Diaphonization / Clearing And Staining Protocol 3o2b52
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Diaphonization / Clearing and Staining protocol
I’m on YouTube! To watch this protocol in live action, click here: https://www.youtube.com/playlist?list=PLE0GkpL0JFYM3eNOuD7CCmcFlb0LQ4g Or scan the following QR code:
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Preface Diaphonization (also known as clearing and staining) is a preservation technique in which the tissues of a specimen is made transparent, and the cartilage and bones are made visible by use of specific dyes. Diaphonization is nothing new – in fact it has been around since before the 1940’ies, where the process enabled zoologists to study the bones while still inside the animal. The flexibility of the specimens even allowed them to study these bones in movement, and it was therefore a powerful tool in order to better understand bone function. While this technique is still used in zoology, there has been a recent spark of interest in diaphonization outside the zoology field due to their visual appeal. Diaphonization is now also part of the taxidermy community, and diaphonized specimens are being sold as artsy display items worldwide. Part of this comes down to the simplicity of the technique; diaphonization is a pretty easy and forgiving process that is possible to do with relatively few remedies and without advanced lab equipment. This means that – in theory – this could be done in the comfort of your own home. While there are a lot of protocols out there, they can be hard to find and navigate without a scientific background. The idea of this protocol is therefore to help bring diaphonization to the layman. About me: I have a background in science (biochemistry) and spend a lot of my time in a lab. I started diaphonization as something fun to do at home, and thought I would share it with the world. While I do have experience in general lab-related tissue/cell dyeing, diaphonization is still new to me, and I am in no way a professional. If you find something gravely wrong, or anything else that could be improved, please don’t hesitate to me J
Table of contents Introduction Materials A ‘quick’ note on safety Calculations F.A.Q. References
3 3 4 5 6 6
Protocol: Experimental overview Part 01: Preparing the specimen/skinning Part 02: Fixation Part 03: Washing Part 04: Cartilage staining (Alcian Blue) Part 05: Rehydration Part 06: Trypsin digestion Part 07: Bone staining (Alizarin Red) Part 08: Clearing Part 09: Results
7 8 9 10 11 12 13 14 15 16
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Introduction This is the boring part, but it needs to be here. It has a list of the materials needed, but it also includes some words about safety when working with these types of chemicals, and a section on how to do some typical lab work calculations that you might need during this protocol.
Materials LIST OF CHEMICALS NEEDED: Name
Formula
State of matter
Distilled/deionized Water
H 2O
Liquid
10% Formalin
CH2O • H2O
Liquid
99% Glacial Acetic Acid
C 2H 4O 2
Liquid
95% Ethanol/Ethyl Alcohol
C 2H 6O
Liquid
Potassium Hydroxide
KOH
Flakes
Sodium Borate (Borax)
Na2B4O7
Powder
3% Hydrogen Peroxide
H 2O 2
Liquid
Thymol
C10H14O
Crystals
Glycerin/Glycerol
C 3H 8O 3
Liquid
Trypsin 1:100, 25g
Powder
Alizarin Red S
Powder
Alcian Blue 8GX
Powder
Notes
93% denatured ethanol will work just fine too
Optional
LIST OF MATERIALS NEEDED: Item
Notes This protocol is mostly suited for small vertebrates like mice – some animals may require tweaking of the protocol in order to work. Frozen feeder animals are good to start with.
Specimens Surgical kit for dissecting
Get at least 1 decent pair of scissors + forceps
Graduated cylinders
A set of e.g. 100 mL, 20 mL and 5 mL will get you a long way
Scale + weighing paper + spatula
You want a scale that is able to weigh down to 0.01g
Jars/containers
These are used throughout the protocol, so you’ll want a good amount of these in various sizes. Old food jars are fine as long as they are cleaned well
Safety
You want gloves, a mask and some googles
Waste containers
Get some containers for your waste liquids, so that you can dispose of this properly (check your local waste rules)
WHERE TO GET EVERYTHING: Sorry, can’t help you there. This is the hardest part, and depending on your country of residence this can be a real struggle. I’ve had some good luck on Amazon and eBay/AliExpress though.
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A ‘quick’ note on safety Some of these chemicals are quite nasty – both for the body and the environment. It is always a good idea to know your chemicals before you start working with them. Detailed safety sheets should come with the chemicals when you buy them, but here is a quick summary: Chemical Formaldehyde/formalin
GHS symbols
Safety measurements Wear gloves, mask, & eye protection. Work in wellventilated area. Can cause cancer and genetic defects. Do not breathe in! Wear gloves, mask, & eye protection. Work in wellventilated area
Potassium hydroxide (KOH) Borax
Wear gloves. Work in wellventilated area/with mask.
Glacial Acetic Acid
Wear gloves, mask, & eye protection. Work in wellventilated area.
Alizarin Red S
Wear gloves. Work in wellventilated area.
Alcian Blue 8GX
Wear gloves, mask, & eye protection.
Trypsin
Wear gloves, mask, & eye protection.
Ethanol
Wear gloves. Work in wellventilated area.
Thymol
Wear gloves. Work in wellventilated area.
Glycerol/Glycerin Water
None None
Other notes Needs proper disposal.
Store dry and away from acids.
Hurts environment. Needs proper disposal.
Wear gloves. None
GHS explanations:
Corrosive
Harmful
Hurtful
Nausea
Flammable
Environmental Hazard
GENERAL SAFETY MEASUREMENTS: • Change gloves often: gloves are not 100% impenetrable, and chemicals will seep through them over time. • Be aware of dust/gas from harmful chemicals: Dust from powders and gas from liquids will enter your system through the skin, nose, mouth and eyes. Keep lids on chemicals as much as possible, work in well-ventilated areas (preferably in a fume hood) and wear eye protection and a filtered mask (needs to be changed regularly to work properly). • Always acid in water: When diluting acids, always put in water first, then add the acid. If adding acid to water high heat can quickly form, and acid can spray everywhere with severe burns. • Properly label and store your chemicals. Keep them away from light, moist, cold, heat, children and (living) animals. Always keep chemicals from eye level down - you do not want these chemicals to drop down on your head while reaching on a top shelf.
