Paxgene-Fixed Cryo-Embedded Tissue H&E Staining Protocol

• Section tissues as written in the PFCE protocol
• This is heavily based on (& in large part copied from) the 10X H&E Staining protocol for fresh-frozen tissue, with modifications based on their protocol for fixed frozen tissue

This is a general protocol for H&E staining that is as RNAse-free and RNA-safe as possible based on what 10X has tested.

Materials

1. Cryosectioned slides (in -80 ºC freezer)
2. Dry ice
3. Incubator
4. RNase decontamination solution
5. 70% ethanol
6. Reagent reservoirs
7. RNAse-free 1X PBS
8. 6 1-L beakers
9. Hematoxylin Solution, Gill No. 2 (Sigma-Aldrich Cat# GHS232)
   1. concentration: 4 g/L
10. Eosin Y Solution, Alcoholic (Sigma-Aldrich Cat# HT110116)
    1. concentration: 0.5 % (w/v) in acidified ethanol
11. Bluing buffer
    1. I make a homemade version for normal H&E staining: https://ihcworld.com/2024/03/05/scotts-tap-water-substitute/
       1. Magnesium sulfate + sodium bicarbonate in RNAse free water (add exact amounts!!)
    2. 10X recommends Bluing Buffer, Dako
12. Glycerol
13. Wide-bore pipette tips
14. Cover slips (Fisherbrand Cat # 12-541-033)
    1. #1.5 thickness, 50 x 24 mm size
15. Nail polish (if sealing slides)

Preparation

1. Clean workspace, pipettes, and gloves with an RNase decontamination solution, followed by 70% ethanol.
   1. I usually then cover the bench with bench protector made of thick paper material to avoid staining the bench but this is optional
2. Preheat incubator to 37 ºC
   1. Ideally you could do this on a thermocycler using the 10X Thermocyler Adapter – I use the incubator genie
3. Warm RNase-free 1X PBS to 37ºC (need 2mL per slide)
4. Clean six 1-L beakers:
   1. Spray with an RNase decontamination solution and leave for 10 sec to 1 min.
   2. Spray with 70% isopropanol or 70% ethanol.
   3. Rinse with Milli-Q water.
      1. Perform these cleaning steps regardless if beakers have been autoclaved.
5. Dispense 800 ml of Milli-Q water into each beaker and label the beakers with #s 1-6.
6. Dispense fresh Alcoholic Eosin into a falcon tube or coplin jar
7. Prepare 85% glycerol solution in a 1.5 mL tube using a wide-bore pipette tip
   1. You need 100uL per slide
   2. This is harder than it sounds - go slow!

Staining

Max 2 at a time. Make sure whenever you place the slide down that the tissue faces up (!!!!). If you have them, use lint-free laboratory wipes. I use kimwipes but they are not recommended.

1. Remove slide in falcon tube or slide mailer from -80 ºC freezer and immediatelly bury in dry ice.
2. Transfer slide to pre-warmed 37ºC incubator for 15 minutes
   1. I place the slide in a cleaned glass petri dish in the incubator
   2. change from 10X protocol - 15 minutes instead of 1 minute
      1. I found that a 15 minute warming compared to a 1 minute warming improved tissue adherence immensely for my specific tissues
3. Place warmed slide on a reagent reservoir
   1. change from 10X protocol - all fixation + post fixation wash steps skipped, tissue already fixed
      1. I tested this and it led to worse outcomes for my specific tissues
4. Immediately cover each slide with 1 mL of warmed 1X PBS
   1. change from 10X protocol: 1X PBS used instead of enhancer mix, but same steps
5. Pour this off immediately into the reservoir and place over the reservoir again (“pour off” from now on)
6. Cover with another 1 mL of warmed 1X PBS
7. Incubate for 1 minute
8. Pour off 1X PBS
9. Add 1 ml Hematoxylin per slide
   1. To avoid disturbing tissue, pipette at least 1 cm away from the tissue edge and avoid pipetting at an angle.
10. Incubate 1 minute
11. Pour off Hematoxylin
12. Immerse slides 5x in Water Beaker 1.
13. Immerse slides 15x in Water Beaker 2.
14. Immerse slides 15x in Water Beaker 3.
15. Wipe excess liquid from back of slides without touching the tissue + return to reservoir
16. Add 1 ml Bluing Buffer per slide
17. Incubate 1 minute
18. Pour off.
19. Immerse slides 10x in Water Beaker 4.
20. Wipe excess liquid from back of slides without touching the tissue
21. Gently immerse slides in alcoholic Eosin solution in coplin jar
22. Incubate 30 sec
23. Quickly and gently remove excess eosin by tapping the bottom edge of the slide to a kim wipe (at an angle)
24. Immerse slides for 30 sec in Water Beaker 5.
25. Immerse slides 10x in Water Beaker 6.
26. Wipe excess liquid from back of slides without touching the tissue + return to reservoir or other non-absorbent surface (not on a kimwipe)

Coverslip mounting

1. Remove excess moisture from slide without touching tissue
   1. Pipette off big drops, use lint-free laboratory wipes, or very carefully use kimwipes to wick away moisture.
2. Place slides on a flat, clean, nonabsorbent work surface. Some residual droplets may remain.
   1. Don’t let the tissue dry
3. Using a wide-bore pipette tip, add 100 µl 85% glycerol to cover the entire tissue section (or less as needed)
4. Slowly apply coverslip at an angle on one end of the slides, without introducing bubbles
5. Let the glycerol spread and settle on a nonabosrbent surface for a few seconds, then wick away any excess from the side of slide using a kimwipe - do not move coverslip.
6. Seal slide with nail polish.

Protocol credit goes to 10X fresh-frozen tissue H&E protocol, and I have made modifications for coral tissue fixed in Paxgene fixative that has been decalcified + cryosectioned.