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Clayton Smith, John Mendenhall, and Masa Kuwajima (K. Harris lab, Ctr for Learning and Memory, UT-Austin.)

0.  Safety Precautions:

• You must complete required lab safety training before starting this procedure.
• If this is your first time doing this procedure, ask to be trained by an experienced lab member.  If you have not done this in a while, you should ask for a refresher.
• Before starting, even if you have done this procedure before,
  • read this protocol entirely
  • ensure you have all reagents and supplies listed below
  • ensure all equipment is in good working order
  • have all waste containers ready (also see Clean-up)
  • plan your schedule well so that you wouldn’t have to rush
  • review SDS and Harris Lab SOP for the following hazardous chemicals used in this procedure:
    • Ethanol: flammable; irritant (eye)
    • Lead nitrate: oxidizer; toxic (ingestion, inhalation); carcinogen; reproductive toxin; causes serious eye damage; environmental toxin
    • LX-112 epoxy resin kit
      • LX-112: irritation (eye, skin, respiratory, oral)
      • DDSA: irritant (eye, skin, respiratory)
      • NMA: irritant (eye, skin, respiratory)
      • DMP-30: permanent eye damage, irritant (skin, respiratory, oral)
    • Osmium tetroxide: acutely toxic; mutagenic
    • Potassium ferrocyanide: contact with acid releases very toxic gas
    • Potassium hydroxide: corrosive; causes severe skin burns and eye damage; irritant (respiratory)
    • Propylene oxide: flammable; acutely toxic (skin, respiratory, oral); serious eye damage; mutagen; carcinogen
    • Sodium cacodylate: carcinogen; irritant (skin, eye); skin permeator
    • Uranyl acetate: fatal (inhalation, ingestion); flammable
• The following Personal Protective Equipment is required for this procedure:
  • Lab coat
  • Nitrile gloves (double-layer required; regularly check for holes)
  • Eye goggles
  • Also recommended when using osmium tetroxide and uranyl acetate: plastic apron and shoulder-length gloves
  • Place a piece of absorbent sheet on the work surface before starting the procedure.  When done, discard into the “Solid Waste – UA” bag

1.  Reagents and supplies

• Worksheet for this procedure [En bloc processing]
• Fixed brain tissue (100 µm thickness), embedded into 7-9% agarose
• H₂O (purified water: double-distilled, ASTM type I, WFI grade, or equivalent; e.g., Fisher 9150-25 or VWR RC915025)
• 0.2M sodium cacodylate buffer (SCB; EMS 11652)
• 0.1M SCB (diluted from 0.2M SCB)
• K₄Fe(CN)₆ · 3H₂O (potassium ferrocyanide, or KFeCN; Sigma P3289-100G)
• 4% aqueous OsO₄ (osmium tetroxide; stored at 4°C; EMS 19190): See 2.4.2
  • Use a fresh ampule for each processing; discard excess
• 1% aqueous uranyl acetate (UA; EMS 22400): See 2.4.1
  • Dissolve 0.1 g UA in 10 ml purified water, using a sonicator bath.
• Lead nitrate (EMS or Sigma-Aldrich)
• Aspertic acid (Sigma-Aldrich A9256)
• Potassium hydroxide (EMS)
• Ethanol (EtOH; EMS 15056); use a fresh bottle for each processing.
• Propylene oxide (PO; EMS 20411)
• Epoxy resin (LX-112 embedding kit; Ladd Research 21210)
• LX-112 (glycerol polyglycidyl ether; Ladd Research 21310)
• DDSA (dodecenyl succinic anhydride; Ladd Research 21340)
• NMA (nadic methyl anhydride; Ladd Research 21350)
• DMP-30 (2,4,6-tri[dimethylaminomethyl] phenol; Ladd Research 21370)
• Absorbent paper with plastic backing
• Disposable pipets (some with their tips trimmed – e.g., for LX-112 components, for transferring tissue)
• Kimwipes
• 100-ml beaker and a magnetic stirrer
• 20-ml syringes and syringe filters (PES membrane; 0.2-µm pore size) × 4 each.
• 20-ml borosilicate glass scintillation vials with caps (e.g., EMS 72634) × 10 (or more depending on the number of tissue), labeled as below:
  
