en bloc only staining and epoxy embedding of chemically fixed tissue (70-250 µm thick tissue)

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

  • review relevant Safety Data Sheets and Harris Lab SOP (also see below)

  • 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:

    • Acetonitrile: flammable; acute toxicity (oral, dermal, inhalation); irritant (eye)

    • Epoxy resin components

      • EMBed-812: irritation (eye, skin, respiratory, oral)

      • NSA: irritant (eye, skin, respiratory)

      • NMA: irritant (eye, skin, respiratory)

      • BDMA: permanent eye damage, irritant (skin, respiratory, oral)

    • Osmium tetroxide: acutely toxic; mutagenic

    • Potassium ferrocyanide: contact with acid releases very toxic gas

    • Pyrogallol: acute toxicity (oral, inhalation, dermal); mutagen; aquatic hazard

    • Sodium cacodylate: carcinogen; irritant (skin, eye); skin permeator

    • Uranyl acetate: fatal (inhalation, ingestion); flammable

    • Acetonitrile, osmium tetroxide, sodium cacodylate, and uranyl acetate must be handled only under a chemical fume hood.

• 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

• Read the following paper:

  • Lu et al. (2023) A Scalable Staining Strategy for Whole-Brain Connectomics. bioRxiv https://doi.org/10.1101/2023.09.26.558265

1.  Reagents and supplies

• Worksheet for this procedure [under development – ask Masa for the current version]

• Vibraslices (70 - 250 µm thickness) of chemically fixed brain tissue, or fixed acute brain slices used for electrophysiology experiments.

  • If the thickness is 100 µm or less, protect the tissue by embedding it in two glass coverslip thickness of 7-9% agarose.

• H₂O (purified water: double-distilled, ASTM type I, WFI grade, or equivalent; e.g., Fisher 9150-25 or VWR RC915025)

• 0.3 M sodium cacodylate buffer (SCB) with 4 mM CaCl₂ and 8 mM MgSO₄: See 2.3.1

• 0.15 M SCB with 2 mM CaCl₂ and 4 mM MgSO₄

• 0.3 M NaCl (aq)

• 0.15 M NaCl (aq)

• 4% aqueous OsO₄ (osmium tetroxide; stored at 4°C; EMS 19190): See 2.3.2

  • Use a fresh ampule for each processing; discard excess

• K₄Fe(CN)₆ · 3H₂O (potassium ferrocyanide, or KFeCN; Sigma-Aldrich P3289)

• Pyrogallol (Sigma-Aldrich 16040)

• Acetonitrile (EMS 10020 or 10021)

• 2% uranyl acetate (UA; EMS 22400) in water (stored at 4°C): See 2.3.1

• Epoxy resin (modified EMBed-812 recipe; EMS)

  • EMBed-812 (glycerol polyglycidyl ether; EMS 14900 or 14901)

  • NSA (noneyl succinic anhydride; EMS 19050 or 19051)

  • NMA (nadic methyl anhydride; EMS 19000 or 19001)

  • BDMA (benzyldimethylamine; EMS 11400 or 11400-25)

• Absorbent paper with plastic backing

• Disposable pipets (some with their tips trimmed – e.g., for transferring tissue)

  • We like fine tipped ones (e.g., Fisher 13-711-26) for measuring out BDMA.

• Kimwipes

• orbital shaker

• ice bath that fit onto the shaker (e.g., a Styrofoam box)

• Processing well inserts (Ted Pella 36169, 36171, or 36173), 1 per tissue

  • Ted Pella 36169 is good for parasagittal sections of prefusion-fixed rat brains. Six of these will fit into one tray.

• Polypropylene tray and lid, 2 sets (or more)

  • Tray: Chemglass Life Sciences CLS3590384 (Fisher 50-194-4006)

  • Lid: Thermo Scientific AB0755 (Fisher AB-0755)

• Instead of the well inserts in a tray, you can use glass shell vials or scintillation vials.

  • Choosing containers for tissue processing:

Tissue processing tray:

• 50- or 60-ml syringes

• syringe filters (Nylon membrane; 0.2-µm pore size) × ?

• 20-ml borosilicate glass scintillation vials with caps (e.g., EMS 72634) × ?