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Calculations No science without math! No worries - this section is meant to help guide you through that. DILUTION aka. “Help! I have solution of X%, but I need a concentration of Y%”: The answer is to make a dilution. Let’s say we have bought some formalin that is 23.4%, but we need to make a 10% dilution, and we want 150 ml of it. In this case we need to do some calculations. The old-school method is to do manual calculations using the formula: 𝑉" =
$% ∙'% $(
, where
C1 = concentration of stock, V1 = volume to take out from the stock, C2 = final concentration, V2 = final volume. So if I have a stock of 23.4% (C1) formalin, and I wish to make 150 ml (V2) of a 10% (C2) formalin solution, I will need to take out: 𝑉" =
$% ∙'% $(
=
")% ∙ ",) -. /0.2%
= 64 𝑚𝑙
from my stock solution, and add up to 150 ml with distilled water, which comes to 𝑉/ − 𝑉" = 150 𝑚𝑙 − 64 𝑚𝑙 = 86 𝑚𝑙 distilled water (see figure to the right) The easy method is to use a dilution calculator, which does exactly the same thing. A good one can be found here: http://www.physiologyweb.com/calculators/dilution_calculator_molarity_percent.html Plot in the numbers of your stock concentration (here 23.4%), the final concentration (here 10%), and the final solution volume (here 150 ml), and click calculate. The calculator will give the answer in yellow. So again, if I want to make 150 ml of 10% formalin from my 23.4% stock, I will need to measure out 64 ml from the stock and add up to 150 ml (150 − 64 = 86 𝑚𝑙) with distilled water. The tool is useful in several ways. Say you have 1L of the 23.4% formalin stock, and you wanted to dilute everything at once, you could leave the “final solution volume” field to calculate the amount of water needed. So if I wanted to dilute my entire 1L bottle, I would need to fill up to 2.34L with water (so a total of 2.34 − 1𝐿 = 1.34𝐿 water) FACTORS aka. “What does X:Y mean?”: Sometimes you will see something along the lines of 1:100 or 1:3 or even 3:1, which is another way to note dilution, or a relationship between two chemicals in a solution. So if you have the notation X:Y, there will be X amount of chemical A, and Y amount of chemical B in the solution. See calculated examples below: Example: Dilution Total parts Total ml Amount of chemical A Amount of chemical B 1:3 4 (1+3) 200 ml 200 𝑚𝑙/4 ∙ 1 = 50 𝑚𝑙 200 𝑚𝑙/4 ∙ 3 = 150 𝑚𝑙 1:1 2 (1+1) 150 ml 150 𝑚𝑙/2 = 75 𝑚𝑙 150 𝑚𝑙/2 = 75 𝑚𝑙 3:1 4 (3+1) 200 ml 200 𝑚𝑙/4 ∙ 3 = 150 𝑚𝑙 200 𝑚𝑙/4 ∙ 1 = 50 𝑚𝑙 1:100 150 ml Solid, so measure out 150 ml (where A is solid) 150 𝑚𝑙/100 = 1.5 𝑔 DISSOLVING aka. “How to go from solid to solution”: So you have some powder, and the protocol says to make a X% solution. What do you do? Well luckily 1 ml = 1 g*. So if I have some powder and I want to make 150 ml of a 1% solution of that chemical, I would need to measure out: 150 𝑚𝑙 ∗ 1% = 150 𝑚𝑙 ∗ 0.01 = 1.5 𝑚𝑙 = 1.5 𝑔 (*Now, technically 1 ml = 1 g is a golden standard that doesn’t apply to all chemicals, and to be absolutely correct you would need to calculate the mass per volume using the molar mass of the specific compound to get the right amount to add. However, when working in the lab scientist like to be lazy – very lazy, so lab protocols are most often written with that in mind – as will this)
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F.A.Q. Which animals can I use? This is a question of trial and error. So far the technique has been successful on birds, mammals, amphibians, reptiles and fish. A general rule is: The larger/more dense the specimen, the harder to clear. Small vertebrates therefore give the best results. that this protocol is optimized to small vertebrates like rodents and birds, and that some of the steps may vary a bit between species. Feather, fur and scales all needs to be removed in order to get a successful clearing. Can I use _____ instead? Again this is a question of trial and error. Diaphonization is pretty forgiving, and many of the chemicals could most likely be substituted with something else. However, it is hard to know what effect it will have on the outcome before actually trying it. For instance, I’ve had success with using 93% denatured ethanol instead of 95% pure ethanol. A natural part of lab work is to just try something to see if it works (but do it safely – you don’t want to accidently create toxic fumes or explosives). You can even experiment with other dyes if you want something other that red and blue. What is this in ounces/gallons/pound/inches? Sorry. In science we use SI-units and don’t measure with our hands and feet. You will just have to learn this wonderful new system – it might be the best thing that ever happened to you! If you live in a country that uses these Neanderthal units, please take action so following generations won’t suffer the way you have. I did everything right and it didn’t work?! Ahhh, this is the beauty of lab work. Sometimes it just doesn’t work, and you have no idea why. Maybe your chemicals are out of date (make sure that your dye solutions haven’t formed crystals), maybe your incubation times were too long/short. Maybe the animal wasn’t fresh enough. The best advice is to just try again, and pay attention to where it might be going wrong. Help! I’m stuck! Check the videos on YouTube (link on Page 1) – maybe it will make more sense then. Also feel free to write in the comments, and I’ll try my best to help.
References PRIMARY SOURCES: “Dyeing The Dead” YouTube series by TacoKel: https://www.youtube.com//TacoKellz
Taylor, W. R. (1967). An Enzyme Method of Clearing and Staining Small Vertebrates. Proceedings of the United States National Museum, 122(3596), 1–17. Weck, B., & Miljak, P. (1998). Give New Life to Old Specimens through Clearing & Staining. The American Biology Teacher, 60(9), 699–702. Cortés-Delgado, N., Pérez-Torres, J., & Hoyos, J. M. (2009). Staining Procedure of Cartilage and Skeleton in Adult Bats and Rodents. International Journal of Morphology, 27(4), 1163–1167.
A very nice Diaphonization series on YouTube, where she shows the entire process. Her videos are more talkative than mine, and she does a great job at explaining everything in an easy to understand language. Very thorough protocol. A bit long for daily use, but good if you are troubleshooting, or to get a nice introduction to diaphonization If you want a “real” protocol to read this is the one. Simple and easy to follow. The bad news is that it requires a subscription and is therefore not freely available for most people Not as nice at the Weck & Miljak one, but still easy to read. This one is much more easy to find, as it is freely available online
SECONDARY SOURCES: Green, M. C. (1952). a Rapid Method for Clearing and Staining Specimens for the Demonstration of Bone. The Ohio Journal of Science, 52(1), 31–33. Tipton, P. W., & Burtt, M. E. (1977). A method for mechanised staining of rat and mouse foetuses for teratological examination. Laboratory Animals, (11), 265–267. Dingerkus, G., & Uhler, L. D. (1977). Enzyme clearing of alcian blue stained whole small vertebrates for demonstration of cartilage. Stain Technology, 52(4), 229–232. Taylor, W., & Van Dyke, G. C. (1985). Revised procedures for staining and clearing small fishes and other vertebrates for bone and cartilage study. Cybium. Armbruster, J. W. (1989). Clearing and Staining Methods. Journal of Chemical Information and Modeling (Vol. 53).
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Protocol: Experimental overview STEP 1: Prepare, fix and wash 100 ml 10% formalin
100 ml Distilled water
100 ml Distilled water
min. 2 days
1 day
1 day
• Skin • Remove organs
STEP 2: Cartilage stain 10 mg Alcian Blue 60 ml 95% Ethanol 40 ml Glacial acetic acid 1 day
STEP 3: Rehydration 100 ml 95% ethanol
100 ml 95% ethanol
100 ml 70% ethanol
100 ml Distilled water
1 day
1 day
1 day
1 day
STEP 4: Trypsin digestion 30 ml Saturated sodium borate 70 ml Distilled water 1 g Trypsin 1-7 days: until body is limp and bones are visible. Change solution every 3 days.
