Histopathology and Cytopathology(Practical) part 3rd

THIS TOPIC IS FOR DIPLOMA IN MEDICAL LABORATORY

1. Histopathology and Cytopathology (Practical) Stain FNAC smears by Giemsa and Papanicolaou methods. Mount stained smears/section. Demonstrate Barr body by Aceto-Orcein staining method.

2. 11. Stain FNAC smears by Giemsa and Papanicolaou methods Stain FNAC smears by Giemsa methods Principle: Giemsa is a differential stain that contains a mixture of basic dyes ( methylene blue and azures) and an acidic dye (eosin).  Basic dyes:  Bind to acidic components of the cell, such as nuclear chromatin and RNA, producing a blue-purple color. Acidic dye:  Binds to basic components, like hemoglobin and cytoplasmic granules, yielding a pink-red color. Proper buffering of the stain is crucial for optimal results.  Required reagents and equipment : Reagents: Absolute methanol (high-grade, acetone-free) Giemsa stock solution Buffered distilled water, pH 6.8 or 7.2 Mounting medium (e.g., DPX, depending on intended storage) Xylene (if mounting) 

3. Equipment and materials: Clean, dry glass slides with FNAC smears Staining jars or racks Measuring cylinder or pipette Filter paper ( Whatman #1) Microscope Fume hood (for methanol and xylene )  Preparation of solutions: Working Giemsa solution: This must be prepared fresh immediately before use, as diluted Giemsa stain deteriorates quickly. Dilute the Giemsa stock solution with buffered distilled water. Slow method:  Mix 1 part Giemsa stock with 20–50 parts buffer (e.g., 1:50) for a 3% solution, and stain for 45–60 minutes. Rapid method:  Mix 1 part Giemsa stock with 9 parts buffer (1:10) for a 10% solution, and stain for 10–15 minutes. 

4. Staining procedure: Smear preparation and drying:  Prepare thin smears of the FNAC material on clean slides and allow them to air-dry completely. Do not apply heat, as this can damage cell morphology. Fixation:  Fix the air-dried slides by dipping them briefly in absolute methanol for 1 to 3 minutes. For very thick smears, air-dry for 24 hours without methanol fixation. Ensure the slides are completely dry again after fixation. Staining: For jar method:  Place the fixed slides in a Coplin jar containing the freshly prepared working Giemsa solution for the specified time (10–60 minutes, depending on the method). For rack method:  Place the slides face up on a staining rack and flood them with the working Giemsa solution.

5. Rinsing:  Gently rinse the slides with buffered distilled water (pH 7.2) to remove excess stain and prevent a metallic surface scum from forming. Drying:  Allow the slides to air-dry completely in a vertical position on a drying rack.  Examination and interpretation: Examine the smear under a microscope, first using a lower power objective to assess cell distribution and then using the oil immersion objective for detailed cellular morphology. Expected colors: Nuclei:  Dark blue to purple. Cytoplasm:  Pale blue or grey-blue. Erythrocytes (if present):  Pink. Lymphocyte cytoplasm:  Light blue. Granules:  Stained in various shades of red, orange, and blue, depending on the cell type. 

6. Stain FNAC smears by Papanicolaou methods A Papanicolaou (Pap) stain protocol for Fine Needle Aspiration Cytology (FNAC) is a multi-step procedure that delivers excellent nuclear and cytoplasmic detail. The following is a general guide; exact times may vary depending on the specific hematoxylin formulation and personal preference.  Equipment and reagents: Equipment: Coplin jars or staining rack Microscope slides Forceps  Reagents: Fixative:  95% Ethanol or commercial cytology spray fixative. Nuclear stain:  Harris' or Gill's Hematoxylin . Bluing agent:  Scott's Tap Water Substitute or weak lithium carbonate solution.

7. Counter stains: Orange G (OG-6). Eosin Azure (EA), such as EA-50 or EA-65. EA-50 is standard, while EA-65 is sometimes used for non-gynecological samples. Graded alcohols:  95% and 100% ethanol for hydration and dehydration. Clearing agent:   Xylene or xylene substitute. Mounting medium:   Permount or DPX.  Preparation and fixation: Immediately after the FNAC, create smears by expelling the aspirate onto a glass slide and spreading it with a second slide. Immediate fixation is critical  to prevent air-drying artifacts, which can compromise cell morphology and nuclear detail. Immediately place wet-fixed smears into a Coplin jar containing 95% ethanol for a minimum of 15 minutes. For smears fixed with a spray fixative, rinse off the fixative with 95% ethanol for 2 minutes. 

8. Staining: Hydrate the slides:  Immerse the fixed smears in 80% ethanol, 70% ethanol, and 50% ethanol for 1–2 minutes each. Finish with a rinse in distilled water for 1–2 minutes. Stain with hematoxylin :  Stain the nuclei by placing the slides in Harris' or Gill's Hematoxylin for 1.5 to 5 minutes, depending on the required intensity. Rinse and differentiate:  Rinse the slides in running tap water for 1–2 minutes. For a regressive stain using Harris' hematoxylin , briefly dip the slides in a 0.5% aqueous solution of hydrochloric acid to remove excess stain. Blue the nuclei:  Immerse the slides in Scott's Tap Water Substitute or a bluing reagent for 30–60 seconds until the nuclei turn blue. Rinse well in running tap water for 1 minute. Dehydrate gradually: Dip in 70% ethanol for 10 dips. Dip in 95% ethanol for 10 dips.