  • OsO₄
    
  • 1
  • 2
  • KFeCN
  • PbA (× 3 vials or more)
  • UA (× 1 vial or more)
• A Pasteur pipet and bulb
• Shell vials (EMS 72631-10 [3.4 ml]), 1 per tissue.
• Embedding mold (e.g., Chien mold, EMS 70140)
• Applicator sticks
• Razor blade
• Block labels
  • Suggested label format:
  • • Font: Tw Cen MT Condensed, 9 pt., with 8 pt. spacing between lines
    • Width = 0.5 in. max; Height = 2 lines max
    • Label information should at least contain: Animal ID (e.g., MK01) and Tissue ID (e.g., R42CA1, for right hemisphere, vibrasection 42, area CA1)
• Aluminum foil for preparing LX-112, as well as for wrapping open OsO₄ ampoule and used Pasteur pipet
• Tri-pour beakers:
  • 6 × 250-ml beakers for mixing LX-112 components (see 2.3)
  • 1 × 250-ml for collecting open OsO₄ ampoule and used Pasteur pipet
  • 1 × 250-ml for washing the processing basket in 0.1 M Na cacodylate buffer, and then for collecting liquid waste during the procedure (labeled “Waste cacodylate, osmium, UA-ethanol”).
  • 1 × 100-ml beaker for infiltration reagents (labeled “ethanol, propylene oxide, and LX-112”)
• Solid waste bags for UA and non-UA chemical-contaminated waste.
• Liquid waste bottles:
  • Formaldehyde, glutaraldehyde, and cacodylate
  • OsO₄, KFeCN, and cacodylate
  • Uranyl acetate and water
  • lead aspertate and potassium hydroxide
  • Flammables
  • Resin and propylene oxide

2.  Reagent/Equipment Preparation (the day before, or on the day of processing)

2.1.  Hybridization incubator

• remove incubation tube.
• turn on the incubator.
• keep the rotisserie off until ready to use.

2.2.  Epoxy resin (can be done the day before)

1. Place stir bars (kept in acetone) in three tri-pour beakers and label "A", "B", and "A+B".
2. Check the weight per epoxide (WPE) of LX-112 indicated on the bottle and record the value in worksheet.
3. Refer to Table 1 to determine the correct amounts of DDSA and NMA. Record the amounts in worksheet.
4. Measure resin components with the scale in fume hood, using disposable pipettes with their tips cut off. 
   
   1. Beaker A: mix DDSA and LX-112 (see Table 2)
   2. Beaker B: mix NMA and LX-112 (see Table 2)
   3. The recipe here should make enough resin for embedding up to 24 pieces of tissue.  
5. Into beaker A+B, add the contents of beakers A and B (see Table 3 for the amounts).  Cover with foil and mix for another 15 min.
6. Add DMP-30 (see Table 3 for the amount) and mix thoroughly for another 15 min. If mixing resin the day before, wait until the day of use to add DMP-30.

2.3.  Reagents for en bloc staining

2.3.1.  Handling sodium cacodylate buffer

**under construction**

2.3.2.  Making stock solution of uranyl acetate, 1% aqueous (Start the day before processing)

1. Wear appropriate PPE (see above).  Place a piece of absorbent pad on work surface in fume hood.
2. Retrieve the following:
   1. Clean 20-ml scintillation vial(s)
   2. A 10-ml serological pipet with pipetter
   3. Solid uranyl acetate (in the desiccator cabinet)
   4. purified water
   5. Kimwipes
   6. A small piece of Parafilm (~1 in. × 1/2 in.)
   7. aluminum foil
   8. “Solid Waste – UA” bag
   9. Sonicator (stored in a cabinet under the lower fume hood), filled with some water
3. Figure out how much uranyl acetate you'll need: ~2 ml per tissue = 0.02 g of UA per tissue
   • Because it is difficult to measure a small amount of UA in a fume hood, it is recommended that you make minimum of 10 ml of this solution.
4. Open bottle of uranyl acetate powder in the fume hood.
5. Gently tap out approx. 0.1 g of uranyl acetate onto the weighing boat, then carefully pour into the scintillation vial. Keep the weighing boat and note the exact amount of uranyl acetate.
6. Based on the amount of uranyl acetate, figure out the volume of water necessary to make the final concentration of 1% (weight-by-volume; e.g., 10.4 ml water for 0.104 g of UA).
7. Use 10-ml glass pipet to dispense purified water into the weighing boat to collect any remaining uranyl acetate.  Pour this into the vial.
8. Cap the vial, wipe the bottle exterior with wet Kimwipes, and wrap around the cap with a piece of Parafilm. Wrap the capped vial in aluminum foil, and store in a secondary container at RT until use.
9. Dispose of the weighing boat and other supplies contaminated with uranyl acetate (e.g., used Kimwipes and outer layer of gloves) in “Solid Waste – UA” bag.
10. Clean and return supplies and equipment to their storage locations.
11. On the day of use, remove the foil from the vial, loosen the cap slightly, and sonicate for at least 15 min.