  • or shell vials, depending on the size of your tissue

• A Pasteur pipet and bulb, a 20-ml borosilicate glass scintillation vial without cap (labeled “OsO₄”)

• Embedding mold (e.g., Slide duplicating mold, EMS 70170; Chien mold, EMS 70140)

• Applicator sticks

• Razor blade

• • Suggested label format for Chein mold:

  • 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 contain at least:

      • Animal ID (e.g., MK01)

      • Tissue ID (e.g., R42CA1, for right hemisphere, vibratome section 42, area CA1)

• Aluminum foil for preparing resin, as well as for wrapping open OsO₄ ampoules and used Pasteur pipets

• Tri-pour beakers:

  • 6 (or more) × 250-ml beakers for mixing resin components (see 2.2)

  • 1 × 250-ml for collecting open OsO₄ ampoule and used Pasteur pipet

  • 1 × 250-ml for collecting liquid waste during the procedure (labeled “Waste cacodylate, osmium, UA”).

• 50-ml conical tube for infiltration reagents (labeled “acetonitrile and resin”)

• Waste containers:

  • Solid waste bags for UA and non-UA chemical-contaminated waste.

  • Liquid waste bottles:

    • Formaldehyde, glutaraldehyde, and cacodylate (use the waste container for perfusion-fixation)

    • OsO₄, KFeCN, and cacodylate

    • pyrogallol and cacodylate

    • Uranyl acetate and acetonitrile

    • Resin and acetonitrile

  • NOTE: Your institution may require a different setup for collecting waste.

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

2.1.  Tissue processing trays and wells

• The day before processing, add 50 ml of SCB to each processing tray and soak the wells overnight.

• Replace with fresh ice-cold SCB before adding tissue.

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 EMBed-812 indicated on the bottle and record the value in worksheet.

   1. Enter WPE value on this Resin Components Calculator to figure out the amount of resin components to use. Remember to record the amounts on your worksheet or notebook.

3. Measure resin components with the scale in fume hood, using disposable pipettes with their tips cut off. 

   1. Beaker A: mix NSA and EMBed-812

   2. Beaker B: mix NMA and EMBed-812

   3. The recipe here should make enough resin for embedding up to 24 pieces of tissue.  (This needs to be verified)

4. Into beaker A+B, add the contents of beakers A and B .  Cover with foil and mix for another 15 min.

5. Add BDMA and mix thoroughly for another 15 min. If mixing resin the day before, wait until the day of use to add BDMA.

• Read this page if the Resin Components Calculator is not working or you want to know how to calculate the amounts.

2.3.  Reagents for osmium fixation, dehydration, UA en bloc staining

2.3.1.  Making sodium cacodylate buffer

**under construction**

• 0.3M sodium cacodylate buffer with 4 mM CaCl₂ and 8 mM MgSO₄ (SCB)

  • In 700 ml purified water, dissolve:

    • 64.2 g sodium cacodylate trihydrate (Ladd)

    • 0.588 g CaCl₂·2H₂O

    • 1.972 g MgSO₄·7H₂O

  • Adjust pH to 7.4.

  • Bring the volume to 1 L

• 0.15M SCB with 2 mM CaCl₂ and 4 mM MgSO₄

  • Dilute 0.3M SCB 1:1 with purified water.

  • Alternatively, this solution can be prepared from the 0.2M stock (e.g., EMS 11653)

    • For 1 L, dissolve 0.294g of CaCl₂·2H₂O and 0.986 g of MgSO₄·7H₂O into 750 ml of 0.2M SCB and bring the volume to 1L with purified water.

2.3.2.  Making stock solution of uranyl acetate, 2% in water

1. Wear appropriate PPE (see above).  Place a piece of absorbent pad on work surface in fume hood.

2. Retrieve the following:

   1. Sonicator (stored in a cabinet under the lower fume hood), filled with some water

   2. A clean 100-ml graduated cylinder

   3. purified water

   4. Solid uranyl acetate (in the desiccator cabinet)

   5. Glass bottle containing 2% uranyl acetate (aq) solution (stored in a secondary container in the fridge)

   6. Kimwipes

   7. A small piece of Parafilm (~1 in. × 4 in.)

   8. “Solid Waste – UA” bag

3. Place a large plastic weighing boat on a scale in fume hood and tare.

4. Open bottle of uranyl acetate powder in the fume hood.

5. Gently tap out approx. 2 g of uranyl acetate onto the weighing boat, then carefully pour into the glass bottle. Keep the weighing boat and note the exact amount of uranyl acetate.

6. Based on the amount of uranyl acetate, measure out the volume of water necessary to make the final concentration of 2% (weight-by-volume; e.g., 98 ml water for 1.96 g of UA).

7. Pour a small volume of water from the graduated cylinder into the weighing boat to collect any remaining uranyl acetate.  Pour this into the bottle.