STEP 5: Bone stain
100 ml 0.5% KOH 10 mg Alizarin Red 1 day
STEP 6: Clearing and bleaching + storage
75 ml 0.5% KOH 25 ml Glycerin (1 ml H2O2) 1 day - 1+ week
50 ml 0.5% KOH 50 ml Glycerin (1 ml H2O2)
1 day - 1+ week
25 ml 0.5% KOH 75 ml Glycerin 1 day - 1+ week
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Glycerin w. thymol crystals Forever…
Part 01: Preparing the specimen/skinning First the specimen needs to be skinned and it’s organs and fat removed, as these things will interfere with the clearing process. THINGS YOU’LL NEED: • Specimens: Can be fresh, frozen of previously preserved. If frozen let thaw completely before use. • Dissection tools: Scissors and forceps as a minimum • Gloves TUTORIAL: The method of skinning is very much up to personal preference. There is no perfect way of doing this. The most important thing is to not disrupt any bones in the process. This is how I do it (using a frozen feeder mouse as an example): 1) Let specimen thaw completely first 2) Make an incision down the back, and start peeling off the skin (Figure 1). Pull the skin down to the ankles and cut off, so that the skin on the feet is left on the body. For the tail pull the skin away carefully, and make small incisions along the way if there is any resistance (Figure 2) 3) Once skin is completely removed, cut open the abdomen to remove all organs. Don’t forget the heart and lungs! They can be reached by cutting through the diaphragm (Figure 3). 4) Lastly remove the fat (Figure 4). Fat can range from white to beige in color, and it is easy to spot after a quick rinse under the faucet. Remove as much as possible. 5) Once done proceed immediately to the fixation step…
Figure 1: Pulling skin
Figure 2: Removal of skin from tail
Figure 3: Removal of heart and lungs
Figure 4: Removal of fat
VIDEO TUTORIALS: Skinning of mouse: https://youtu.be/5eO1DkXtUCM Skinning of chicken: https://youtu.be/b6OfTxtizrU
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Part 02: Fixation To preserve the specimens and stop them from decaying they are fixed in formaldehyde. After this step the body will become rigid and slightly dull in color.
THINGS YOU’LL NEED: • Specimen(s): Skinned • 10% formalin*: Dependent on the concentration of your stock solution, you may need to make a dilution (see dilution calculations for help). Dilute with distilled water. • Measuring cylinder • Jar with lid • Forceps • Protection: Gloves, mask, eyewear *WARNING! Formalin is very toxic, so be sure to wear protective gear and work in a well ventilated area. TUTORIAL: After skinning and removal of fat and organs, the specimen needs to be fixed in formalin. 1) Make a 10% formalin solution and pour it into a jar with a lid. There should be enough solution to cover the specimen(s) entirely (usually 100-200 ml in a glass jar is sufficient for small animals) 2) Transfer the specimens into the formalin solution, making sure they are completely covered and close lid. 3) Leave in solution for at least 2-3 days (this time frame is fit for small animals like mice, but may vary for other species). If you need to take a break in the protocol, this is a good time to do so, as the time in formalin is not that critical. In fact the animals can be stored in formalin for years if needed (Weck & Miljak, 1998).
Figure 5: Specimens freshly placed in formalin
VIDEO TUTORIAL: https://youtu.be/zmJp7kQem10
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Part 03: Washing After fixation the specimen needs to be washed in order to rehydrate the tissues and to remove excess formaldehyde. This is done through a series of water baths over several days. THINGS YOU’LL NEED: • Specimen(s): Fixed* • Distilled water • Measuring cylinder • Jar with lid • Forceps • Protection: Gloves, mask, eyewear *WARNING! Formalin is very toxic, so be sure to wear protective gear and work in a well ventilated area. TUTORIAL: The specimen needs to go through a series of water baths. The baths are as following: Day 1: Distilled water, leave overnight Day 2: Distilled water, leave overnight 1) Transfer the specimen from the formalin to a new empty jar. 2) Pour distilled water over until it’s completely covered 3) Put lid on and leave overnight The next day: 4) Transfer the specimen to a new empty jar 5) Pour distilled water over until it’s completely covered 6) Put lid on and leave overnight The next day: 7) Move on to cartilage staining. (If needed, the specimens can be left in the water solution for a few days before moving on)
Figure 6: During water baths. The specimens are stiff and colorless after fixation.
VIDEO GUIDE: https://youtu.be/B1-VyqGwqjU
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Part 04: Cartilage staining (Alcian Blue) This is the fun part! In this step the cartilage will be stained blue using the dye Alcian Blue.
THINGS YOU’LL NEED: • Specimen(s): Fixed and washed • Ethanol 95% • Alcian Blue 8GX • Glacial Acetic Acid • Measuring cylinder • Jar with lid • Forceps • Scale + weighing paper • Spatula • Protection: Gloves, mask, eyewear TUTORIAL: For the staining we need to make the dye solution: Cartilage Dye Recipe Alcian Blue 8GX Ethanol 95% Glacial Acetic Acid Total:
Factor 0.01-0.02%* 3:2 2:3
For a 200 ml solution: 0.02-0.04 g 120 ml 80 ml 200 ml
*Dependent on specimen size and density. 0.01% should be fine for small animals like mice
1) Measure out an appropriate amount of Alcian Blue powder using a scale. The easiest way is to use weighing paper and scoop the product out with a spatula (Figure 7). For 200 ml you’ll need to measure out 20 mg (= 0.02 g) Alcian Blue (See Table). 2) Pour the Alcian Blue powder into an empty jar. 3) Add the 95% Ethanol to the jar (see Table for amount) 4) Add the glacial Acetic Acid to the jar (See Table for amount) 5) Put lid on and mix by shaking gently 6) Transfer the specimen to the jar with the dye mix, and make sure that it is completely covered. 7) Let it soak in the dye for 1 day before moving on to rehydration… → The dye solution can be used more than once. Stop using if build-up starts to appear at the bottom.
