second extraction from that same 4 oz. roots at the workshop. We added more water to the roots, plus calcium carbonate and soda ash to get a really high pH–it was like pH 10 or 11! I figured this wouldn’t hurt the roots. Plus we were pressed for time and getting more water to dilute the bath involved running up a flight of stairs and down a hallway to a bathroom with a shallow sink. So, pH 11 was good enough for the time being. We let that heat up on the low/simmer side of the hotplate for about 45 minutes.

When we came back outside to make the dyebaths, we strained out the roots and combined the high-pH extraction with my original extraction. The lower-pH bath was a much more orange color, whereas the high pH bath was much more of a cherry-red. We could see a difference in the color of the foam at the surface when we poured the two baths together. Together they made a sensible pH for wool. We lowered the first 4-oz. skein into the dyebath so that participants could see the color “strike”. That’s what it’s called when color first starts to attach to the fiber. It happens quickly with some dyes and more slowly with others.

The next day, I re-heated the original skein in the original bath for an hour at 140 degrees or so, and let it sit overnight. Then I exhausted that bath with a second 4 oz. skein. After that, I extracted the same roots two more times with calcium carbonate and soda ash to keep the pH around pH 8-9. I combined the two extractions to make a third dyebath, and dyed a third woolen skein that had been premordanted with alum. The combined exhaust dyebath was also pH 9. The color was practically indistinguishable from the original exhaust skein. Here are the three skeins together once they were all rinsed and dried:

It is possible to obtain much richer shades of red with madder root by making a more concentrated dye bath, but then you are looking at days and days of exhausting the bath, resulting in a small amount of reddish yarn and lots and lots of pink yarn. I tried to go easy on myself this time.

Weld–This is another pH sensitive dye plant that makes a kind of blah greenish yellow at a neutral pH and eye-poppingly bright yellows at a higher pH. We used 4 oz. of dried plant material. In an ideal world this would have had time to heat, soak, then sit around a while and steep. As it was, the plant material heated up for 45 minutes to an hour, then we strained it right away to make the dyebath. We added soda ash and calcium carbonate to get a bath of pH 8, and after all was said and done I got a nice yellow. I did not exhaust that bath.

Orange Cosmos–This is also a pH sensitive dye, which becomes more red in color at a higher pH. However, we didn’t bother to modify the pH in this workshop. We used 4 oz. of frozen flowers. Like the others, this dyebath heated up for 45 minutes to an hour before straining. The next day when I was dyeing the skein at a more leisurely ace, I left the pH alone. I didn’t even check it, so I can’t tell you what it was. The color was a pleasing tangerine, but not dark enough to bother exhausting.

Marigolds–We used 4 oz. of dried flowers to make the dyebath with no pH modification. The resulting color after heating for a full hour on Sunday was a sort of mustardy yellow or yellow-orange.

Together they make a nice range of colors. Here are all the skeins again (this is the same photos as my earlier post), minus the third madder exhaust:

From left to right: madder exhaust, first madder bath, orange cosmos, marigold, weld. I decided to use the same ratio of plant material to fiber for every dyebath, namely a one-to-one ratio. This was primarily to keep things simple for all of us. We could have made a more concentrated bath of weld, but 4 oz. of plant material filled up the pot, so that was really our limiting factor. A higher ratio of plant material would make for a stronger dyebath.

I hope some of these educators got inspired to dye with their students (or at home just for fun)!


On October 21st, 2017 I presented a workshop on growing and using dye plants with kids at the Inside-Outside Conference in Keene, NH. The conference was a collaboration of several local organizations, including Antioch University New England, the Monadnock Region Placed-Based Education Committee, the Harris Center for Conservation Education, the Caterpillar Lab, Symonds Elementary School (where the conference was held), and the Keene School District. The theme was “Promising Practices in Nature- and Place-Based Elementary Education.” You can view the full brochure here.

The audience was K-6 educators from a variety of educational settings. I don’t mention this very often on this blog, but I actually am a teacher! I co-teach in a combined first and second grade at the Common School in Amherst, MA, where I’ve been working since 2004. Most of the time, I am in the classroom doing all the usual academic things: reading, writing, word study, math, science, social studies, arts and crafts. I do fiber and dye projects with kids when I can, and the rest of the time I squeeze it in on weekends and vacations.

