PERMANENT SLIDE PREPARATION

 

A permanent slide is a prepared microscope slide that can be stored for a long time without deteriorating. It is commonly used in biological and medical laboratories for the observation of specimens, such as cells, tissues, microorganisms, and other biological samples. The preparation of permanent slides involves the following basic steps:

Steps in Permanent Slide Preparation:

1. Fixation: Preserves the specimen structure and prevents decomposition.
2. Dehydration: Removes water from the specimen to prepare it for the mounting medium (especially if using organic solvents like Canada balsam or for specimens that will be mounted with resins).
3. Clearing: Makes the specimen transparent and prepares it for embedding in the mounting medium.
4. Staining: Adds color to specific parts of the specimen (e.g., cells, tissue, bacteria) to improve contrast for microscopic observation.
5. Mounting: Involves placing the specimen in a mounting medium to preserve and protect it long-term.
6. Sealing: Seals the specimen and mounting medium under a cover slip.

 

Materials Needed:

• Glass slide: Standard microscope slide.
• Cover slip: Thin glass cover (round, square, or rectangular) that protects the specimen and provides a flat surface for observation.
• Specimen: The biological sample to be observed (e.g., tissue, microorganisms, or cells).
• Fixative: Chemical solutions used to preserve the specimen, such as formaldehyde, ethanol, or acetic acid.
• Staining agents: Dyes used to color the specimen, such as Gram stain, Methylene blue, Iodine, Safranin, or Hematoxylin.
• Mounting medium: A liquid medium that helps to preserve the specimen and provide clarity for viewing (e.g., glycerin, Canada balsam, or DPX).
• Forceps: For handling small specimens and cover slips.
• Dissecting needle or brush: To handle small specimens, especially for bacterial slides.
• Distilled water: For rinsing specimens during preparation.

 

Steps for Preparing a Permanent Slide:

1. Collection and Preparation of Specimen

• Collection: Obtain a small sample of the specimen to be examined.
• Cutting (for tissue samples): If preparing a tissue sample, cut it into thin sections using a sharp scalpel or razor blade. The thickness of the sections depends on the specimen and the desired observation level (usually around 5–10 microns for tissues).
• Air-drying (for wet specimens): If the specimen is moist, allow it to air-dry for a few minutes.

 

2. Fixation

·        The fixation process preserves the specimen and maintains its structure by killing cells and microorganisms and preventing decomposition. A good fixative penetrates tissue quickly, reduces the shrinkage, and does not change the biological components of the tissue. The fixation of two types:

·        Physical fixation: by heating or cooling to the extent of freezing.

·        Chemical fixation: by using chemical fixative like formaldehyde in light microscope. In the electron microscope different fixative is used such as glutaraldehyde and potassium permanganate ( KMNO4).

Depending on the specimen, choose an appropriate fixative. For example:

• Formalin (10% formaldehyde) is commonly used for tissue specimens.
• Alcohol (e.g., ethanol) is used for microorganisms.
• Acetic acid is used for some types of plant material.
• Procedure:
• Place the specimen on the slide and add a few drops of the fixative.
• If using a tissue sample, immerse it in the fixative for 10–20 minutes. For microorganisms, a quick immersion (1–2 minutes) is usually sufficient.
• Rinse: After fixation, rinse the specimen with distilled water to remove excess fixative.

·        The goal of this process is to maintain the composition of cells and preventing the growth of bacteria in the sample and prevent decomposition also leads to hardening samples and make it possible to cut into thin slices.

3. Dehydration and Clearing

·        For long-term preservation, the slide must be properly dehydrated. The sample is exposed to an escalating series of alcohols like ethanol starting from 50%, 70%, 90%, 100% alcohol to gradually remove water from the specimen.  After dehydration, clearing agents like xylene or toluene are used to make the specimen transparent, allowing the mounting medium to flow easily and preserve clarity. Add a drop of xylene or toluene to the specimen and let it sit for a few minutes. Clearing is the process of making the sample or section clear and free of impurities and remove the dehydration solutions.

 

4. Staining

Staining enhances the contrast of the specimen, making it easier to observe under the microscope. Staining in the light microscope are by different dyes such as Eosin, Hematoxylin, Safranin, Toluidine blue while in Electron Microscope by uranyl acetate or lead citrate.

• Prepare Staining Solution: Depending on the specimen, use an appropriate dye or stain:
• For bacteria: Gram stain or Methylene blue.
• For plant cells: Iodine or Safranin.
• For animal tissues: Hematoxylin and Eosin (H&E).
• Staining Process:
• Place the slide with the specimen on a staining tray or dish.
• Add a few drops of the staining solution and let it sit for a period (usually 1-2 minutes for bacterial specimens, and up to 10 minutes for tissue specimens, depending on the stain).
• After staining, rinse the slide gently with distilled water to remove excess stain.
• Dehydration: If the specimen needs to be dehydrated (as with some histological preparations), pass it through a series of alcohol solutions (50%, 70%, 95%, 100%) to remove water from the sample. This step may be necessary for certain tissues, especially when using thicker stains like hematoxylin.