Figure 8: Specimens in dye solution
Figure 7: Measuring out Alcian Blue
VIDEO TUTORIAL: https://youtu.be/SIAypcts298
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Part 05: Rehydration Here the specimen gets rehydrated and neutralized after the acetic cartilage staining by going through a series of baths of ethanol and water. THINGS YOU’LL NEED: • Specimen(s): Stained with Alcian Blue • Ethanol 95% • Distilled water • Measuring cylinder • Jar with lid • Forceps • Protection: Gloves, mask, eyewear TUTORIAL: The specimen needs to go through a series of baths. The baths are as following: BATH I: 95% ethanol, 2 hours - 1 day BATH II: 95% ethanol, 2 hours - 1 day BATH III: 70% ethanol, 2 hours - 1 day BATH IV: Distilled water, 2 hours - 1 day Day 1, Bath I: 1) Transfer specimen to a new jar and pour over 95% ethanol until specimen is covered. Let sit overnight Day 2, Bath II: 2) Same as day 1. Let sit overnight Day 3, Bath III: 3) Make a 70% ethanol dilution: If making a total volume of 200 ml, measure out 150 ml 95% ethanol and 50 ml distilled water (See page 5 for help with calculations) 4) Transfer the specimen to the 70% ethanol and soak for 1 day Day 4, Bath IV: 5) Pour out ethanol and add distilled water. Soak for 1 day, before moving on to trypsin digestion…
Figure 9: Specimens in bath IV (water only)
VIDEO TUTORIAL: https://youtu.be/K-tIGuw2jIs
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Part 06: Trypsin digestion Trypsin is a digestive enzyme that breaks down many proteins, but leaves collagen. The breaking down of muscle and other tissues will help to make the animal transparent, while the collagen will keep the specimen from falling apart. THINGS YOU’LL NEED: • Specimen(s): Rehydrated • Sodium Borate (Borax) • Distilled water • Measuring cylinder • Small pot (to boil water) • Jar with lid • Weight + weighing paper + spatula • Forceps • Protection: Gloves, mask, eyewear TUTORIAL: 1) Since Trypsin works best at a slightly elevated pH (~7.5-8.5), an alkaline buffer of sodium borate (borax) is made: a) Boil some distilled water (you’ll need 60 ml for a 200 ml solution) b) Transfer the hot water to a heatsafe container and add borax until saturated. It is saturated when small insoluble crystals form at the bottom. c) Trypsin is temperature sensitive, so cool to room temperature before moving on. 2) Once the borax buffer has cooled completely, you’ll need to make the digestion mixture: Digestion mixture Borax buffer solution Distilled water Trypsin Total
Factor 30% 70% 1:100
For a 200 ml solution: 60 ml 140 ml 2g 200 ml
3) 4) 5) 6)
Measure out the borax buffer, water and trypsin (see table for volumens) into an empty jar. Put on lid and shake gently to mix. Transfer specimens to the digestion mixture The incubation time will depend on size and build of the specimen. Check on the specimens daily to look for signs of proper digestion. You want the specimen to be limp and bones to be slightly visible (Figure 10 + Figure 11) before moving on to the next step. 7) Continue immediately to bone staining once ready… → Usually by 2-3 days with small animals like mice. Change the solution after 3 days if longer incubation is needed.
Figure 10: Bone visibility in a mouse after 3 days of digestion. Ribcage is clearly visible.
Figure 11: Bone visibility in a mouse after 3 days of digestion. Bones in hind legs are slightly visible (yellow arrow)
VIDEO TUTORIAL: https://youtu.be/Y_5HTxB0kXM
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Part 07: Bone staining (Alizarin Red) This is the second dye step. Here the bones will be stained that characteristic purply red color using a dye called Alizarin Red.
THINGS YOU’LL NEED: • Specimen(s): Digested • Potassium hydroxide (KOH) • Alizarin Red S • Distilled water • Measuring cylinder • Jar with lid • Weight + weighing paper + spatula • Forceps • Protection: Gloves, mask, eyewear TUTORIAL: Dye recipe KOH Alizarin Red S Distilled water Total 1) 2) 3) 4) 5)
Factor: 0.5-2%* 1:10,000
For a 250 ml solution: 1.25-5 g 0.025 g 250 ml 250 ml
*Dependent on specimen size and density. 0.5% is fine for small animals like mice
Measure out the KOH needed (see Table) and add to a new jar Add distilled water Measure out the Alizarin Red S (you only need a tiny amount) and add to the jar (Figure 12) Put on lid and shake to mix Transfer the specimens to the dye solution and stain for 1 day before moving on to clearing… → The dye solution can be reused as long as there is no precipitation.
Figure 12: Ooh, pretty
Figure 13: Specimens in Alizarin dye solution
VIDEO TUTORIAL: https://youtu.be/YhxgNgd1OMQ
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Part 08: Clearing This is the longest part of the protocol, but also the part where we see the biggest changes. The specimen will go through a series of baths to make it transparent. The longer the time spent in these baths, the better the clearing – however be careful not to leave specimens for too long as this might make them come apart. THINGS YOU’LL NEED: • Specimen(s): Dyed with Alizarin Red • Potassium hydroxide (KOH) • Distilled water • Glycerin • 3% Hydrogen Peroxide (H2O2): Optional. Will bleach brown areas. Often not needed for smaller animals
• • • • •
Measuring cylinder Jar with lid Weight + weighing paper + spatula Forceps Protection: Gloves, mask, eyewear
TUTORIAL: The specimen needs to go through a series of clearing baths. The baths are as following: BATH I: 3:1 0.5% KOH to Glycerin (Optional: 1:100 3% H2O2), leave 1 day to over a week BATH II: 1:1 0.5% KOH to Glycerin (Optional: 1:100 3% H2O2), leave 1 day to over a week BATH III: 1:3 0.5% KOH to Glycerin, leave 1 day to over a week 1) Start by making a 0.5% KOH stock solution. Make enough for all 3 baths: 0.5-2% KOH stock solution KOH Distilled water Total
Factor 0.5-2%**
For a 300 ml solution: 1.25-5 g 250 ml 250 ml
**Dependent on specimen size and density, 0.5% is fine for small animals like mice, but 2% is better for larger and/or more dense animals
2) Make the 3 baths in separate jars, according to following table:
KOH solution (0.5-2%): Glycerin: H2O2 (optional): Total:
BATH I Factor: 3:1 1:3 1:100
200 ml: 150 ml 50 ml 2 ml 202 ml
BATH 2 Factor: 1:1 1:1 1:100
150 ml: 75 ml 75 ml 1.5 ml 151.5 ml
BATH 3 Factor: 1:3 3:1
200 ml: 50 ml 150 ml 200 ml
3) Transfer the specimens to bath I and incubate for 1 day to over a week 4) Transfer the specimens to bath II and incubate for 1 day to over a week 5) Transfer the specimens to bath III and incubate for 1 day to over a week
Figure 14: Specimens in Bath III of the clearing process
VIDEO TUTORIAL: https://youtu.be/EVQ28494ymc
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Part 09: Results Now it’s time to finish up the specimens. The specimens should at this point be quite transparent with visible staining. The last bit of transparency we’ll get by the use of magic! Or well, basic science. how trypsin digested everything but collagen? Collagen and glycerin has the same refractive index, meaning they bend light in the same manner. When light can freely through two objects without bending they will appear transparent. That means that placing the collagen-rich specimen into 100% glycerin will give the illusion that it´s even more transparent. Neat, right?
THINGS YOU’LL NEED: • Specimen(s): Cleared and stained • Glycerin • Thymol Crystals • Jars for storage • Dissection tools: Scissors and forceps as a minimum • Protection: Gloves, mask, eyewear TUTORIAL: 1) Take the specimen out of the jar for inspection 2) Cut off any loose tissue or small imperfections that doesn’t look good (Figure 15) 3) Once satisfied put the specimen into a jar that fits the specimen nicely 4) Fill the jar with glycerin 5) Add a pinch of thymol crystals – this will work as an anti-fungal agent 6) Use forceps to play around with the placement of the specimen in the jar, add more glycerin if needed, and put the lid on once satisfied. Sometimes the specimen can be a little floaty and hard to work with. This is due to tiny air bubbles stuck inside cavities - don’t worry, they will disappear over time. Just leave the specimen for a few days/weeks to allow the bubbles to clear, and then try to reposition it again. 7) You are done! Go enjoy your morbidly cool looking creatures!
