Histology Protocol 1

Histology Protocol

Methods for histology protocol follows these basic procedures

• Stepwise infiltration with ethanol (graded percentages of alcohol) to dehydrate tissue
• Using molds to embed tissue onto microtome chucks
• Tissue sectioning onto slides using a typical sliding microtome
• Tissue water bath and positioning tissue onto slides
• Staining tissue (using Cresyl violet)
• Cover slipping slides using Permount
• Unbiased cell counting using optical fractionation

Contents 1 Infiltration 2 Tissue Sectioning 3 Staining 4 Cover-Slipping 5 Cell Counting Using Stereo Investigator

Infiltration

This is a typical stepwise infiltration schedule using increasing percentages of alcohol:water in order to dehydrate tissue. This specific schedule is adapted from a protocol for Technovit 7100 plastic; however wax paraffin embedding follows the same basic procedures.

• Label 20ml scintillation vials with tissue ID information
• If tissue sample is larger than 1.5 cubic centimeters, cut tissue in half
• Place tissue into vial and fill vial with the appropriate concentration of infiltration solution (below)

Day 1 (24 hours) - 70% EtOH 30% ddH2O
Day 2 (24 hours) - 80% EtOH 20% ddH2O
Day 3 (24 hours) - 95% EtOH 5% ddH20
Day 4 (4 days) - 25% Infiltration Sol. 75% EtOH (100% concentration)
Day 8 (4 days) - 50% Infiltration Sol. 50% EtOH (100% concentration)
Day 12 (4 days) - 75% Infiltration Sol. 25% EtOH (100% concentration)
Day 16 (4 days) - 100% Infiltration Sol
Day 20 (24 hours) embed into molds
Day 21 (24 hours) embed molds onto chucks

Day 1 Notes: Put tissue into the vial containing 70% EtOH. Put them onto a rotation table in a plastic container with a lid. Optimal setting for rotation table is around 55 RPM. Set time accordingly. Lay the vials perpendicular as to maximize infiltration solution surface area, which allows the solution to circulate more freely. Minimize light factor. Leave on table for 24 hours.

Day 4 Notes: Infiltration solution is made by using the Technovit kit components. Mix 1 bag (1g) of activaor with 100 ml of the Technovid liquid solution in a flask using a stir bar, but do not heat. This makes a 100% infiltration solution. This is the minimum amount you can make. To make a 25% infiltration solution, mix the 100% infiltration solution with 100% EtOH in the desired amount.

Day 21 notes: Embed in molds and harden for 24 hours. A small piece of kim-wipe should be placed between the cut edge of the tissue and the bottom of the mold in order to allow the hardening solution to completely engulf the tissue (this prevents any tissue from protruding out of the plastic when removed from mold.

Tissue Sectioning

Prior to sectioning using experimental tissue, it is wise to determine the necessary amount of slices per specimen that need to be saved. This can be done with a trial stereology run-through using practice tissue. By performing counts on every 5 section, then every 10 sections, then every 20 sections, one can find a happy balance between counting the least amount of slices while maintaining the necessary amount of statistical power. The experimental procedure is as follows.

• WEAR GLOVES to prevent oils from hands to touch slides or tissue (prevents tissue loss during staining).
• Mount chuck into microtome. Be sure tissue float is filled with deionized water and that the water is near 35 degrees Celsius.
• Set appropriate slice thickness (30 micrometers for brain tissue is a typical standard) and trim plastic/paraffin until the first tissue section is acquired. Reset counter to 0.000.
• Wet the embedded tissue using a brush (anti-static paint brushes work best) with clean D.I. water at a minimum of 3 slices prior to collected slice; that is, if you are collecting every 10th section, wet section number 8, wet section number 9, wet and collect section number 10.

! - If excessive amounts of plastic/paraffin are around the tissue section, use a scalpel to etch plastic around tissue on the 9th slice (1 slice prior to the collected section).>

• The wet slection to be kept should be collected off the blade with the brush (not your hands) and placed into the tissue bath.

! - This is done in order to prevent permanent tissue wrinkling which is cause by folding of dry plastic (folding of wet plastic is ok).>

! - There should be no tension or sticking of the plastic, if it becomes stretched it will cause the slice to later peel from slide.>

• Float the slice in the warm water for about a minute, the surface tension should cause the tissue to unfold and lay flat.

