Masson’s Trichrome Stain (Masson’s Trichrome) Standard Operating Procedure (SOP)

I. Purpose and Scope

1.1 Purpose

Masson’s trichrome staining is used to differentiate and visualize the distribution and relative proportion of tissue components in paraffin-embedded tissue sections. Typical readouts include:

(1) Connective tissue components such as collagen fibers, mucin, and cartilage.

(2) Muscle fibers, cytoplasm (cell cytoplasm), erythrocytes, etc.

(3) Cell nuclei.

This method is widely used for:

① Assessment of fibrosis severity in various organs (liver, heart, lung, kidney, etc.).

② Qualitative/semi-quantitative analysis of muscle fibers versus collagen fibers.

③ Observation of thrombi, fibrin deposition, basement membrane structures, and related features.

1.2 Applicable specimens

(1) Paraffin tissue sections (3–5 μm) fixed in routine 10% neutral buffered formalin.

(2) Specimens fixed with other fixatives may also be processed using this SOP; however, staining time and contrast may require minor optimization.

II. Principle and Staining Mechanism

2.1 Overview of staining principles

(1) Hematoxylin (Regaud or Weigert type) binds nucleic acids in nuclei, producing a black-blue nuclear stain and providing good resistance to subsequent acidic dyes.

(2) Ponceau–acid fuchsin is a mixed acidic anionic dye system that preferentially binds basic groups in proteins (e.g., cytoplasm, muscle fibers, fibrin), staining these structures red.

(3) Phosphomolybdic acid displaces part of the acidic dye from collagen fibers (acting as a differentiator for ponceau and acid fuchsin), thereby creating binding sites for the subsequent collagen-selective dye.

(4) Aniline blue or light green has strong affinity for collagen fibers and—after phosphomolybdic acid pretreatment—preferentially stains collagen blue (aniline blue) or green (light green).

(5) A low concentration of glacial acetic acid improves color purity and enhances contrast between red and blue (or red and green).

2.2 Interpretation of staining colors

(1) Collagen fibers, mucin, cartilage: blue (aniline blue) or green (light green).

(2) Muscle fibers, cytoplasm, fibrin, neuroglia: red.

(3) Nuclei: black-blue or dark blue.

III. Reagents and Materials

3.1 Specimen and slide preparation

(1) Fixation: 10% neutral buffered formalin fixation, typically ~24 h.

(2) Embedding: routine paraffin embedding.

(3) Sectioning: 3–5 μm thick sections mounted on adhesive-coated slides; bake at 60°C for 30–60 min.

3.2 Key reagent formulations

(1) Regaud’s hematoxylin

① Hematoxylin 1 g

② 95% ethanol 10 mL

③ Glycerol 10 mL

④ Distilled water 80 mL

Preparation notes:

① Dissolve hematoxylin in a portion of distilled water with gentle heating; cool, then add 95% ethanol and glycerol.

② Bring to final volume with distilled water and mix well.

③ Allow to ripen for several days at room temperature protected from light before use.

④ Filter before use.

(If Weigert iron hematoxylin is routinely used for nuclear staining in your laboratory, it may be substituted according to the current laboratory formulation.)

(2) Masson’s ponceau–acid fuchsin solution

① Ponceau (Ponceau red) 0.7 g

② Acid fuchsin 0.3 g

③ Distilled water 99 mL

④ Glacial acetic acid 1 mL

Preparation notes:

① Dissolve ponceau and acid fuchsin thoroughly in distilled water.

② Add glacial acetic acid and mix well.

③ Store at room temperature protected from light; filter before use.

(3) Aqueous glacial acetic acid solution

① 0.2% glacial acetic acid: 0.2 mL glacial acetic acid brought to 100 mL with distilled water.

(Some laboratories use 2% glacial acetic acid for a brief dip. This may be adjusted according to established SOPs, but should be standardized once set.)

(4) 1% phosphomolybdic acid aqueous solution

① Phosphomolybdic acid 1 g

② Distilled water to 100 mL

Dissolve completely before use and replace regularly.

(5) Aniline blue solution

① Aniline blue 2 g

② Distilled water 98 mL

③ Glacial acetic acid 2 mL

(6) Light green solution (optional; alternative to aniline blue)

① Light green 1 g

② Distilled water 100 mL

3.3 Other reagents and materials

(1) Xylene: deparaffinization and clearing.

(2) Graded ethanol: 100%, 95%, 80%, 70%.

(3) Tap water and distilled water.

(4) Neutral mounting medium (neutral resin).

(5) Routine staining rack, staining jars, microscope, etc.

IV. Procedure

4.1 Deparaffinization and rehydration to water

(1) Bake slides at 60°C for 30–60 min.

(2) Xylene, 2 changes, 10 min each.

(3) 100% ethanol, 2 changes, 5 min each.

(4) 95% ethanol, 80% ethanol, 70% ethanol, 2–3 min each.

(5) Rinse thoroughly in tap water, then rinse in distilled water; keep sections in distilled water until staining.