Figure 15: Removing excess tissue
Figure 16: The final product! If you were not careful during the skinning process it will show (hence the broken leg)
VIDEO TUTORIAL: https://youtu.be/fH_5GvM8UdI
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I’m on YouTube! To watch this protocol in live action, click here: https://www.youtube.com/playlist?list=PLE0GkpL0JFYM3eNOuD7CCmcFlb0LQ4g Or scan the following QR code:
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Preface Diaphonization (also known as clearing and staining) is a preservation technique in which the tissues of a specimen is made transparent, and the cartilage and bones are made visible by use of specific dyes. Diaphonization is nothing new – in fact it has been around since before the 1940’ies, where the process enabled zoologists to study the bones while still inside the animal. The flexibility of the specimens even allowed them to study these bones in movement, and it was therefore a powerful tool in order to better understand bone function. While this technique is still used in zoology, there has been a recent spark of interest in diaphonization outside the zoology field due to their visual appeal. Diaphonization is now also part of the taxidermy community, and diaphonized specimens are being sold as artsy display items worldwide. Part of this comes down to the simplicity of the technique; diaphonization is a pretty easy and forgiving process that is possible to do with relatively few remedies and without advanced lab equipment. This means that – in theory – this could be done in the comfort of your own home. While there are a lot of protocols out there, they can be hard to find and navigate without a scientific background. The idea of this protocol is therefore to help bring diaphonization to the layman. About me: I have a background in science (biochemistry) and spend a lot of my time in a lab. I started diaphonization as something fun to do at home, and thought I would share it with the world. While I do have experience in general lab-related tissue/cell dyeing, diaphonization is still new to me, and I am in no way a professional. If you find something gravely wrong, or anything else that could be improved, please don’t hesitate to me J
Table of contents Introduction Materials A ‘quick’ note on safety Calculations F.A.Q. References
3 3 4 5 6 6
Protocol: Experimental overview Part 01: Preparing the specimen/skinning Part 02: Fixation Part 03: Washing Part 04: Cartilage staining (Alcian Blue) Part 05: Rehydration Part 06: Trypsin digestion Part 07: Bone staining (Alizarin Red) Part 08: Clearing Part 09: Results
7 8 9 10 11 12 13 14 15 16
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Introduction This is the boring part, but it needs to be here. It has a list of the materials needed, but it also includes some words about safety when working with these types of chemicals, and a section on how to do some typical lab work calculations that you might need during this protocol.
Materials LIST OF CHEMICALS NEEDED: Name
Formula
State of matter
Distilled/deionized Water
H 2O
Liquid
10% Formalin
CH2O • H2O
Liquid
99% Glacial Acetic Acid
C 2H 4O 2
Liquid
95% Ethanol/Ethyl Alcohol
C 2H 6O
Liquid
Potassium Hydroxide
KOH
Flakes
Sodium Borate (Borax)
Na2B4O7
Powder
3% Hydrogen Peroxide
H 2O 2
Liquid
Thymol
C10H14O
Crystals
Glycerin/Glycerol
C 3H 8O 3
Liquid
Trypsin 1:100, 25g
Powder
Alizarin Red S
Powder
Alcian Blue 8GX
Powder
Notes
93% denatured ethanol will work just fine too
Optional
LIST OF MATERIALS NEEDED: Item
Notes This protocol is mostly suited for small vertebrates like mice – some animals may require tweaking of the protocol in order to work. Frozen feeder animals are good to start with.
Specimens Surgical kit for dissecting
Get at least 1 decent pair of scissors + forceps
Graduated cylinders
A set of e.g. 100 mL, 20 mL and 5 mL will get you a long way
Scale + weighing paper + spatula
You want a scale that is able to weigh down to 0.01g
Jars/containers
These are used throughout the protocol, so you’ll want a good amount of these in various sizes. Old food jars are fine as long as they are cleaned well
Safety
You want gloves, a mask and some googles
Waste containers
Get some containers for your waste liquids, so that you can dispose of this properly (check your local waste rules)
WHERE TO GET EVERYTHING: Sorry, can’t help you there. This is the hardest part, and depending on your country of residence this can be a real struggle. I’ve had some good luck on Amazon and eBay/AliExpress though.
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A ‘quick’ note on safety Some of these chemicals are quite nasty – both for the body and the environment. It is always a good idea to know your chemicals before you start working with them. Detailed safety sheets should come with the chemicals when you buy them, but here is a quick summary: Chemical Formaldehyde/formalin
GHS symbols
Safety measurements Wear gloves, mask, & eye protection. Work in wellventilated area. Can cause cancer and genetic defects. Do not breathe in! Wear gloves, mask, & eye protection. Work in wellventilated area
Potassium hydroxide (KOH) Borax
Wear gloves. Work in wellventilated area/with mask.
Glacial Acetic Acid
Wear gloves, mask, & eye protection. Work in wellventilated area.
Alizarin Red S
Wear gloves. Work in wellventilated area.
Alcian Blue 8GX
Wear gloves, mask, & eye protection.
Trypsin
Wear gloves, mask, & eye protection.
Ethanol
Wear gloves. Work in wellventilated area.
Thymol
Wear gloves. Work in wellventilated area.
Glycerol/Glycerin Water
None None
Other notes Needs proper disposal.
Store dry and away from acids.
Hurts environment. Needs proper disposal.
Wear gloves. None
GHS explanations:
Corrosive
Harmful
Hurtful
Nausea
Flammable
Environmental Hazard
GENERAL SAFETY MEASUREMENTS: • Change gloves often: gloves are not 100% impenetrable, and chemicals will seep through them over time. • Be aware of dust/gas from harmful chemicals: Dust from powders and gas from liquids will enter your system through the skin, nose, mouth and eyes. Keep lids on chemicals as much as possible, work in well-ventilated areas (preferably in a fume hood) and wear eye protection and a filtered mask (needs to be changed regularly to work properly). • Always acid in water: When diluting acids, always put in water first, then add the acid. If adding acid to water high heat can quickly form, and acid can spray everywhere with severe burns. • Properly label and store your chemicals. Keep them away from light, moist, cold, heat, children and (living) animals. Always keep chemicals from eye level down - you do not want these chemicals to drop down on your head while reaching on a top shelf.
