Why would a conservator want to live in a town of 30,000 that is not on the road system and a three hour flight to Seattle?
Why would a conservator want to live in a town of 30,000 that is not on the road system and a three hour flight to Seattle?
A visit to the Alaska State Conservation lab, January 27, 2009. There are two main areas, we call them the “wet lab” and the “dry lab.” Until just a few years ago, the entire conservation operation was located in the wet lab area.
A list of irons in the fire in my lab:
1. PEG BASKETRY RESEARCH: Ready to put basketry fragments in the freezer in the final step before examining how the samples held up. They are (likely) spruce root basketry fragments, perhaps a few hundred years old, from a waterlogged archaeological context. All we treated with 20% PEG 400, half at room temp and half heated. Then I tried various amounts of PEG 3350 to test my hypothesis that poor results reported for some PEG treated basketry might be from too much low mw PEG and not enough high mw PEG. I plan to publish the results, as well as info about other PEG basketry treatment done at the Alaska State Museum, but it would be much more fun if this goes well. I do believe there will be valuable information if there is NOT success, but it will be less enjoyable to write up and more of a duty thing. Incidentally, if you’re reading this and have dabbled in PEG, I’d love to chat with you.
2. PEG BASKETRY CONSOLIDATION: Some of the oldest archaeological basketry on the Northwest Coast (5,000 year old) is in the Alaska State Museum collection, but the PEG treatment done for it years ago was only partially successful. The fragments are preserved and look nice, but they are fragile and are rather like fibrous pieces of dried meat. I think careful consolidation with Butvar B-98 applied with a nebulizer might do the trick, allowing them to be exhibited and perhaps even travel.
3. BASKETRY SURVEY: Begun in 2007 as the first of a series of materials-based conservation surveys of the collection. Almost none of the ASM’s collection has condition descriptions in the database, and a large percentage of the collection is not photographed. The survey is capturing this basic information, as well as providing interesting insights and an inventory along the way. Survey is approximately 20% complete, but I have offered to give presentations at the “Alaska Anthropological Association” conference and the Tlingit clan conference in Juneau this March, so this might kick into high gear soon of my proposals are accepted.
4. NATURAL HISTORY SURVEY: Begun in 2008 with a consultation with Catharine Hawks which was incredibly helpful. The natural history collection is rather small, and has been somewhat neglected for years, both in terms of storage conditions and curation. The herbarium contains more than 5,000 specimens dating back into the 1920’s. The geology collection is undoubtedly of some scientific value, even without the boxes and boxes of well-identified specimens sitting on the floor of my lab because they’ve never been added to the collection. The marine shell collection might be significant, I am not sure. There are hundreds of taxidermy birds, which I am arsenic testing at the moment. Some of those are almost 100 years old and indeed the vast majority are positive for arsenic. Small but unmolested collection of fossils, including many mammoth teeth. This survey is about 25% complete.
5. SHIPWRECK ITEMS FROM THE TORRENT
Twelve small plastic tubs of artifacts from the 1868 Torrent shipwreck are currently desalinating in the lab. Munitions,metal items, ceramics, and glass mostly. A bronze howitzer will be contracted out and is still in the lab of state archaeologist, Dave McMahan. So are several portholes, and an unusual ceramic and lead toilet. Shipwreck items and waterlogged archaeological items regularly make their way to the Alaska State Museum. I’m fascinated by the demands of these materials, but equally fascinated by the diverse approaches and strained history between AIC-affiliated conservators and the majority of conservators treating this kind of material who have been trained at the Texas A&M program. A can of worms I’m sure, but I want to know more…
6. FUR ID PROJECT
More info in a previous posting. This is the project I am enjoying the most right now.
7. PARKA TREATMENTS
Tear repair needed for a child’s parka made from baby caribou and another parka made from marmot skins that had been turned into a bomber-jacket style around WWII. I plan to try Reemay and BEVA with a heat spatula.
8. GUTSKIN TREATMENTS
Three items made of gutskin, probably seal, came as part of as part of a recent acquisition. Two were decorative bags, and then this fabulous hat. They are thought to be Aleut or Alutiiq, and the rim of the hat in particular needs some humidification in order to be exhibited. Reference literature suggests the welting is made of sea lion esophagus.