4.2 Nuclear staining

(1) Hematoxylin nuclear staining

① Stain sections in Regaud or Weigert hematoxylin for 5–10 min.

② Rinse thoroughly in tap water.

③ If nuclei are overstained, differentiate briefly in 1% acid alcohol for a few seconds.

④ Blue in tap water (for iron hematoxylin, bluing may be omitted or performed according to laboratory practice).

(2) Rinse in distilled water to remove salts from tap water.

4.3 Ponceau–acid fuchsin staining (cytoplasm/muscle)

(1) Immerse sections in Masson’s ponceau–acid fuchsin solution for 5–10 min (typically 5–8 min).

(2) Rinse quickly in distilled water for a few seconds to remove excess surface dye and prevent overly intense red staining that may obscure subsequent collagen staining.

4.4 Differentiation and collagen staining

(1) Acetic acid rinse

① Immerse sections in 0.2% aqueous acetic acid for ~30 s to 1 min.

② Gently agitate the staining jar to obtain clearer, more saturated tones.

(2) Phosphomolybdic acid differentiation

① Transfer sections into 1% phosphomolybdic acid for 3–5 min.

② During differentiation, check 1–2 slides microscopically: an appropriate endpoint is when muscle fibers and fibrin remain red while collagen fibers appear only faintly pink.

(3) Collagen staining (aniline blue or light green)

① Remove sections from phosphomolybdic acid, gently drain excess solution, and do not rinse in water.

② Immediately immerse in aniline blue or light green solution for ~5 min.

③ Gently agitate during staining to ensure uniform coloration.

(4) Acetic acid rinse

① Immerse sections in 0.2% aqueous acetic acid for ~30 s to 1 min.

② Gently agitate to remove excess unbound dye and enhance red/blue (or red/green) contrast.

4.5 Dehydration, clearing, and mounting

(1) 95% ethanol, 1–2 changes, 1–2 min each.

(2) 100% ethanol, 1–2 changes, 1–2 min each.

(3) Clear in xylene, 2 changes, 5 min each.

(4) Mount with neutral resin mounting medium and apply coverslip, avoiding bubbles; air-dry at room temperature.

V. Staining Results and Interpretation

5.1 Expected/ideal staining appearance

(1) Collagen fibers, mucin, cartilage:

① Aniline blue system: uniform blue.

② Light green system: uniform green.

(2) Muscle fibers, cytoplasm, fibrin, neuroglia:

Bright red to brick red with clear boundaries.

(3) Nuclei:

Black-blue or dark blue, with discernible nuclear membrane and chromatin structure.

5.2 Interpretation notes

(1) Color separation between collagen and muscle fibers should be high; tones should not appear muddy or gray.

(2) For fibrosis analysis, image analysis software (e.g., ImageJ/Fiji) may be used to quantify collagen area/volume fraction; ensure consistent staining conditions across batches.

(3) Abnormal findings should be interpreted in conjunction with H&E staining, clinical context, and other special stains.

VI. Quality Control and Common Troubleshooting

6.1 QC key points

(1) Include a known fibrosis-positive control tissue in each run to monitor staining consistency.

(2) Filter all dye solutions before use to prevent precipitates/crystals/particulates that may cause background contamination or focal uneven staining.

(3) Replace phosphomolybdic acid solution and Masson’s ponceau–acid fuchsin solution periodically based on usage frequency and staining performance.

6.2 Common problems and adjustment strategies

(1) Poor delineation between collagen and muscle fiber colors

① Possible cause: insufficient phosphomolybdic acid differentiation.

② Recommendation: extend phosphomolybdic acid differentiation time and monitor the endpoint microscopically.

(2) Muscle/cytoplasmic red staining too pale

① Possible cause: insufficient ponceau staining time or over-differentiation with phosphomolybdic acid.

② Recommendation: extend ponceau staining time or shorten phosphomolybdic acid differentiation.

(3) Gray background or dull color tones

① Possible cause: aged dye solutions, insufficient acetic acid rinsing, or non-standard dehydration/clearing.

② Recommendation: prepare fresh dye solutions, moderately extend running-water or acetic acid rinses as appropriate, and strictly control dehydration/clearing times.

(4) Nuclear staining too dark or too light

① Too dark: shorten hematoxylin staining time or increase the speed/intensity of differentiation.

② Too light: extend hematoxylin staining time or optimize the bluing step.

VII. Safety and Waste Disposal

7.1 Operational safety

(1) When handling acidic dyes, phosphomolybdic acid, glacial acetic acid, and organic solvents, wear gloves and protective eyewear and work in a well-ventilated area.

(2) Avoid contact of staining solutions and organic solvents with skin or mucosa; in case of contact, rinse immediately with copious running water.

7.2 Waste disposal

(1) Waste liquids containing dyes and strong acids must be collected and disposed of as hazardous chemical waste according to laboratory regulations; do not discharge into the sewage system.

(2) Used xylene and alcohol-containing organic waste should be collected separately and transferred to qualified waste handlers for proper recovery/disposal.

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