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Calculations No science without math! No worries - this section is meant to help guide you through that. DILUTION aka. “Help! I have solution of X%, but I need a concentration of Y%”: The answer is to make a dilution. Let’s say we have bought some formalin that is 23.4%, but we need to make a 10% dilution, and we want 150 ml of it. In this case we need to do some calculations. The old-school method is to do manual calculations using the formula: 𝑉" =
$% ∙'% $(
, where
C1 = concentration of stock, V1 = volume to take out from the stock, C2 = final concentration, V2 = final volume. So if I have a stock of 23.4% (C1) formalin, and I wish to make 150 ml (V2) of a 10% (C2) formalin solution, I will need to take out: 𝑉" =
$% ∙'% $(
=
")% ∙ ",) -. /0.2%
= 64 𝑚𝑙
from my stock solution, and add up to 150 ml with distilled water, which comes to 𝑉/ − 𝑉" = 150 𝑚𝑙 − 64 𝑚𝑙 = 86 𝑚𝑙 distilled water (see figure to the right) The easy method is to use a dilution calculator, which does exactly the same thing. A good one can be found here: http://www.physiologyweb.com/calculators/dilution_calculator_molarity_percent.html Plot in the numbers of your stock concentration (here 23.4%), the final concentration (here 10%), and the final solution volume (here 150 ml), and click calculate. The calculator will give the answer in yellow. So again, if I want to make 150 ml of 10% formalin from my 23.4% stock, I will need to measure out 64 ml from the stock and add up to 150 ml (150 − 64 = 86 𝑚𝑙) with distilled water. The tool is useful in several ways. Say you have 1L of the 23.4% formalin stock, and you wanted to dilute everything at once, you could leave the “final solution volume” field to calculate the amount of water needed. So if I wanted to dilute my entire 1L bottle, I would need to fill up to 2.34L with water (so a total of 2.34 − 1𝐿 = 1.34𝐿 water) FACTORS aka. “What does X:Y mean?”: Sometimes you will see something along the lines of 1:100 or 1:3 or even 3:1, which is another way to note dilution, or a relationship between two chemicals in a solution. So if you have the notation X:Y, there will be X amount of chemical A, and Y amount of chemical B in the solution. See calculated examples below: Example: Dilution Total parts Total ml Amount of chemical A Amount of chemical B 1:3 4 (1+3) 200 ml 200 𝑚𝑙/4 ∙ 1 = 50 𝑚𝑙 200 𝑚𝑙/4 ∙ 3 = 150 𝑚𝑙 1:1 2 (1+1) 150 ml 150 𝑚𝑙/2 = 75 𝑚𝑙 150 𝑚𝑙/2 = 75 𝑚𝑙 3:1 4 (3+1) 200 ml 200 𝑚𝑙/4 ∙ 3 = 150 𝑚𝑙 200 𝑚𝑙/4 ∙ 1 = 50 𝑚𝑙 1:100 150 ml Solid, so measure out 150 ml (where A is solid) 150 𝑚𝑙/100 = 1.5 𝑔 DISSOLVING aka. “How to go from solid to solution”: So you have some powder, and the protocol says to make a X% solution. What do you do? Well luckily 1 ml = 1 g*. So if I have some powder and I want to make 150 ml of a 1% solution of that chemical, I would need to measure out: 150 𝑚𝑙 ∗ 1% = 150 𝑚𝑙 ∗ 0.01 = 1.5 𝑚𝑙 = 1.5 𝑔 (*Now, technically 1 ml = 1 g is a golden standard that doesn’t apply to all chemicals, and to be absolutely correct you would need to calculate the mass per volume using the molar mass of the specific compound to get the right amount to add. However, when working in the lab scientist like to be lazy – very lazy, so lab protocols are most often written with that in mind – as will this)
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F.A.Q. Which animals can I use? This is a question of trial and error. So far the technique has been successful on birds, mammals, amphibians, reptiles and fish. A general rule is: The larger/more dense the specimen, the harder to clear. Small vertebrates therefore give the best results. that this protocol is optimized to small vertebrates like rodents and birds, and that some of the steps may vary a bit between species. Feather, fur and scales all needs to be removed in order to get a successful clearing. Can I use _____ instead? Again this is a question of trial and error. Diaphonization is pretty forgiving, and many of the chemicals could most likely be substituted with something else. However, it is hard to know what effect it will have on the outcome before actually trying it. For instance, I’ve had success with using 93% denatured ethanol instead of 95% pure ethanol. A natural part of lab work is to just try something to see if it works (but do it safely – you don’t want to accidently create toxic fumes or explosives). You can even experiment with other dyes if you want something other that red and blue. What is this in ounces/gallons/pound/inches? Sorry. In science we use SI-units and don’t measure with our hands and feet. You will just have to learn this wonderful new system – it might be the best thing that ever happened to you! If you live in a country that uses these Neanderthal units, please take action so following generations won’t suffer the way you have. I did everything right and it didn’t work?! Ahhh, this is the beauty of lab work. Sometimes it just doesn’t work, and you have no idea why. Maybe your chemicals are out of date (make sure that your dye solutions haven’t formed crystals), maybe your incubation times were too long/short. Maybe the animal wasn’t fresh enough. The best advice is to just try again, and pay attention to where it might be going wrong. Help! I’m stuck! Check the videos on YouTube (link on Page 1) – maybe it will make more sense then. Also feel free to write in the comments, and I’ll try my best to help.
References PRIMARY SOURCES: “Dyeing The Dead” YouTube series by TacoKel: https://www.youtube.com//TacoKellz
Taylor, W. R. (1967). An Enzyme Method of Clearing and Staining Small Vertebrates. Proceedings of the United States National Museum, 122(3596), 1–17. Weck, B., & Miljak, P. (1998). Give New Life to Old Specimens through Clearing & Staining. The American Biology Teacher, 60(9), 699–702. Cortés-Delgado, N., Pérez-Torres, J., & Hoyos, J. M. (2009). Staining Procedure of Cartilage and Skeleton in Adult Bats and Rodents. International Journal of Morphology, 27(4), 1163–1167.
A very nice Diaphonization series on YouTube, where she shows the entire process. Her videos are more talkative than mine, and she does a great job at explaining everything in an easy to understand language. Very thorough protocol. A bit long for daily use, but good if you are troubleshooting, or to get a nice introduction to diaphonization If you want a “real” protocol to read this is the one. Simple and easy to follow. The bad news is that it requires a subscription and is therefore not freely available for most people Not as nice at the Weck & Miljak one, but still easy to read. This one is much more easy to find, as it is freely available online
SECONDARY SOURCES: Green, M. C. (1952). a Rapid Method for Clearing and Staining Specimens for the Demonstration of Bone. The Ohio Journal of Science, 52(1), 31–33. Tipton, P. W., & Burtt, M. E. (1977). A method for mechanised staining of rat and mouse foetuses for teratological examination. Laboratory Animals, (11), 265–267. Dingerkus, G., & Uhler, L. D. (1977). Enzyme clearing of alcian blue stained whole small vertebrates for demonstration of cartilage. Stain Technology, 52(4), 229–232. Taylor, W., & Van Dyke, G. C. (1985). Revised procedures for staining and clearing small fishes and other vertebrates for bone and cartilage study. Cybium. Armbruster, J. W. (1989). Clearing and Staining Methods. Journal of Chemical Information and Modeling (Vol. 53).
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Protocol: Experimental overview STEP 1: Prepare, fix and wash 100 ml 10% formalin
100 ml Distilled water
100 ml Distilled water
min. 2 days
1 day
1 day
• Skin • Remove organs
STEP 2: Cartilage stain 10 mg Alcian Blue 60 ml 95% Ethanol 40 ml Glacial acetic acid 1 day
STEP 3: Rehydration 100 ml 95% ethanol
100 ml 95% ethanol
100 ml 70% ethanol
100 ml Distilled water
1 day
1 day
1 day
1 day
STEP 4: Trypsin digestion 30 ml Saturated sodium borate 70 ml Distilled water 1 g Trypsin 1-7 days: until body is limp and bones are visible. Change solution every 3 days.
STEP 5: Bone stain
100 ml 0.5% KOH 10 mg Alizarin Red 1 day
STEP 6: Clearing and bleaching + storage
75 ml 0.5% KOH 25 ml Glycerin (1 ml H2O2) 1 day - 1+ week
50 ml 0.5% KOH 50 ml Glycerin (1 ml H2O2)
1 day - 1+ week
25 ml 0.5% KOH 75 ml Glycerin 1 day - 1+ week
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Glycerin w. thymol crystals Forever…
Part 01: Preparing the specimen/skinning First the specimen needs to be skinned and it’s organs and fat removed, as these things will interfere with the clearing process. THINGS YOU’LL NEED: • Specimens: Can be fresh, frozen of previously preserved. If frozen let thaw completely before use. • Dissection tools: Scissors and forceps as a minimum • Gloves TUTORIAL: The method of skinning is very much up to personal preference. There is no perfect way of doing this. The most important thing is to not disrupt any bones in the process. This is how I do it (using a frozen feeder mouse as an example): 1) Let specimen thaw completely first 2) Make an incision down the back, and start peeling off the skin (Figure 1). Pull the skin down to the ankles and cut off, so that the skin on the feet is left on the body. For the tail pull the skin away carefully, and make small incisions along the way if there is any resistance (Figure 2) 3) Once skin is completely removed, cut open the abdomen to remove all organs. Don’t forget the heart and lungs! They can be reached by cutting through the diaphragm (Figure 3). 4) Lastly remove the fat (Figure 4). Fat can range from white to beige in color, and it is easy to spot after a quick rinse under the faucet. Remove as much as possible. 5) Once done proceed immediately to the fixation step…
Figure 1: Pulling skin
Figure 2: Removal of skin from tail
Figure 3: Removal of heart and lungs
Figure 4: Removal of fat
VIDEO TUTORIALS: Skinning of mouse: https://youtu.be/5eO1DkXtUCM Skinning of chicken: https://youtu.be/b6OfTxtizrU
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Part 02: Fixation To preserve the specimens and stop them from decaying they are fixed in formaldehyde. After this step the body will become rigid and slightly dull in color.
THINGS YOU’LL NEED: • Specimen(s): Skinned • 10% formalin*: Dependent on the concentration of your stock solution, you may need to make a dilution (see dilution calculations for help). Dilute with distilled water. • Measuring cylinder • Jar with lid • Forceps • Protection: Gloves, mask, eyewear *WARNING! Formalin is very toxic, so be sure to wear protective gear and work in a well ventilated area. TUTORIAL: After skinning and removal of fat and organs, the specimen needs to be fixed in formalin. 1) Make a 10% formalin solution and pour it into a jar with a lid. There should be enough solution to cover the specimen(s) entirely (usually 100-200 ml in a glass jar is sufficient for small animals) 2) Transfer the specimens into the formalin solution, making sure they are completely covered and close lid. 3) Leave in solution for at least 2-3 days (this time frame is fit for small animals like mice, but may vary for other species). If you need to take a break in the protocol, this is a good time to do so, as the time in formalin is not that critical. In fact the animals can be stored in formalin for years if needed (Weck & Miljak, 1998).
Figure 5: Specimens freshly placed in formalin
VIDEO TUTORIAL: https://youtu.be/zmJp7kQem10
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Part 03: Washing After fixation the specimen needs to be washed in order to rehydrate the tissues and to remove excess formaldehyde. This is done through a series of water baths over several days. THINGS YOU’LL NEED: • Specimen(s): Fixed* • Distilled water • Measuring cylinder • Jar with lid • Forceps • Protection: Gloves, mask, eyewear *WARNING! Formalin is very toxic, so be sure to wear protective gear and work in a well ventilated area. TUTORIAL: The specimen needs to go through a series of water baths. The baths are as following: Day 1: Distilled water, leave overnight Day 2: Distilled water, leave overnight 1) Transfer the specimen from the formalin to a new empty jar. 2) Pour distilled water over until it’s completely covered 3) Put lid on and leave overnight The next day: 4) Transfer the specimen to a new empty jar 5) Pour distilled water over until it’s completely covered 6) Put lid on and leave overnight The next day: 7) Move on to cartilage staining. (If needed, the specimens can be left in the water solution for a few days before moving on)
Figure 6: During water baths. The specimens are stiff and colorless after fixation.
VIDEO GUIDE: https://youtu.be/B1-VyqGwqjU
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Part 04: Cartilage staining (Alcian Blue) This is the fun part! In this step the cartilage will be stained blue using the dye Alcian Blue.
THINGS YOU’LL NEED: • Specimen(s): Fixed and washed • Ethanol 95% • Alcian Blue 8GX • Glacial Acetic Acid • Measuring cylinder • Jar with lid • Forceps • Scale + weighing paper • Spatula • Protection: Gloves, mask, eyewear TUTORIAL: For the staining we need to make the dye solution: Cartilage Dye Recipe Alcian Blue 8GX Ethanol 95% Glacial Acetic Acid Total:
Factor 0.01-0.02%* 3:2 2:3
For a 200 ml solution: 0.02-0.04 g 120 ml 80 ml 200 ml
*Dependent on specimen size and density. 0.01% should be fine for small animals like mice
1) Measure out an appropriate amount of Alcian Blue powder using a scale. The easiest way is to use weighing paper and scoop the product out with a spatula (Figure 7). For 200 ml you’ll need to measure out 20 mg (= 0.02 g) Alcian Blue (See Table). 2) Pour the Alcian Blue powder into an empty jar. 3) Add the 95% Ethanol to the jar (see Table for amount) 4) Add the glacial Acetic Acid to the jar (See Table for amount) 5) Put lid on and mix by shaking gently 6) Transfer the specimen to the jar with the dye mix, and make sure that it is completely covered. 7) Let it soak in the dye for 1 day before moving on to rehydration… → The dye solution can be used more than once. Stop using if build-up starts to appear at the bottom.