Back to the conference: My time-frame was 2:15-3:45 in the afternoon. This is a normal amount of time for most workshops at a day-long conference like this, even hands-on workshops. The only trouble is that all the steps in natural dyeing take a long time. If you want to make a dye bath with fresh, frozen, or dried plant material it takes at least 45 minutes to an hour, and then in an ideal world you let that sit overnight. Dyeing the skeins of yarn takes the same length of time. I always let the skeins or fiber sit overnight if I can, and I let them dry before I rinse them. Also, you have to mordant the fiber ahead of time. You cannot possibly fit it all in to an hour and a half. Nevertheless, I had committed to teach this thing. Making color with plants is so magical and so do-able that I am always happy to encourage people to try it.

So, I was excited about it, but also anxious. Basically I was counting on the fact that this would be a group of folks who are interested in process over product, and would want to see and participate in how the dye baths are made. Hopefully people felt satisfied with what we were able to do in that time:

  • make the dye baths (measure/weigh the materials, add pH amendments, test pH, set the baths to heat up)
  • talk about some considerations for setting up a dye plant garden or incorporating dye plants into an existing garden
  • look at some examples of projects I’ve done with kids
  • browse some of my favorite reference books
  • strain the dyebaths and put in the yarn

We were literally putting in the last skein of yarn at 3:45!

True to its name, we were outside for part of the workshop, and inside for part of it. When I set up in the morning, the outdoor space was shady and pleasant. A helpful custodian helped me run two long extension cords out of two windows, down to the ground on the playground below. There, I set up two portable electric burners. I set up everything we would need for dyeing outdoors, I set up books, hand-outs, and project samples in the classroom, and then headed off to enjoy the rest of the conference.

By 2:15, the sun had come around to our side of the building and it was blazingly hot. You wouldn’t have anticipated a blazingly hot day in late October, perhaps, but such it was. I think it was about 80 degrees. I had a great group of participants, about 20 folks, who obligingly tolerated the blazing heat for a few minutes while we got started. But it was really uncomfortable! I was feeling bad about it, but also feeling kind of stuck and unsure of what to do. Then, someone pointed out that it was shady just around the corner of the building. Yay! We moved things around the corner, and Ellen Doris (a former colleague at the Common School and my contact for the conference) brought an extra extension cord. We all breathed a sigh of relief in the small patch of shade behind a wall, and proceeded to set up the dye baths.

Here are the skeins drying afterwards on a rack at home:

We used four dyes for this workshop: madder, weld, orange cosmos, and marigolds. In my next post I will go into all the details of exactly how we obtained these colors!

Japanese Indigo August 2017

Way back in August I ran a Japanese indigo vat. Here’s what the bed of Japanese indigo plants looked like on August 20th:

I have only dyed with fresh Japanese indigo leaves a few times, so I am still trying to develop skill with the process. An important part of developing skill is repetition. Another important piece is learning and testing new things, and then trying to understand why they do or don’t work. Luckily, this vat afforded me all of those opportunities!

I picked 22 oz. of plant material, which yielded exactly 1 pound (16 oz.) of leaves trimmed off of the stems. Here are the tips of the plant stalks that I harvested:

On the left are the stems, and on the right is the bag with just the leaves in it. It’s a really beautiful plant! It has sweet little hairs, wrapped-around layers, exciting color contrasts, and an interesting juxtaposition of rigid and luscious textures.

I wanted to over-dye six small skeins (about two ounces each) of pale blue cotton yarns (commercial 10/2). They had all previously been dyed with woad, and several of them had gone through other processes, too. Two had been dyed in an umbilicate lichen vat, but had only become vaguely pinkish beige in that process. Five had been soaked in a gallotannin solution in an attempt to achieve a teal or blue-green color with woad. One had been in a weld exhaust bath after several dips in a woad vat. All the skeins were still disappointingly pale. I find cotton very difficult to dye!

I used the canning jar “double boiler” method again for this vat. I’ve described this process before, but I figured it was worth repeating here.

I crammed the leaves into half gallon and quart jars, filled them to the shoulder with cold tap water, and put the lid on. I set the jars inside a pot with water about three quarters of the way up the height of the jar.

Here’s all the equipment and the way the jars were arranged in the pots:

I slowly heated the pots of water over the course of two hours. One of the pots accidentally got up to 180º for the last fifteen minutes, but I was aiming for 160º. Here’s what the liquid in the jar looked like after two hours. On the left is the top of the jar and on the right is the liquid in the bottom of the jar:

This is what the leaves looked like when I opened the jars. The metallic sheen that you can see on the right is always a good sign!

Once I strained out the leaves, I had about two gallons of liquid. I used ammonia to get the pH up to 9:

The color changes dramatically with the pH shift. This is true with woad, too. But in my experience, Japanese indigo and woad don’t act or look the same way. On the left below is the greenish color that came out of the jars. Usually, extracted woad is pink or red. On the right is the way it looked after I added a lot of ammonia. The liquid is completely opaque, but now it looks sort of red. Usually with woad, the liquid turns dark green after I add the ammonia.

In this case, the photo on the left is before I added ammonia, and the photo on the right is after:

Then I aerated the liquid for about 10 minutes by pouring the liquid back and forth between different buckets. I had set the timer for ten minutes, but my back started hurting.

I decided to use thiourea dioxide for the reducing agent. I usually use Rit Color Remover, which contains sodium hydrosulfite. But, I had some thiox left over from an indigo vat in the spring, and I have heard it has a short shelf-life. I followed the recommendations from Rita Buchanan in A Weaver’s Garden of 1 Tbsp thiox or 2 Tbs. sodium hydrosulfite per gallon of liquid.

After 50 minutes, the vat didn’t really look reduced, but I stuck in a skein anyway. Usually I look for a murky greenish yellow, but I was kind of impatient!

When you dip fiber into a woad or indigo vat that has been chemically reduced, the reducing agent also functions as a color-stripper. So, if there is color on the skein already, you have to be careful not to let it sit in the vat too long. If you leave it in too long, the original color will be stripped off. On the other hand, I have found that the “quick dips” that are recommended for indigo vats using powdered indigo don’t work for me when I’m using a vat with fresh leaves. So, I left the first skein in for ten minutes.

That worked fine. I put the second skein in for ten minutes, then bumped the pH back up (it had gone down to 8) with a little more ammonia. The third and fourth skeins were in for 20 minutes, the 5th skein was in for 30 minutes, and the last skein was in for about an hour and a half.

While the skeins were oxidizing but still wet, the color was very promising!

While I was getting ready for this vat, I re-read my notes from a dyeing workshop with Joan Morris at Long Ridge Farm. I noticed that she recommended neutralizing cellulose fiber with tannic acid rather than acetic acid (vinegar) after a vat. Since the pH of a vat is very high, you are supposed to neutralize the fibers by soaking them in a mild acidic solution afterwards. I always do this with wool, since protein fibers are damaged with a high pH. Somehow I had forgotten that this was also important with cellulose fibers. And I had completely missed the tannic acid recommendation. So, I thought I’d better look into it.

A little bit of poking around on line led me to Catherine Ellis’ blog Natural Dye: Experiments and Results. What a fabulous resource! I found this post about over-dying with indigo especially interesting.

I had two kinds of tannins at that moment (not including black tea–my favorite is Barry’s gold blend, which I would rather drink than use for dyeing!). I decided to use Earthhues gallotannin, which is very light.

I always wait until the fiber is completely dry, if I can, before rinsing, so it took a couple days to get to the neutralizing and rinsing stage. I dissolved 1 tsp. of gallotannin in about 2 gallons of water (a dishpan) to soak all the cotton skeins before rinsing. The pH was between 7-8. That seemed weird for something called tannic “acid”. In case you need a refresher on your pH scale, 7 is considered neutral, and anything below that is considered acidic. Anything above 7 is considered alkaline. Adding an acid to water ought to make an acidic solution. Or so I thought.

I tested the pH of the plain, hot tap water. I got a surprising pH 8-9. What? I was shocked. I checked around, and apparently it is not unusual for the pH of tap water to be this high. So much for the notion that water is “neutral”.

Soaking in the not-acidic pH solution with tannic acid didn’t help much. A lot of color rinsed out from the cotton skeins. Perhaps I should have made a stronger solution with tannic acid. I was reluctant to make it too strong, though, because tannins can also shift the color to a duller or browner tone. I didn’t want the blues to get muddy.

I did a second soak and rinse with laundry detergent in hot water, but color was still rinsing off. So, I made a new solution with acetic acid (white vinegar) to make a mildly acidic bath of pH 6, and soaked all the skeins in that. It seemed to do the trick.

I clearly need to do more reading about why tannic acid is better for cellulose fibers. Meanwhile, the dried skeins are a *bit* but not a *lot* darker than before. I am not sure it is worth it to me to continue banging my head against cotton yarns.


Historic Eastfield Village 2017

On Saturday September 23rd, I demonstrated the flax-to-linen process at Historic Eastfield Village’s Founder’s Day celebration. It was a lovely day! We had a heat wave later that week, but under the oak trees that day it was pleasantly cool and shady.

I brought dried flax stalks with the seeds on, retted flax, and all the tools to break, scutch, and hetchel the fibers. I also had some commercial linen yarns that I dyed with madder, weld, woad, and black walnut.

Historic Eastfield Village is a very interesting place. You can read more about their history, buildings, and classes on their website. Last year, I attended Founders Day with Lisa Bertoldi, on the invitation of Niel DeMarino of the Georgian Kitchen, whom we had met at the Flax and Linen Symposium in August 2016.

This was my favorite image from Founders Day 2016:

It really captures what motivates me, and what I aspire to in my own knowledge and skills. Not that I am always successful! Still, words to live by.

Last year I got to wander all over the site and take photos of various artisans and craftspeople at work. Here’s the printer, for example:

This year, I didn’t get to see all the other demonstrations and vendors at work. But I did get to talk to a lot of interesting people!

Early in the morning, things were slow. So, I started breaking flax to see if I could lure people over to the tent.

It worked! Some of the other demonstrators and living history folk came over to visit.

This fellow, Henry Cooke, is a well-known 18th century tailor, but was there in his capacity as a militia member from the early 1800s. He was at Historic Deerfield the following weekend, as part of their Historic Trades series.

And there were lots of regular non-historical people, too:

In this photo I look like I am telling a tall fish tale. “It was this big, I swear!” Actually, I am describing the height that fiber flax can reach under ideal conditions:

Here I am explaining the dyes I used to make these colors:

Matthew took some better crowd shots, which you can see on the New England Flax and Linen‘s recent Facebook post.

Electra Update Part Two

As I mentioned in my last post, this is a “retro time” account of my flax harvest this year, not a “real time” account. Here’s the belated next installment.

I started digging up the Electra plot on July 31st. I didn’t finish until August 12th. Now it’s all pulled up, dry, and stored safely in the back of the van. Because that’s where the flax gets stored.

The yield was small but the effort was mighty! I could only work for a couple hours a day, and some days I didn’t work at all. This summer taught me a profound lesson in the privileges and assumptions I have carried with me all my life as an able-bodied, pain-free person. My motto used to be, “Do all the things!”* This summer, not so much.

On August 4th, Lisa and Carolyn from my flax and linen study group came to help with my flax harvest. Three people work much faster than one!


Here’s our haul after a couple hours. We were very happy with this lovely pile, and celebrated with Chinese food.

Granted, under normal circumstances, this wouldn’t be a lot to show for that much work. However, since I wasn’t able to weed earlier in the summer, most of the plants we were removing weren’t actually flax. It was more like mining or excavating for flax. A lot of rubble punctuated by gleaming moments of excitement.

To maximize the learning opportunities (and to slow this whole process down to the speed of cold molasses!), I sorted most of the bundles according to stem thickness, height, and branching pattern. Why on earth would I do such a thing? Well. I often have questions regarding what I read or hear about flax processing. For example, I have heard that thinner stalks produce finer fiber. I have also heard that thinner stalks take longer to ret. I have also heard that branching at the base of the stalk is not desirable, even though it’s so close to the root that I can’t imagine it makes a difference in the length of the extracted fiber. Many times, I have wished for and wanted a way to prove to myself whether these things are true or not. Hopefully, my sorted bundles will let me test out some of these questions.

* N.B. My motto was inspired by this brilliant artist. Neither myself nor the artist, apparently, is good at being a grown up. Like her, I have embraced my hatred of going to the bank and cleaning.  I very seldom do either. Ha! Since I am resigned to my fate, I decided that the “things” in my motto are the things I want to do. If I’m going to put a lot of energy into something, it will be my personal obsessions and desires (plants, fibers, stinky creative messes, and saving obscure knowledge from obsolescence…).

Electra Progress Report Part One

I had meant to post updates about my flax crop this summer in “real time”. However, “retro time” will have to do.

Here are a few things that I observed and learned as the 2017 flax was growing and maturing.

First, the flax chewers who devastated my crop in 2016, and half of my crop in 2015, were back at it again this year. However, when you have 1500 square feet of the same variety (Electra from Biolin), rather than tiny test plots of 12 square feet or less, the effect of the damage isn’t as troubling. I found dozens of chewed up flax stalks, but it was a negligible percentage of the whole crop. I am sticking to my hypothesis that the culprits are rodents of some kind. Here’s some scat that may or may not belong to them:

Second, the chewers are not solely interested in flax. It might not even be their favorite or preferred plant to chew. The fact that flax is *my* preferred plant in that location means that it bothers me when they chew it. I don’t care about the other plants, so I’m less inclined to notice their demise. Predation of “weeds” is a boon, from a flax-grower’s point of view. But it’s possible that from the chewers’ point of view, it’s the flax that’s a nuisance.

The two most significant weeds in my plot were lambsquarters and campion. Since apparently most people spell that “lamb’s quarters” I guess I should adopt the convention. I found chewed stalks of both. Here are a few lamb’s quarters plants that had been chewed as they were going to seed.  On the left you see the entire stalks, and on the right a close up of the stalk where it was chewed. The diagonal angle is similar to the way the flax is chewed:

Campion plants were also chewed as they went to seed:

For comparison, here are some chewed flax stalks:

The pillowcase is there for contrast and scale. Yes, the lamb’s quarters were as tall as (and in many cases taller than) the flax.

Third, I can understand why campion is so hard to weed out. It had all gone to seed by the time I was harvesting the flax. Every time I pulled up a campion plant, I had an image of a million baby seedlings sprouting from my hand as I flung the plant aside. Basically I was sowing next year’s campion crop. This is the number of seeds from one pod. Every plant makes lots of pods….

Fourth, not everyone knows what lamb’s quarters are. I was describing to someone how difficult it was to pull up my flax this year, and made reference to how many “lamb’s quarters” there were in the plot. They asked enthusiastically, “How much is that?” It had never occurred to me to think of the name as a unit of measurement, but now I love the idea! It does sound like a old fashioned measurement term.

The plant to which I am referring is a Chenopodium album. I went looking for the etymology of the name lamb’s quarters, but I’m not sure if I got to the heart of it. One source refer to Lammas, a harvest celebration on August 1 to bless the first loaves of bread. Personally, I am not sure how Chenopodium album fits into the picture. The seeds from C. album are edible and nutritious, so maybe they were an ingredient in a harvest-time loaf. I have never eaten the seeds, but I have eaten my fair share of lamb’s quarters. They are a delectable green vegetable, even more nutritious than the seeds.  But once the plant has gone to seed, the greens are well past their prime. In fact, they shrink up and often turn pink and fall off. Here’s another interesting article about lamb’s quarters. I may have to follow up on this question with better information.

Anyway, it isn’t a unit of measurement, it is a plant. A very useful and nutritious edible plant. It just so happens that it can grow as tall as me, and twice as tall as a flax stalk, and digging it up is very difficult.