! - If the slice is still folded or sinks after 1 minute, use a tool with a flat surface to guide the slice out of the water and then slowly lowering the slice back into the water allowing the surface tension to make it lay flat.>

• If your slices was etched, gently remove any excess plastic from around the tissue.
• Place a +charged slide (Fisher Superfrost Plus) into the water beneath the floating tissue, and guide the flattened tissue onto the slide.

! - Do not manipulate the tissue after it has settled on the slide, this will cause the tissue to stretch and as the tissue dries it will peel from slide.>

! - If tissue collection fails, discard tissue and collect the next slice on the chuck.>

• Once the slide is full, place vertically for about 5 minutes to dry and then place at 45 degree angle for 24 hours prior to staining.

Staining

In order to facilitate penetration of the H2O based stain, mounted specimen sections are softened using ethanol and rehydrated to accept stain. Adequately stained specimens are then differentiated and again ehydrated through increasing concentrations of ethanol. Finally the specimens are cured and covered using xylene and a xylene compatible adhesive.

30 s each:
	1.  100% EtOH
	2.  95% EtOH
	3.  50% EtOH
	4.  100% dd H2O

20 minutes
	5.  Cresyl Violet1

30 s each:
	6.  100% dd H2O
	7.  50% EtOH
	8.  Differential solution2
	9.  95% EtOH
	10.  100% EtOH

Prep Cresyl Violet Stain (~400 ml)

• 90% EtOH (1 L)
  • add 52 ml ddH2O to 948ml 95% EtOH

• 6% Acetic Acid (Differential Solution (400 ml)
  • add 24 ml glacial acetic acid to 376 ml 90% EtOH

• 0.2 M Acetate Buffer
  • add 1.64 g anhydrous sodium acetate to 100 ml H2O

Cresyl Violet solution (400 ml)

Combine

• 0.8 g cresyl violet acetate
• 40 ml 0.2 M Acetate Buffer (pH 4.8)
• 120 ml 100% EtOH
• 240 ml dd H2O
• Stir for 1 h
• Filter solution into a clean staining container
• Use Qualitateive paper filters

Cover-Slipping

Coverslip using Permount

Application: Permount is clear with permanent adhering qualities. Absolutely neutral. Will not become acidic or discolor with age. 115C melting point makes it more suitable than balsam or gum damar for microprojection work. Refractive index of Permount is slightly higher than that of Canada balsam. Can be used with either artificial light or daylight. Permount is thinner than commonly used balsam and damar solutions, hence easier to apply. Has little tendency to trap bubbles beneath cover glass.

Notes:

1. Place the slide on a clean horizontal surface, preferably upon an unused kim-wipe.
2. Apply a drop of Permount to each tissue section and also put a drop of Permount at the far end of the slide away from the frosted end.
3. Carefully apply cover-slip by placing one end of slip on top of the drop of Permount at the far end of the slide. SLOWLY AND GENDLY "roll" the cover-slip down the slide, only using enough pressure to allow the medium to spread evenly. Do not attempt to clean excess Permount from edges of the slide. This will usually result in spreading the medium upon the surface of the cover-slip, resulting in a reduction in clarity.
4. Place the slides flat upon a kim-wipe in the bench-top slide warmer for 15-20 minutes. The slide warmer should be set at ~70 C. This will help to eliminate any bubbles under the cover-slip. Once the bubbles dissipate, decrease the temperature of the slide warmer to around 50 C for 1 hour.
5. Remove the slides from the warmer and allow them to cool to room temperature for 24 hours. The slides are now ready for microscopic visualization. It was found that some particular tissue sections, especially gastrointestinal sections or necrotic areas on the sections, have a significant number of microscopic air bubbles when Permount is not completely dry. Thorough drying of the sections cover-slipped with Permount helps to reduce this limitation.

Cell Counting Using Stereo Investigator

1. Turn on Opti Scan
2. Open Stereo Investigator Software
3. Turn on microscope
4. Set a reference point
5. Tools - serial section manager - new - enter parameters
6. Options - preferences - line diameter = 10 (be sure that scope is set at 4x)
7. Outline section
8. Oil Section
9. Move microscope and program to 60x
10. Focus on the very top of tissue
11. Move - Set Z stage - ok
12. Probes - Optical Fractionator
13. Parameters for OF

	Scan grid size = 1000 x 1000
	Dixector Probe
		Distance from top = 2.5
		3D frame height = 12.0
		Section Thickness = 25.0