 

8- Mounting

Mounting involves placing the specimen in a mounting medium that will help preserve it and provide a medium that will support the cover slip. A drop of resinous substance such as Canada balsam, DPX, or PVA, is placed on the slide and covered with a glass cover and the slide is saved until the study.

 

• Common mounting media include:
• Glycerin: Used for temporary mounts or when long-term preservation is not necessary.
• Canada balsam: A commonly used resinous medium for permanent mounts. It’s clear, hardens over time, and provides excellent transparency.
• DPX (Distyrene Plasticizer Xylene) is another permanent mounting medium often used in histology.
• Place the Cover Slip: Gently lower a cover slip over the specimen using forceps. Be careful not to trap air bubbles between the slide and cover slip.
• Press Gently: Apply gentle pressure to the cover slip to spread the mounting medium evenly and expel excess fluid.

9. Sealing the Slide

To ensure the specimen remains securely mounted for long-term observation, the edges of the cover slip are sealed with a sealant.

• Sealant Application: After the cover slip is in place, apply a small bead of clear nail polish, paraffin wax, or Permount around the edges of the cover slip. This prevents the mounting medium from evaporating and helps to keep the cover slip in place.
• Allow the slide to dry for several hours to ensure that the sealant hardens.

11. Storage and Labeling

Once the slide is prepared and sealed, it can be stored for future use:

• Labeling: Write the name of the specimen, the preparation date, and any relevant details on the slide or slide holder (using a permanent marker or label).
• Storage: Store permanent slides in a cool, dry place. Ideally, slides should be kept in a slide box or slide rack to prevent contamination and damage.

For histological sections, embedding and sectioning can also be done.

Embedding

Paraffin wax is used to embed the samples to be examined by light microscope. After immersion in the wax, sample is poured in templates and left to cool and solidify.  In the electron microscope embedding material used must be more solid and more resistant to temperatures.

Sectioning

Sectioning is the process of getting thin slices of a certain thickness, using mechanical or manual devices called microtome. Microtome is a tool used to cut extremely thin slices of material, known as sections. Important in science, microtomes are used in microscopy, allowing for the preparation of sample for observation under transmitted light or electron radiation. Microtomes use steel, glass, or diamond blades depending upon the specimen being sliced and the desired thickness of the sections being cut. Steel blades are used to prepare sections of animal or plant tissues for light microscope histology, Glass knives are used to slice sections for light microscopy and to slice very thin sections for electron microscope. Industrial grade diamond knives are used to slice hard materials such as bone, teeth and plant matter for both light microscopy and for electron microscopy. Diamond knives are used for slicing thin sections for electron microscope.

 There are many types of microtome such as Sledge microtome for the preparation of large samples, such as those embedded in paraffin for biological preparations, Rotary microtome for hard materials, such as a sample embedded in a synthetic resin, Cryo-microtome for the cutting of frozen samples, Ultramicrotome can allow for the preparation of extremely thin sections, Vibrating microtome is usually used for difficult biological samples. Saw microtome is especially for hard materials such as teeth or bones. Laser microtome can also be used for very hard materials, such as bones or teeth as well as some ceramics.

Conclusion

Permanent slide preparation is a process that requires attention to detail to ensure the specimen is properly preserved, stained, and mounted for long-term observation. By following the proper steps of fixation, staining, dehydration, clearing, and mounting, the slide can be kept intact for years, making it useful for further study or reference. This process is widely used in microbiology, histology, and other biological fields.

 

 

Fungal Staining

Fungi are eukaryotic organisms with both macroscopic and microscopic characteristics.  Lactophenol Cotton Blue (LPCB) Staining is a simple histological staining method used for the microscopic examination and identification of fungi. The fungal spore cell wall is made up of chitin, which, the components of the Lactophenol Cotton Blue solution stains for identification. The lactophenol cotton blue solution acts as a mounting solution as well as a staining agent. The solution is blue in color and it is made up of a combination of three main reagents:

·        Phenol: It acts as a disinfectant by killing living organisms

·        Lactic acid: It preserves the fungal structures

·        Cotton blue: It stains the chitin on the fungal cell wall and other fungal structures

 

The stain will give a blue-colored appearance to the fungal spores and structures, such as hyphae.

 

Reagents of Lactophenol Cotton Blue (LPCB) Staining

 

A preparation of 50ml Lactophenol cotton Blue staining solution is made up of:

·        Distilled water 50ml

·        Cotton Blue (Aniline Blue) 0.125g

·        Phenol Crystals (C₆H₅O₄)  50g

·        Glycerol 100ml

·        Lactic acid (CH₃CHOH COOH) 50ml

·        70% ethanol

 

Preparation of Lactophenol Cotton Blue solution

Lactophenol Cotton Blue solution is prepared for over two days leaving the reagents undisturbed to allow dissolving and maturation.