There are also a pair of child’s boots from the Sheldon Jackson Museum, possibly Aleut, that are made of some inner membrane of a marine mammal. Possibly esophagus? How to distinguish between all these stomaches and guts and bladders and windpipes and the like? Two graduate students have separately contacted me about this kind of material identification: Linda Lin at the UCLA/ Getty program working with Ellen Pearlstein and Amy Tjiong at the NYU Conservation Center working with Linda Nieuwenhuizen. I’m hoping to stir up some collaboration. Stay tuned for a blog musing on the issue…
9. STORAGE ISSUES
With no collection manager on staff, I have to pitch in with storage solutions. Fine art has long since outgrown its digs, and I’ve been looking into ModuPanel kits as the solution, designed by Peter Diemand at Biblio Design and now sold by a number of vendors. We’ve also got watercraft, vintage snow machines, small airplane engines, houseposts, a mammoth skull, salmon canning equipment and other oversize items that could really use some nice big rolling shelves without pesky cross-bracing. I know other institutions have done it, but finding a source has proven elusive. Any ideas, folks?
10. IN-SITU DEHUMIDIFICATION
Here’s the problem: in a water emergency (and so many of them are) salvage and triage guides all talk about freezing your paper-based collections within 2-3 days to prevent mold growth, and then using a freeze-drier to recover them. As far as I know, there is not a place to freeze-dry materials like this anywhere in Alaska. Furthermore, my inquiries in Juneau have not come up with a way to even freeze more than a couple of boxes of wet material. There are refrigerated containers that could be rented from various barge companies, but only if one is available. Apparently, in the summer all of them are booked solid with shipments of fresh fish and beer from the Alaskan Brewery. Juneau is not on the road system. A lot of towns in Alaska are not on the road system. Is in-situ dehumidification our answer? My understanding is that the method uses a large truck to suck moist air out of an interior space using big flexible hoses, dehumidifies it in the vehicle, and then pumps the air back in over and over until the entire space and everything in it is dry. Is this method possible for remote places in Alaska? Could we set up something where the guts of the operation could be flown in and rigged up anywhere? That would be tremendous. Maybe someone with BMS-CAT or another one of the major disaster response service providers might be willing to work with us on this…am I crazy to hope for this?
2. I’ve been using cheap plastic test tubes, bought in bulk from Fisher Scientific, and when I sample I use two swabs, each one going over the entire specimen in all the likely spots. Break off the swab ends, put them both directly into the tube. No intermediate ziplock bag to label and throw away.
3. Take your “known negative” sample right there with the same distilled water so you can have lots of confidence in your negative.
PREPPING THE TEST
1. Cut up Parafilm to make lids for the test tubes. The Parafilm I’ve got has squares printed on it and cutting them in quarters is perfect.
2. Put on gloves. The mercury bromide test papers are not healthy.
3. Cut up test papers in 1/8 pieces…four strips lengthwise and then each in half. Then snip a slit in the end and fold to make a little propeller-like thingy that will prevent the strip from falling into the tube. I call these “helicopters” and I make a bunch ahead of time and keep them in a beaker with a watchglass for a lid.
4. Use one empty test tube as a model to make all the lids you need using the squares of Parafilm you’ve cut, slit with scalpel, insert a “helicopter” into each one.
5. Set your tubes in a rack in the fume hood, set the little lids in front of each
6. Make a reporting sheet for your results
7. Tape down plain paper strips to the rack to keep it clean. Remove the topmost sample swab from each beaker using a straight-beaked tweezers and set it on the paper right in front of the tube it came from. Make sure other swab is fuzz-down in the tube. In this way, you’ve got a backup swab but you’re being super efficient about making not making more mess and labels.
8. Run your known negative and a known positive EVERY time to give confidence in results. I’ve been saving the spare swab from strongly positive specimens to use as my known positive in future tests.
9. Have all reagents ready to go. The reaction makes a gas as soon as all the components come together, and that is what the test paper is sensitive to. You’ve got to have all your ducks in a row to get that lid on quickly. The acid dropper that came with the acid in the Merckoquant test kit is annoying. Remove it with straight beaked tweezers, slowly using one beak to lift gently around all sides. I use plastic pipettes for the acid and base and little glass beakers to hold them when not in use. Acids and bases scare me, so I have a fanatical little routine of diluting and rinsing and throwing away the pipettes after a few rounds of testing.
10. Put on goggles
11. Put 2 drops KOH in each tube Leave tubes in rack while doing this. We got our 1 molar solution made up by the local pharmacist. NOTE: this version of the test DOES NOT WORK if you don’t use the KOH. This is a major difference between the Hawks method and Odegaard et al.
12. Add 7 drops HCl to each tube. Keep careful track when reloading the dropper if you run out partway through counting your drops. Leave tubes in rack while doing this.