Here’s what it looked like relative to the flax:

And here is a lone flax stalk surrounded by campion and lamb’s quarters:

Lastly, there is a lot more going on with regard to the relationship of insects to flax than I have any idea about. What are these creatures doing?

They were on the bolls as they were ripening. My guess is that they’re either eating them or laying eggs inside them. At the time I didn’t investigate further, but it might be important knowledge to possess.


Green Yarn

This has been an extremely prolific year for Queen Anne’s Lace, also known as wild carrot or Daucus carota. It is absolutely everywhere!

Back in July I ran two dyebaths with fresh Queen Anne’s Lace flowers. Since it’s so abundant, I decided to just use the flowers this time, though you can use the whole plant. For the first dyebath, I had no trouble collecting 30 oz. of flowers from various spots around Amherst, including the sides of parking lots, the side of the road, and next to bus stops.

The flowers are incredibly fragrant and sticky, and consequently they host a huge range of insects. When you pick the flowers, all the insects come along, too. This fact gave rise to a new house-hold rule:

I weighed the plant material outdoors! I also made the first dyebath outside on the portable electric stove outdoors. We had some rainy weather after that, so I made the second dyebath indoors using 24 oz. of flowers that I picked in Hadley.

Here’s a pot full of flowers:

Here’s a close-up. It’s a really beautiful plant:

For the first dyebath, I filled the pot with water, heated it to 140 degrees, maintained that for an hour, and then let the plants soak in the pot overnight. The relatively low temperature was due to the fact that my portable electric burner has two rings. One of them can get very hot, but the other only has a “simmer” setting. The Queen Anne’s Lace was on the simmer side, while I mordanted yarn on the other burner.

Here are the strained flowers after they were heated, soaked, and cooled:

The dyebath looked reddish in the pot, but when I put some of the liquid in a jar, it was light gold. The little white dots are flower petals and maybe pollen that didn’t strain out.

For this project, I decided to over-dye some blue woad-dyed woolen yarn from last summer. I hadn’t bothered to mordant the yarn for the woad vat originally, so I had to mordant the skeins before overdyeing with Queen Anne’s Lace. I used aluminum sulfate at the ratios recommended in Rita Buchanan’s A Dyer’s Garden (1 tablespoon per 4 oz. of fiber). It looked pretty funny to put the blue yarns in a pot with clear water:

To mordant wool with aluminum sulfate, pre-soak the scoured yarn in water for at least an hour in a separate tub. Dissolve the mordant in a pot of hot water, then add the wetted out yarn. Bring the temperature up to 180 degrees, maintain that for an hour, then shut off the heat and let the fiber cool in the mordant bath as long as possible. In this case, it cooled overnight.

The next day, I put a 6 oz. skein into the first Queen Anne’s Lace dyebath, heated it to 160 degrees, and kept it between 160-180 degrees for an hour. I let it cool from 10 a.m. to 4 p.m., then pulled it out to dry before rinsing it. I got a very nice shade of green!

Here’s the skein while it’s still in the dyepot, shown with another skein of the same, original shade of blue for comparison. Colors are always darker when they are wet:

Here’s the green skein dripping and drying outside, amidst all the other gorgeous greens of July:

I used the first dyebath again to over-dye a 3 oz. woad-blue skein of wool, and got more of a slate shade of green (less yellow, more blue). It’s in the center in the photo below. When you use the same dyebath again it’s called “exhausting” the bath, and usually results in a lighter color. After the exhaust bath, I poured out the liquid.

The second dyebath I made a couple days later wasn’t quite as strong, only 24 oz. I heated it to 200 degrees, maintained that for an hour, and cooled it overnight. Again, I got a beautiful shade of green on a woad-blue skein. Here are the three skeins once they were all rinsed and dried:

I used the exhaust bath from the second dye bath to over-dye 7 oz. of mohair. It was an extremely pale gray-blue from an exhausted woad vat last summer. I ended up with a sort of pale silvery gray. which was not what I was expecting. In the photo below, there’s a light-yellow lock on the top left corner that shows what the color would have been if the mohair wasn’t already gray-blue.