8. Pour the remaining water into the bottle.

9. Cap the bottle (but do not tighten), wipe the bottle exterior with wet Kimwipes, and sonicate for at least 15 min.

10. Tighten the cap, remove the bottle from sonicator, and wipe the bottle exterior with Kimwipes.

11. Wrap around the cap with a piece of Parafilm and place the bottle in a secondary container (e.g., 500-ml tri-pour beaker) before storing in the fridge.

12. 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.

13. Clean and return supplies and equipment to their storage locations.

14. glass bottle ordering info: Qorpak GLC-14183 (Safety-Coated Packer Bottles, Wide Mouth, Amber, 120ml, Green F217/PTFE Lined cap)

2.3.3.  Dispensing osmium tetroxide solution from a glass ampule

1. Wear appropriate PPE (see above).  Place a piece of absorbent pad on the work surface in the fume hood.

2. Retrieve the following:

   1. A Pasteur pipet and bulb

   2. A 20-ml borosilicate glass scintillation vial without a cap (labeled “OsO₄” and placed in the vial rack)

   3. A 250-ml tri-pour beaker

   4. A piece of Aluminum foil (large enough to wrap the beaker with pipet in it)

   5. A glass serological pipet

   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. "Solid Waste – No UA" bag

   9. "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 the fume hood, while still in the plastic sleeve with the red cap on, break open the ampule.

6. Remove the red cap and discard it into "Solid Waste – No UA" bag.

7. Remove the broken ampule top with forceps and place it on the foil. Keep the ampule bottom (with osmium tetroxide solution) in the plastic sleeve.

8. Use a 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 the 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.  Amount of reagents needed

2.3.5.  Handling acetonitrile

• Open and use acetonitrile only under a chemical fume hood.

2.3.6.  Reagents for dehydration and infiltration

• In conical tubes (15 ml or 50 ml, depending on your tissue/setup), prepare the following:

  • 1:3 = acetonitrile : 0.15 M NaCl (aq)

  • 1:3 = acetonitrile : water

  • 1:1:2 = acetonitrile : water : 2% UA (aq)

  • 1:1 = acetonitrile : 2% UA (aq)

  • 3:1 = acetonitrile : water

  • 1:1 = acetonitrile : Resin

  • 1:3 = acetonitrile : Resin

3. Tissue Processing Protocol

Day 1

1. Wash fixed tissues in 0.15 M SCB for 30 min in total, on ice.

   • Change buffer for at least three times: 10 min + 10 min + 10 min, or 5 min + 10 min + 15 min

   • Waste --> “Aldehydes-Cacodylate”

2. Make reduced osmium solution in SCB during wash

   1. for each of the 20-ml scintillation vial,

      1. solution 1: dissolve 1.2 g of potassium ferrocyanide in 20 ml of 0.15 M SCB

      2. solution 2: dilute 10 ml of 4% OsO₄ stock in 10 ml of 0.3 M SCB

   2. keep the vials on ice until use.

   3. immediately before use, mix solutions 1 and 2 into a processing tray. 40 ml total vol

3. Stain in reduced osmium for 45 min, on ice 

   • waste --> “KFeCN-OsO₄”

4. Wash in the buffer for 30 min in total, on ice

   • waste --> “KFeCN-OsO₄”

5. Make 320 mM pyrogallol (aq) during wash

   • Dissolve 2.01776 g pyrogallol in 50 ml water and keep on ice until use.

6. Stain in fresh 320 mM Pyrogallol for 30 mi, on ice 

   • waste --> “Pyrogallol”

7. wash in the buffer for 30 min in total, on ice

   • waste --> “Pyrogallol”

8. prepare 2% OsO₄ in SCB during wash

   • 1:1 dilution of 4% OsO4 stock solution in 0.3 M SCB, 40 ml total; keep on ice until use

9. stain in buffered 2% OsO₄ for 45 min, on ice 

   • waste --> “KFeCN-OsO₄”

10. wash twice in the buffer for 20 min in total, on ice

    • waste --> “KFeCN-OsO₄”

11. wash once in purified water for 10 min, on ice

    • waste --> “KFeCN-OsO₄”

12. prepare resin during wash, RT

13. dehydrate in series of acetonitrile solutions, 10 min per step, RT

    1. 1:3 = acetonitrile : water

    2. 1:1:2 = acetonitrile : water : 2% UA (aq)

    3. 1:1 = acetonitrile : 2% UA (aq)