Figure 8: Specimens in dye solution
Figure 7: Measuring out Alcian Blue
VIDEO TUTORIAL: https://youtu.be/SIAypcts298
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Part 05: Rehydration Here the specimen gets rehydrated and neutralized after the acetic cartilage staining by going through a series of baths of ethanol and water. THINGS YOU’LL NEED: • Specimen(s): Stained with Alcian Blue • Ethanol 95% • Distilled water • Measuring cylinder • Jar with lid • Forceps • Protection: Gloves, mask, eyewear TUTORIAL: The specimen needs to go through a series of baths. The baths are as following: BATH I: 95% ethanol, 2 hours - 1 day BATH II: 95% ethanol, 2 hours - 1 day BATH III: 70% ethanol, 2 hours - 1 day BATH IV: Distilled water, 2 hours - 1 day Day 1, Bath I: 1) Transfer specimen to a new jar and pour over 95% ethanol until specimen is covered. Let sit overnight Day 2, Bath II: 2) Same as day 1. Let sit overnight Day 3, Bath III: 3) Make a 70% ethanol dilution: If making a total volume of 200 ml, measure out 150 ml 95% ethanol and 50 ml distilled water (See page 5 for help with calculations) 4) Transfer the specimen to the 70% ethanol and soak for 1 day Day 4, Bath IV: 5) Pour out ethanol and add distilled water. Soak for 1 day, before moving on to trypsin digestion…
Figure 9: Specimens in bath IV (water only)
VIDEO TUTORIAL: https://youtu.be/K-tIGuw2jIs
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Part 06: Trypsin digestion Trypsin is a digestive enzyme that breaks down many proteins, but leaves collagen. The breaking down of muscle and other tissues will help to make the animal transparent, while the collagen will keep the specimen from falling apart. THINGS YOU’LL NEED: • Specimen(s): Rehydrated • Sodium Borate (Borax) • Distilled water • Measuring cylinder • Small pot (to boil water) • Jar with lid • Weight + weighing paper + spatula • Forceps • Protection: Gloves, mask, eyewear TUTORIAL: 1) Since Trypsin works best at a slightly elevated pH (~7.5-8.5), an alkaline buffer of sodium borate (borax) is made: a) Boil some distilled water (you’ll need 60 ml for a 200 ml solution) b) Transfer the hot water to a heatsafe container and add borax until saturated. It is saturated when small insoluble crystals form at the bottom. c) Trypsin is temperature sensitive, so cool to room temperature before moving on. 2) Once the borax buffer has cooled completely, you’ll need to make the digestion mixture: Digestion mixture Borax buffer solution Distilled water Trypsin Total
Factor 30% 70% 1:100
For a 200 ml solution: 60 ml 140 ml 2g 200 ml
3) 4) 5) 6)
Measure out the borax buffer, water and trypsin (see table for volumens) into an empty jar. Put on lid and shake gently to mix. Transfer specimens to the digestion mixture The incubation time will depend on size and build of the specimen. Check on the specimens daily to look for signs of proper digestion. You want the specimen to be limp and bones to be slightly visible (Figure 10 + Figure 11) before moving on to the next step. 7) Continue immediately to bone staining once ready… → Usually by 2-3 days with small animals like mice. Change the solution after 3 days if longer incubation is needed.
Figure 10: Bone visibility in a mouse after 3 days of digestion. Ribcage is clearly visible.
Figure 11: Bone visibility in a mouse after 3 days of digestion. Bones in hind legs are slightly visible (yellow arrow)
VIDEO TUTORIAL: https://youtu.be/Y_5HTxB0kXM
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Part 07: Bone staining (Alizarin Red) This is the second dye step. Here the bones will be stained that characteristic purply red color using a dye called Alizarin Red.
THINGS YOU’LL NEED: • Specimen(s): Digested • Potassium hydroxide (KOH) • Alizarin Red S • Distilled water • Measuring cylinder • Jar with lid • Weight + weighing paper + spatula • Forceps • Protection: Gloves, mask, eyewear TUTORIAL: Dye recipe KOH Alizarin Red S Distilled water Total 1) 2) 3) 4) 5)
Factor: 0.5-2%* 1:10,000
For a 250 ml solution: 1.25-5 g 0.025 g 250 ml 250 ml
*Dependent on specimen size and density. 0.5% is fine for small animals like mice
Measure out the KOH needed (see Table) and add to a new jar Add distilled water Measure out the Alizarin Red S (you only need a tiny amount) and add to the jar (Figure 12) Put on lid and shake to mix Transfer the specimens to the dye solution and stain for 1 day before moving on to clearing… → The dye solution can be reused as long as there is no precipitation.
Figure 12: Ooh, pretty
Figure 13: Specimens in Alizarin dye solution
VIDEO TUTORIAL: https://youtu.be/YhxgNgd1OMQ
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Part 08: Clearing This is the longest part of the protocol, but also the part where we see the biggest changes. The specimen will go through a series of baths to make it transparent. The longer the time spent in these baths, the better the clearing – however be careful not to leave specimens for too long as this might make them come apart. THINGS YOU’LL NEED: • Specimen(s): Dyed with Alizarin Red • Potassium hydroxide (KOH) • Distilled water • Glycerin • 3% Hydrogen Peroxide (H2O2): Optional. Will bleach brown areas. Often not needed for smaller animals
• • • • •
Measuring cylinder Jar with lid Weight + weighing paper + spatula Forceps Protection: Gloves, mask, eyewear
TUTORIAL: The specimen needs to go through a series of clearing baths. The baths are as following: BATH I: 3:1 0.5% KOH to Glycerin (Optional: 1:100 3% H2O2), leave 1 day to over a week BATH II: 1:1 0.5% KOH to Glycerin (Optional: 1:100 3% H2O2), leave 1 day to over a week BATH III: 1:3 0.5% KOH to Glycerin, leave 1 day to over a week 1) Start by making a 0.5% KOH stock solution. Make enough for all 3 baths: 0.5-2% KOH stock solution KOH Distilled water Total
Factor 0.5-2%**
For a 300 ml solution: 1.25-5 g 250 ml 250 ml
**Dependent on specimen size and density, 0.5% is fine for small animals like mice, but 2% is better for larger and/or more dense animals
2) Make the 3 baths in separate jars, according to following table:
KOH solution (0.5-2%): Glycerin: H2O2 (optional): Total:
BATH I Factor: 3:1 1:3 1:100
200 ml: 150 ml 50 ml 2 ml 202 ml
BATH 2 Factor: 1:1 1:1 1:100
150 ml: 75 ml 75 ml 1.5 ml 151.5 ml
BATH 3 Factor: 1:3 3:1
200 ml: 50 ml 150 ml 200 ml
3) Transfer the specimens to bath I and incubate for 1 day to over a week 4) Transfer the specimens to bath II and incubate for 1 day to over a week 5) Transfer the specimens to bath III and incubate for 1 day to over a week
Figure 14: Specimens in Bath III of the clearing process
VIDEO TUTORIAL: https://youtu.be/EVQ28494ymc
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Part 09: Results Now it’s time to finish up the specimens. The specimens should at this point be quite transparent with visible staining. The last bit of transparency we’ll get by the use of magic! Or well, basic science. how trypsin digested everything but collagen? Collagen and glycerin has the same refractive index, meaning they bend light in the same manner. When light can freely through two objects without bending they will appear transparent. That means that placing the collagen-rich specimen into 100% glycerin will give the illusion that it´s even more transparent. Neat, right?
THINGS YOU’LL NEED: • Specimen(s): Cleared and stained • Glycerin • Thymol Crystals • Jars for storage • Dissection tools: Scissors and forceps as a minimum • Protection: Gloves, mask, eyewear TUTORIAL: 1) Take the specimen out of the jar for inspection 2) Cut off any loose tissue or small imperfections that doesn’t look good (Figure 15) 3) Once satisfied put the specimen into a jar that fits the specimen nicely 4) Fill the jar with glycerin 5) Add a pinch of thymol crystals – this will work as an anti-fungal agent 6) Use forceps to play around with the placement of the specimen in the jar, add more glycerin if needed, and put the lid on once satisfied. Sometimes the specimen can be a little floaty and hard to work with. This is due to tiny air bubbles stuck inside cavities - don’t worry, they will disappear over time. Just leave the specimen for a few days/weeks to allow the bubbles to clear, and then try to reposition it again. 7) You are done! Go enjoy your morbidly cool looking creatures!
Figure 15: Removing excess tissue
Figure 16: The final product! If you were not careful during the skinning process it will show (hence the broken leg)
VIDEO TUTORIAL: https://youtu.be/fH_5GvM8UdI
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