13. Hold the rack with one hand and gently rattle tubes in the holes with the other. This just makes me feel secure that all the liquids are getting on the swab OK.
14. Lift out each tube and add scoop (approx 0.5g) of zinc dust. I find the coordination works better if you are trying to bring two hands together than if you’re trying to hit the target with one hand and the tube is in the rack. It also makes the next step easier. Since I had an old Merckoquant test kit, I’ve been using that nice scoop, but if I didn’t have it I would scheme a way to have the right quantity of zinc ready ahead of time. Adding the zinc makes the reaction happen.
12. Quickly but without panic, grab each lid by one corner and poke the paper into the tube, squeezing down the edges of the Parafilm to seal the lid in place, set back into rack.
13. Contents should be bubbling vigorously. If they are not, something is wrong. Bubbling has nothing to do with a positive or negative result, it just means the test is running properly.
14. References suggest results are reliable after 30 minutes. I usually wait overnight, just for convenience.
REPORTING THE RESULTS
15. Dipping the paper in tap water and immediately holding it up to plain white paper helps a lot to see faint yellow positives.
16. The test is not known for false positives. If your negative control did not turn and your positive did, you ought to be OK.
17. The degree of color change is not indicative of the amount of arsenic on the sample. Your sampling technique or the application method of arsenic over various areas are examples of varying factors that make this test qualitative and not quantitative.
18. The “helicopters” of test paper are hazardous waste. The rest of the materials are not, particularly if you only throw them away 10 at a time and you keep your extra positives for future testing. With the Odegaard et al method, you’ve potentially got a bunch of contaminated water to deal with.
TROUBLESHOOTING AND NOTES
I’ve gotten positive results on really small specimens, including a hummingbird that was only about an inch square.
Things that test positive for arsenic can also have insect infestations. A bird mount that tested positive had a previous infestation in the feet. Tons of larval casings and frass. Cathy Hawks tells me that feet were generally not treated on traditional bird mounts and have a bit of meat left intact which can be attractive to insects.
I am still investigating, but it seems that a number of specimens in our collection that were freeze dried are coming up positive for arsenic. Cathy Hawks told me that the craze for freeze drying really got going after arsenic was going out of favor, but I guess that doesn’t mean that the two are mutually exclusive.
Once, I accidentally ran my two swabs at once, using only the normal amount of reagent. The result still came out positive.
UPDATE January 18, 2011. Cathy Hawks just sent me a link to an arsenic test kit on the website for Gallade Chemical:
Cathy Hawks generously gave me permission to post the version of the arsenic test she uses.
An improvement on the Gutzeit test for arsenic is a modification of the Merckoquant (EM Quant) Arsenic Test kit, which was formerly supplied by EM Science. The kit was designed to detect the presence of arsenic (3+ or 5+ valance states) in water, soil extracts, pharmaceuticals, prepared biological materials, and liquid foods. Unfortunately, the kit is no longer available.
Kits can be created using chemicals commonly found in labs (1M or 1N KOH [potassium hydroxide] solution, zinc dust, small quantity of concentrated HCl [hydrochloric acid]) and by purchasing test strips impregnated with mercury bromide to detect the arsine gas evolved in the process.
The test strips are available from:
http://www.sigmaaldrich.com/catalog/search/ProductDetail/FLUKA/83544 or http://www.ctlscientific.com/cgi/display.cgi?item_num=90762 and probably from other suppliers. The procedure given below could then be used for testing.
Always wear a lab coat, a rubber lab apron, safety goggles, and nitrile gloves while doing the tests. Conduct the tests in a chemical vapor hood or in a well-ventilated area. Never attempt to test more than 10 samples at a time.
Take samples using cotton-tipped swabs dampened very lightly with deionized water. Roll one side of the swab gently on the surface of specimens/objects, over as much area as possible. When dealing with taxidermy mounts and animal skins, concentrate on the areas around the eyes, nose, mouth, ears, ventral suture (if present), base of tail, and bottom of feet. Collect samples from each object/specimen on at least 3 swabs.
Always conduct the tests using a negative control—a fresh cotton swab tip, dampened with deionized water. As long as there is no discoloration of the test strip over this swab, the results for the actual samples should be valid.