There is still plenty of Queen Anne’s Lace blooming now that it’s mid-August, but I may turn my sights to other plants next. Goldenrod, perhaps.

Past Speaking Engagements

Over the past year, I have had several opportunities to demonstrate flax processing and talk about natural dyeing. Here is a quick summary of four events that I didn’t get around to writing about when they happened. I just want to document and share them before too much more time passes.

Last August (2016) I did a flax processing demonstration at the Amherst History Museum, in conjunction with the art exhibit “Artifacts Inspire” by the Fiber Artists of Western Massachusetts. The museum asked the participating artists to create original works inspired by objects in the museum’s collection. Two of the pieces in the show were created by Martha Robinson, inspired by two antique hetchels, which are flax processing tools. There’s a good photo of one of her felted pieces here. It was great fun to show people how flax was processed in the past, and to let folks try their hand at using the tools.

Here’s a shot of crowd at the beginning of the demo:

Here I am by the brake and the scutching board:

And here’s Marianne, their consulting curator, getting a kick out of using a hetchel:

The next gig I wanted to mention was my presentation to the Weavers Guild of Springfield on March 4, 2017. I showed a slideshow about planting, growing, harvesting, and retting flax:

Then, I did a quick demonstration of how to use the tools:

It was lovely to meet a new group of weavers, and inspiring to see some old acquaintances there, too.

The third event I wanted to note was the FIBERuary panel I was part of on Feb. 19, 2017 at Sheep and Shawl in Deerfield. FIBERuary is a relatively new event here in Western MA celebrating our local fiber farmers and fiber artists. It’s spearheaded by Carole Adams of Whispering Pines Farm in Colrain, MA. In the past two years it has included a month-long blog and speaker series at Sheep and Shawl. On our panel, we addressed dyeing natural fibers from three perspectives: Linnie Dugas of Woollies of Shirkshire Farm talked about dyeing wool with natural dyes, and brought some luscious dyed batts and roving. I talked about natural dyeing skeins of linen. Scott Norris of Elam’s Widow talked about his process using Procion fiber reactive dyes to dye the linen yarns he uses in his spectacular handwoven kitchen towels.

Last but not least, I was a presenter on a panel at the Fashion Institute of Technology’s Sustainability in Textiles Summer Institute in New York on June 7, 2017. Our panel was called “Local Fiber Connections” on the theme of “Farm to Fashion”. The other panelists were Jeffrey Silberman, Chair of the Textile Development and Marketing Department at FIT, Mimi Prober, designer, and Sara Healy of Buckwheat Bridge Angoras. My portion of the panel was a slideshow about retting and processing flax, and basic information about spinning and weaving linen. Sara has worked with Mimi to create custom blended batts for felted garments in Mimi’s collection.

Jeff, Mimi, some other folks at FIT, myself and other members of the New England Flax and Linen Study Group are working on a Farm to Fashion project in which we are collaborating to grow and process flax, spin and weave it, and produce garments for a runway show! At this point, the flax is still in the field, but it’s an exciting prospect.


Swamp Milkweed Sightings

I first learned to identify swamp milkweed (Asclepias incarnata) in 2012 after discovering some lovely fibers near my sister’s apartment in Maryland. In 2015 I acquired some plants from Nasami Farm in Whately, MA for the Common School‘s fiber and dye plant garden at Bramble Hill Farm. For all this time, I have been keeping an eye out for it “in the wild” but haven’t seen it. Until now!

This month I have been spotting swamp milkweed all over the place. The first place I noticed it was in the bluebird field at Amherst College on July 6th. Admittedly, these photos are a bit like photos of Big Foot: blurry and indistinct. Trust me, though, it is swamp milkweed!

The next place I caught a sighting was in the Lawrence Swamp area of the Norwottuck Rail Trail in Amherst. It was right in the swamp, aptly. We could see several plants further out, but ran into the same blurry Big Foot photo problem. This one was close to the edge of the trail:

Continue reading “Swamp Milkweed Sightings”