2.3.3.  Dispensing osmium tetroxide solution from glass ampule (on the day of processing)

1. Wear appropriate PPE (see above).  Place a piece of absorbent pad on work surface in fume hood.
2. Retrieve the following:
   1. A Pasteur pipet and bulb
   2. A 20-ml borosilicate glass scintillation vial without cap (labeled “OsO₄” and placed in the vial rack)
   3. A 250-ml tri-pour beaker
   4. A piece of Aluminum foil
   5. A glass 10-ml serological pipet with pipetter
   6. A 10-ml ampule of 4% aqueous solution of osmium tetroxide (stored encased in a plastic sleeve in a metal can in the fridge)
   7. A pair of forceps (located in fume hood)
   8. purified water
   9. "Solid Waste – No UA" bag
   10. "Waste OsO₄-KFeCN" bottle
3. Open the metal can to retrieve a 10-ml ampule of osmium tetroxide solution. Keep the ampule encased in the plastic sleeve.
4. Close and return the metal can to the fridge.
5. In fume hood, while still in the plastic sleeve with the red cap on, break open the ampule.
6. Remove the red cap and discard into "Solid Waste – No UA" bag.
7. Remove the broken ampule top with forceps and place on the foil. Keep the ampule bottom (with osmium tetroxide solution) in the plastic sleeve.
8. Use Pasteur pipet to dispense osmium tetroxide solution into a 20-ml borosilicate glass scintillation vial.
9. Once empty, keep the pipet tip in the ampule bottom, remove the pipet bulb, place the ampule top into plastic sleeve, and loosely wrap them in the foil.  Place the foil-wrapped waste into a 250-ml tri-pour beaker and discard into "Solid Waste – No UA" bag.
10. Dispense osmium tetroxide solution out of the scintillation vial for tissue processing using a glass serological pipet.
11. Discard any remaining osmium tetroxide solution into "Waste OsO₄-KFeCN" bottle.
12. Discard the scintillation vial into "Solid Waste – No UA" bag.

2.3.4.  Reagents for processing up to 6 pieces of tissue (on the day of processing)

• In 15- or 50-ml conical tubes, add:
  • ddH₂O: 50 ml at RT (~150 ml total will be needed); additional ~100 ml should be warmed to 50°C (use a water bath or hybridization incubator)
  • Na cacodylate buffer (SCB; 0.2M): ~7 ml
  • Na cacodylate buffer (0.1M): 50 ml (~100 ml total will be needed)
• In the labeled scintillation vials (Mix well by shaking):

              * See above (2.4.3) for how to dispense 4% OsO₄.

**Reduced osmium:

1. Add 3 ml of KFeCN solution to vial 2.
2. Place vials 1 and 2 on ice.
3. When ready, pour vials 1 and 2 back and forth to mix well before adding to tissue.

***2% OsO₄: Keep vial on ice until use.

2.3.5.  Making stock solution of Lead Aspartate (perform on Day 1 of processing)

1. Mix 0.16 g of Aspartic Acid in 40 ml of purified water. Stir > 1 hr on stir plate. 
2. Adjust pH to 3.8 with 1 M KOH and then add 0.26 g lead nitrate to solution.
3. Adjust pH to 5.5 with KOH
4. Store and label solution in scintillation vials and place in refrigerator.
5.  *heat solution at 60°C in oven for 15 min and filter before use*

2.4.  Reagents for infiltration

2.4.1.  Handling propylene oxide

**under construction**

2.4.2.  Reagents for infiltration (on the day of processing)

• Prepare ~2 ml per tissue of the following:
  • 1:1 = EtOH : propylene oxide (PO)
  • 1:2 = EtOH : PO
  • 100% PO
  • 1:1 = PO : Resin
  • 1:2 = PO : Resin

3.  Processing (4 days; all steps [staining, dehydration, infiltration] on rorator at RT, unless otherwise specified)

Day 1 (estimated work duration: 2 hrs + enough time for dissection and agarose embedding)

1. Place a piece of absorbent paper on work surface in the fume hood.
2. Prepare reagents and equipment, as described in Section 2 above.
3. Fill shell vials with SCB (~2 ml each).
4. Transfer the tissue (embedded in 7-9% agarose) to the shell vials (one each).
5. SCB washes, 5 min × 5.
6. Reduced osmium (1% OsO₄ + 1.5% K₄Fe(CN)₆ in SCB), 5 min.
   