The test is designed to detect tri- or pentavalent arsenic by converting the arsenic to arsine gas. This is accomplished by adding zinc dust to the samples dissolved in KOH, then adding hydrochloric acid. The reaction between the metal dust and the acid generates hydrogen gas that combines with arsenic in the samples to produce arsenic trihydride (arsine, AsH3), a very toxic, colorless gas with an odor of garlic (consequently, it is important to do the test with closed caps and to do it in a well ventilated area).
The arsine gas reacts with the treated potion of the test strip to produce a color change that is indicative of the presence of arsenic in the sample. The treated portion of the strip contains mercury bromide, which reacts with the arsine gas to form a colored compound. The test should be considered to qualitative (i.e., a positive or negative test). Used in the way described, it is not reliable for quantitative measurements.
While the strips purportedly give a quantitative indication, the concentration is largely irrelevant for most museum objects because arsenic salts were never applied evenly, and the concentration in the sample is an artifact of the sampling procedure, rather than a reflection of the overall concentration of arsenic on the objects. Consequently, the test results should be regarded as simply positive or negative for arsenic. If test results are equivocal (it is uncertain whether there has been a color change on the test strips for at least 2 of the 3 test samples), repeat the tests with fresh samples.
When the tests are completed, push the test strips down into the vials, collect the used vials in a sealed, heavy-weight, ziptop, plastic bag. The bag can be disposed of in ordinary trash because the amount of arsenic involved in 10 or less samples does not constitute hazardous waste (at least, based on testing to date), and because the potassium hydroxide helps to neutralize the acid.
Since 2001, I have supervised more than a dozen interns and even more volunteers. I’ve come to a few conclusions. One is that the balance of supervision invested versus useful work produced cannot be met without a significant time commitment from both parties. For me, this has either been a minimum commitment of one day per week over a long period, or a concentrated chunk of time where the person comes in almost every day. In order to utilize interns and volunteers well, the supervisor needs to have a certain level of organization already in place. Otherwise, the time is simply spent facilitating work I could have done faster myself. Most of my supervisory experience took place when I was a curator of collections and exhibits. The interns were mainly grad students from the Texas Tech Museum Studies program. The most difficult aspect of supervising them was striking the right balance of adequate instruction and appropriate correction. I have no stomach for providing criticism. Written manuals of museum procedures are most helpful, particularly for processing collections. With the volunteers, who were mostly retired local folks, it was more delicate because few of them had computer skills, and volunteering is partly a social activity. However, most of them had deep and valuable knowledge about community history. It was most helpful for me to schedule them all on the same day of the week. This way, I could spend some time the day before preparing everyone’s projects and lining up my own questions for them. The volunteers mostly knew each other and enjoyed each other’s company.
In 2007, I supervised my first conservation graduate students, Samantha Spring from the Delaware program and Molly Gleeson from the UCLA/Getty program. An article about the experience can be found at (forgive me, I’m just learning links!):
This was a very rewarding experience for me, since we all came from similar educational backgrounds and could cover considerable ground quickly. They had knowledge of AIC standards and guidelines for practice, basic ethics, and knew the components of a treatment report. The exchange of knowledge and skills was a two-way street, with the students providing the latest information from their training and me providing the pragmatic “what they don’t tell you in school” and some meaty projects. They presented a paper about their work at the ICOM-CC conference in New Delhi India in September 2008.
I’ve found it rather difficult to come up with a reliable barometer about how well an intern or volunteer will work out. For a while, I thought that young people who were not yet in grad school didn’t take the experience seriously and were flaky. But I was proven wrong on that a couple of times, thanks to Katie Mahoney and Dean Duryea, Jr. And the best resume I’ve ever seen ended up being a poor fit. And now that I am focused mainly on conservation, I thought perhaps I should stick only to conservation grad students. But I’ve got a volunteer, Sadie Beck Ingalls, who is so sharp I could totally use her full-time, and I think her educational background is a recent Greek and Classical Studies degree.
I would love to have an intern around most of the time…so I am entertaining the idea of a summer intern and perhaps a third-year conservation grad student. The main difficulty is money. There isn’t funding from my end, although conservation students often have some limited resources if the want this particular experience bad enough. Juneau is an expensive place to live, and since it is a summer tourist destination, living accomodations are tricky. On the upside, my boss, Bruce Kato, is very supportive of conservation and the projects we’re tackling. And Juneau is a fantastic wonderland.
I really believe in the intern/mentor relationship. I love seeing things anew and critically through the eyes of a grad student. I like being challenged and questioned so I don’t get complacent (there are only 4 conservators in all of Alaska and I am married to one of them.) And I hope that these relationships are destined to turn into a colleague-colleague peer thing.