1.     Day 1: Dissolve the cotton blue in distilled water and leave to rest overnight.

2.     Day2: Using protective gloves, add phenol crystals to lactic acid in a glass beaker and stir using a magnetic stirrer until the crystals dissolve.

3.     Add glycerol

4.     Filter the Cotton blue and the distilled water into the phenol + glycerol +lactic acid solution and mix.

5.     Store at room temperature.

 

Procedure of Lactophenol Cotton Blue (LPCB) Staining

1.     On a clean microscopic glass slide, add a drop of Lactophenol Cotton Blue Solution

2.     Add the fungal specimen to the drop of alcohol using a sterile mounter such as an inoculation needle.

3.     Tease the fungal sample using needle, to ensure the sample mixes well with the alcohol.

4.     Carefully cover the stain with a clean sterile coverslip without making air bubbles to the stain.

5.     Examine the stain microscopically at 40X, to observe for fungal spores and other fungal structures.

 

 

Results and Interpretation

Fungal spores, hyphae, and fruiting structures stain blue while the background stains pale blue.

 

          

                                     Penicillium                                 Aspergillus

Limitations

·        It can only be used as a presumptive identification method of fungi which should be followed up with other diagnostic tools such as biochemical and cultural examination.

·        The components of the solution should be used before expiry, including the use of the solution before it expires.

·        The solution may disrupt the original morphology of the fungi.

·        The stain can only be used to identify mature fungi and its structures and not the young vegetative forms of fungi.

·        The stain can not be stored for a long period of time.

 

Applications

Clinical diagnosis:

·        Identifying fungal infections in various body sites like skin, lungs, blood, and cerebrospinal fluid. 

·        Differentiating between different types of fungi based on their staining characteristics. 

·        Monitoring the effectiveness of antifungal treatments.

 

Research applications:

·        Studying fungal cell wall composition and structure 

·        Investigating fungal interactions with host tissues 

·        Analyzing fungal diversity in environmental samples 

·        Evaluating antifungal efficacy in vitro 

 

 

Common fungal stains and their characteristics:

 

·        Periodic Acid-Schiff (PAS):

Used for detection of fungi in tissue sections. Stains fungal cell walls pink, making it useful for identifying both yeast and hyphae. Oxidation of fungal polysaccharides results in the production of aldehydes that react with Schiff reagent and fungal cell walls appear pink/magenta in colour.

Procedure

1. Tissue sections are treated with periodic acid.
2. Stain with Schiff reagent.
3. Observe under bright-field microscope.

Results:

• Fungal structures appear magenta/red; tissue background lightly stained.

Applications:

• Detection of fungi in tissue sections (biopsy specimens).
• Identification of pathogenic fungi in histopathology.
• Examples: Candida albicans, Histoplasma capsulatum.

           

                                                           Candia hyphae overlying a     plaque of squamous cells- Fungal esophagitis- PAS stain

Calcofluor White:

• A fluorescent stain that binds to chitin and cellulose in fungal cell walls, allowing for rapid visualization with a fluorescence microscope. Fungi fluoresce bright blue-white under UV light.

Procedure

1. Place fungal material on a slide.
2. Add a drop of Calcofluor White.
3. Observe under fluorescence microscope.

Results:

• Fungal structures glow bright blue/white; background remains dark.

Applications:

• Rapid detection of fungi in clinical specimens (e.g., skin scrapings, sputum).
• Study of spores, hyphae, and yeast forms.
• Examples: Candida, Trichophyton.

 

Calcofluor white stained Candida albicans

showing true hyphae (*) and pseudohyphae (+).

Gomori's Methenamine Silver (GMS):

Gomori's Methenamine Silver (GMS), is a histological stain that highlights polysaccharides in tissue, primarily used to detect fungi by staining them black. It oxidises  carbohydrates in fungal cell walls to aldehydes, which then reduce silver ions to visible black metallic silver,  and is good for diagnosing fungal infections like Pneumocystis carinii pneumonia (PCP) and identifying opportunistic pathogens. GMS is considered highly sensitive for detecting fungi in tissue samples.   

 

GMS

Fungal staining methods with specific applications

Staining Method	Principle	Procedure (Brief)	Appearance / Results	Specific Applications / Examples
1. Lactophenol Cotton Blue (LPCB) Staining	Cotton Blue binds to chitin in fungal cell walls; phenol kills fungus, lactic acid preserves structure.	Mix fungal material with LCB, cover with coverslip, observe under bright-field microscope.	Hyphae, conidia, and spores appear blue; background pale.	- Identification of fungi in culture (e.g., Aspergillus, Penicillium). - