Histological and histochemical methods

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Hematoxylin - Eosin Stain (H&E)

H&E is a good general stain, staining nucleic acids in the nucleus blue and the cytoplasm pink.

Procedure:

1. Cut tissue sample on a microtome 10 to 12 µm thick and place onto microscope slides (microscope slides should be pre-rinsed/cleaned in an ethanol solution). Leave slides at room temperature for at least 5 min.

2. Fix sections in 4% formaldehyde-Ca²⁺ (see notes) for 3-5 min.

3. Rinse for 5 min in dH₂O.

4. Stain sections with Hematoxylin (Haemalaun acc. Meyer) for 5 min.

5. Rinse continuously under tap water for 20 min to rinse off remaining Hematoxylin stain.

6. Rinse for 5 min in d₂H₂O.

7. Stain sections with a 0.1 % solution of Eosin (Potassium salt) in dH₂O for 10 min.

8. Rinse for 3 min in dH₂O.

9. Wash with: 70% Ethanol for 1 min

96% Ethanol for 1 min

absolute Ethanol for 3 min

10. Fix with Xylol for between 5 and 60 min (dispose of Xylol in a proper receptacle after use).

11. Mount a cover slip using a non-aqueous cover slip medium (ie. Histofluid).

Notes:

- To prepare the 4% Formaldehyde solution you add 1 g CaCl2 (water-free - VERY important) for every 100 mL of 4% formaldehyde. A 4% formaldehyde solution can be made from stock 37% Formalin by dilution. To the CaCl2/4% formaldehyde solution you add CaCO3. You need to add enough so that it does not dissolve and it is visible at the bottom of the jar. Basically the solution has to be supersaturated. In our solution you can see about 2 mm of CaCO3 on the bottom of the jar in a slurry. Just make sure to filter the solution before using it. The CaCO3 will go into solution so make sure that you add enough. It should settle to the bottom after awhile.

- The hematoxylin solution is purchased pre-mixed. We use Mayers Hematoxylin.
The 0.1% Eosin solution is made with water. Simply mix 0.1 g of Eosin in 100 mL of water.

- Unless otherwise noted solutions are to be kept at 4 °C, are good for up to one month, and can be disposed of down the sink.

Aniline Blue - Orange G Stain

This stain provides a good contrast between muscle fibers (orange), connective tissue (blue), and fat cells (white).

Procedure:

1. Cut tissue sample on a microtome 10 to 12 mm thick and place onto microscope slides (microscope slides should be pre-rinsed/cleaned in an ethanol solution). Leave slides at room temperature for at least 5 min.

2. Fix sections using Müller´s reagent for 1 min.

3. Rinse for 1 to 2 min in d₂H₂O.

4. Stain sections with Orange G for 3 min.

5. Rinse for 2 min in dH₂O (twice).

6. Stain sections with Aniline Blue for 1 to 3 seconds.

7. Rinse (with gentle shaking) immediately for 2 to 5 minutes in dH₂O (twice) (until no blue color is seen in the water)

8. Wash with: 70% Ethanol for 5 to 10 sec

96% Ethanol for 5 to 10 sec

absolute Ethanol for 1 to 2 min

9. Fix with Xylol for between 5 to 60 min (dispose of in proper receptacle).

10. Mount a cover slip using a non-aqueous cover slip medium (i.e. Histofluid).

Solutions and Reagents:

Müller´s reagent (Fresh Daily)

Potassium dichromate (K₂Cr₂O₇) 2.5 g

Sodium sulfate (Na₂SO₄-10H₂O) 1.0 g

dH₂O 100 mL

Phenol pinch (tip of a scupula, see notes)

Orange G solution (Store at room temperature, good for over 1 year)

Orange G 3.0 g

Acetic Acid, conc. 5.0 mL

dH₂O 100 mL

Bring the solution to a boil, cool to room temperature, and filter.

Aniline Blue solution (Store at room temperature, good for over 1 year)

Aniline Blue 0.5 g

Acetic Acid, conc. 5.0 mL

dH₂O 100 mL

Bring the solution to a boil, cool to room temperature, and filter.

Notes:

- The phenol added to the aniline blue/orange G stain is used as a preservative to stop bacterial growth. The recipe is over 100 years old and was used when water sources were not as clean as they are today. Imagine the end of a pen and use an amount of phenol equivalent to the tip of the pen. Its basically just a few grains.

Mitochondria (Nitro BT) Stain

This stain preferentially locates muscle fiber mitochondria. Muscle fiber type can thus be distinguished by the density of mitochondria. Discrimination is observed between three fiber types, oxidative fibers, type I (high density), glycolytic fibers, type IIB (low density), and an intermediate fiber type, type IIA (medium density).

Procedure:

2. Fix sections in 4% formaldehyde-Ca²⁺ for 45 sec (muscle samples 24 h post mortem) or 5 min (muscle samples 30 min post mortem).

3. Rinse for 5 min in dH₂O.

4. Incubate sections in Staining Medium for 60 min at 37 °C.

5. Rinse for 5 min in dH₂O.

6. Mount a cover slip using an aqueous cover slip medium (ie. Glycerine Gelatine).

Staining Medium (Fresh Daily)

NADH-Na₂ 16 mg

Sodium Phosphate Buffer 0.1M (pH 7.4) 3.2 mL

Nitro-BT (4-nitro blue tetrazolium chloride) (1 mg/mL) 4.0 mL

DH₂O 4.8 mL

Notes:

- Unless otherwise noted solutions are to be kept at 4 °C and can be disposed of down the sink.

Reference

Novikoff, A; Shin, W.Y.; Drucker, J. (1961) Mitochondrial localization of oxidative enzymes: Staining results with two tetrazolium salts. Journal of Biophysical and Biochemical Cytology 9: 47-61

Myofibrillar ATPase Stain

This stain allows a clear discrimination between three fiber types, one slow-twitch fibre, type I (white fibres) and two fast-twitch fiber types, types IIA (light blue) and IIB (dark blue).

Procedure:

2. Wash twice for 1 min each with Tris-Ca²⁺ (Pre-Rinse Solution).

3. Incubate for 5 min in the Alkaline Pre-Incubation Solution.

4. Wash twice for 30 sec each with Tris-Ca²⁺ (Pre-Rinse Solution).

5. Incubate for 90 min at 37 °C in the Incubation Solution.

6. Wash 4 times for 20 – 30 sec each in CaCl₂ Wash Solution.

7. Rinse for 3 min in a 2% CoCl₂ Solution (mix immediately before use).

8. Rinse 4 times for 20 – 30 sec each in dH₂O.

9. Stain for 28 sec in a 1% Azure Stain (mix immediately before use, see notes).

10. Rinse continuously under tap water for 5 min to rinse off remaining Azure Stain.

11. Rinse once with dH₂O.

12. Wash with: 50 % Ethanol for 5 to 10 sec

70% Ethanol for 5 to 10 sec

96% Ethanol for 5 to 10 sec

absolute Ethanol for 1 to 2 min

13. Fix sections using a 1:1 (v:v) solution of Xylol:absolute Ethanol (dispose of in proper receptacle after use).

14. Mount a cover slip using a non-aqueous cover slip medium (i.e. Histofluid).

Solutions and Reagents:

Pre-Rinse Solution (Fresh Daily) 100 mL 0.18 M CaCl₂

(pH 7.3) 12.1 g Trishydroxymethylaminomethane

add dH₂O to 1000 mL

Alkaline Pre-Incubation Solution 10 g CaCl₂

(pH 10.4 with 1 N NaOH) 7.44 g Glycine

100 mL Formaldehyd (37%)

add dH₂O to 1000 mL

Incubation Solution (Fresh Daily) 20 mL 0.1 M Glycine Buffer

(pH 9.4, warm to 37 °C) 10 mL 0.18 M CaCl₂(2%)

0.152 g ATP

add dH₂O to 100 mL

0.1 M Glycine Buffer 125 mL Glycine (7.51g/250 mL dH₂O)

(pH 9.4) 42 mL 0.4 M NaOH (8 g/250 mL dH₂O)

add dH₂O to 500 mL

0.18 M CaCl₂ 19.98 g CaCl₂ in 1000 mL dH₂O

CaCl₂ Wash Solution 13.3 g CaCl₂ in 1000 mL dH₂O

Notes:

- Azure A is the most commonly used for this purpose. The 1% Azure A stain is made by dissolving 1 g of Azure A in 100 mL water.

- Unless otherwise noted solutions are to be kept at 4 °C, are good for up to one month, and can be disposed of down the sink.

Reference

Brooke, M. H., and K. K. Kaiser. 1970. Muscle fiber types: how many and what kind ? Arch. Neurol. 23:369-379.

Szentkuti, L., and A. Eggers. 1985. Eine zuverlässige Modifikation der Myosin-ATPase-Reaktion zur histochemischen Darstellung von drei Fasertypen in der Skelettmuskulatur von Schweinen. Fleischwirtsch. 65:1398-1404.