9. Stain with Orange G (OG-6):  Dip the slides in OG-6 for 1.5 to 2 minutes to stain keratinized cells orange. Rinse:  Rinse in two changes of 95% ethanol. Stain with Eosin Azure (EA):  Dip the slides in EA-50 or EA-65 for 2.5 to 5 minutes to stain cytoplasm and other components. Final dehydration: Rinse in two changes of 95% ethanol. Rinse in two changes of 100% ethanol for 1 minute each. Clear the slides:  Place the slides in two changes of xylene for 1–2 minutes each until clear.  Mounting: Apply a drop of permanent mounting medium, such as DPX, to the smear. Carefully place a coverslip over the smear, avoiding air bubbles. Allow the slides to dry before viewing under a microscope. 

10. Expected results: A properly executed Papanicolaou stain should yield a transparent, multi-colored smear with the following characteristics:  Nuclei:  Crisp and clear, with well-defined chromatin patterns, stained blue to black. Cytoplasm: Keratinized cells appear bright orange. Metabolically active cells (e.g., intermediate and parabasal cells) stain blue-green. Superficial epithelial cells and nucleoli stain pink. Background:  Clean and clear of cellular debris and excessive blood. 

11. 12. Mount stained smears/section Mounting a stained smear or section is the final step in preparing a microscopic specimen for long-term preservation and observation. The process involves applying a protective liquid medium and a glass coverslip to the specimen, which is already fixed and stained on a glass slide.  Principles of mounting: Preservation : A suitable mounting medium protects the tissue or cells from drying out, fading, and deterioration over time. Optical clarity : The medium fills the space between the specimen and the coverslip , preventing air bubbles that can obstruct the view. Mounting media have a refractive index close to glass, which minimizes light refraction and increases the visibility of the specimen. Firm sealing : The mounting medium solidifies to securely hold the coverslip in place and seal the sample. 

12. Method for mounting a stained tissue section: This method is typically used for histology samples that have been processed and stained.  Materials: Stained tissue section on a glass slide High-grade xylene (for clearing) Mounting medium (e.g., Canada balsam, DPX, or other resinous media) Glass coverslips Forceps Lint-free wipes or blotting paper  Procedure: Dehydrate the section : After staining, the slide is typically run through a series of increasing concentrations of alcohol (e.g., 70%, 95%, 100%) to remove all water.

13. Clear the section : Immerse the slide in xylene or a similar clearing agent. This makes the tissue transparent and ensures the mounting medium will mix correctly. Apply the mounting medium : Place a small drop of mounting medium over the stained tissue section on the slide. The exact amount depends on the size of the coverslip . Alternatively, place a drop of the mounting medium on a clean coverslip Lower the coverslip : Using forceps, hold the coverslip at a 45-degree angle with one edge touching the slide next to the mounting medium. Slowly lower the coverslip over the specimen to allow the medium to spread evenly and push air bubbles out. Remove excess medium : Gently press on the coverslip with a clean wipe or paper towel to remove any excess mounting medium that may seep out from the edges. Dry the slide : Place the slide flat to dry, preferably in a fume hood, until the mounting medium has hardened completely. This can take several hours or days, depending on the medium. 

14. Method for mounting a stained smear This method is used for thin preparations of cells, such as blood or bacterial smears.  Materials Stained and air-dried smear on a glass slide High-grade xylene (if using a resinous medium) Water-insoluble mounting agent (e.g., Permount or similar) Glass coverslips Forceps  Procedure: Prepare the smear : Ensure the smear has been properly air-dried and heat-fixed before staining. After staining and rinsing, blot the slide dry.

15. Mount the smear : For a permanent mount using a resinous medium, clear the completely dried slide by dipping it in xylene . Place a small drop of the mounting agent directly onto the smear. Lower a coverslip over the smear using the same technique as with tissue sections to avoid air bubbles. Dry the slide : Allow the slide to dry completely on a flat, level surface.  Common issues and tips Air bubbles : The most common problem. To prevent them, lower the coverslip slowly at a 45-degree angle. If bubbles form, they can sometimes be worked out by gently pressing on the coverslip before the medium hardens. Uneven smears : An unevenly prepared smear can result in an inconsistent final product. Practice the smearing technique for uniform cell distribution. Coverslip movement : Ensure the mounting medium is fully hardened before handling or storing the slides vertically, as the coverslip may shift. 

16. 13. Demonstrate Barr body by Aceto-Orcein staining method To demonstrate a Barr body using the aceto-orcein staining method, a buccal smear is prepared and stained with 1% aceto-orcein solution, which makes the Barr body appear as a darkly staining, small mass against the nucleus Step 1: Sample Collection: Ask the subject to rinse their mouth with water before collection. Use a wooden or sterile spatula to gently scrape the inside of the cheek to collect buccal cells. Smear the collected cells onto a clean glass slide. 

17. Step 2: Staining: Place a drop of 1% aceto-orcein solution onto the cells on the slide. Allow the cells to stain for about 20 minutes. This is a fresh, unstable stain and should be prepared just before use for best results Step 3: Mounting and Observation: Gently place a coverslip over the stained cells to avoid air bubbles. Dry the preparation to make it suitable for viewing. Examine the slide under an optical microscope, preferably using a higher magnification like oil immersion, to observe the Barr bodies. 

18. Interpretation In a female:  The nucleus will show one or more Barr bodies, which appear as darkly staining, small masses pressed against the nuclear membrane. In a male:  No Barr bodies will be visible, as only females typically have two X chromosomes, one of which becomes inactivated and condenses into a Barr body.