   • Pour vials 1 and 2 back and forth to mix before adding to tissue.
7. SCB washes, 5 min × 3.
8. Purified water washes, 5 min × 3.
9. 2% aqueous OsO₄, 30 min.
   
   • Sonicate 1% aqueous UA solution at this point.
10. Purified water washes, 5 min × 5.
11. 1% aqueous UA, overnight at RT.
    • Dispense the UA solution with syringe-filter unit (0.1 um pore)

Day 2 (estimated work duration: 7 hrs )

1. Make a fresh batch of resin (See 2.3.). This step can be done while performing step 2.
2. 1% aqueous UA, 120 min at 50°C.
   • Place the vials into hybridization incubator (inside hybridization tube), rotate at the lowest speed.
3. Purified water washes, 5 min x 5 at 50°C
4. Lead aspartate, 120 min at 50°C
5. Purified water washes, 5 min x 3 at 50°C
6. Purified water washes, 5 min x 3
7. 50% EtOH, 5 min.
8. 70% EtOH, 5 min.
9. 100% EtOH, 5 min.
10. 100% EtOH, 5 min.
11. 1:1 = EtOH : PO, 10 min.
12. 1:2 = EtOH : PO, 10 min.
13. 100% PO, 15 min × 2.
14. 1:1 = PO : Resin, 1 hr.
15. 1:2 = PO : Resin, overnight.

Day 3 (estimated work duration: 3-4 hrs)

1. Make a fresh batch of resin (See 2.3.).
2. Infiltration with 100% Resin, 1hr × 3.
3. Place block labels writing side up in embedding molds and cover with a small amount of fresh resin.
4. Bevel a wooden applicator stick with a razor blade.  Using the stick, transfer the tissue from shell vials into the mold.
5. Under a stereomicroscope, move the tissue to desired position. 
   
   • Small blocks of polymerized resin can be used to support the tissue inside the mold.
6. Make sure that the resin is slightly convex from top of the mold.
7. Polymerize resin for 48 hr at 60°C.
   
   
   

Day 4

Continue resin polymerization. You may check and adjust positioning at this point. 

Day 5 (estimated work duration: 1-2 hrs)

1. Take the resin-embedded tissue blocks out of oven.  Remove the blocks out of the mold immediately and store in the cardboard pill boxes labeled with block identification numbers. 
2. Take solidified resin waste into “Solid Resin” waste drum.
3. Examine the blocks under the stereomicroscope and record their images before trimming and cutting.
4. Scan worksheet into a pdf file.

4.  Clean-up

4.1.  Waste containers

• Hazardous Liquid Waste: Pour all waste into the proper waste collection bottles available in fume hood.
  • Aldehyde-Cacodylate (fixative solution, cacodylate buffer)
  • Osmium-Ferrocyanide (reduced osmium/osmium tetroxide): Discard excess K₄Fe(CN)₆ and OsO₄.
  • Uranyl acetate-water (aqueous UA solution)
  • Lead aspertate-potassium hydroxide
  • Flammable Solvents (propylene oxide and alcohols that do NOT contain other chemicals such as uranyl acetate and Epon)
  • Resin-PO (mixture of LX-112 and propylene oxide)
    • One resin beaker should be used to collect all excess LX-112 resin and aluminum foil, plastic pipets, applicator sticks, and vials (uncapped) used with it.  This should be cured in the oven with the blocks and discarded in hazardous solid waste container labeled “solid LX-112 resin”.
• Hazardous Solid Waste: Place all contaminated solid waste (e.g., gloves, pipets, vials, processing dishes, etc.) into hazardous waste bags in fume hood.  All vials must be uncapped and empty.  Make sure to separate all waste contaminated with UA.

4.2.  Used supplies and equipment

• Supplies (e.g., vials and dishes) that came in contact with UA should be rinsed with ethanol or water into the UA waste bottle and disposed of in the solid UA waste bag.  All vials must be uncapped and empty.  Make sure to separate all waste contaminated with UA.
• Used absorbent paper should be discarded into the UA solid waste bag.
• Wipe inside hybridization incubator with wet Kimwipe (discard into the UA waste bag).
• Hybridization tube: First rinse with water (discard into UA-water waste bottle), then wash the bottle with lab soap. Rinse extensively with RO water before rinsing twice with purified water. Air dry completely.
• Stir bars (for resin mixing) and processing baskets should be placed in jars of acetone in the fume hood.
• Monitor the area for radioactivity (see Harris lab SOP for uranyl acetate).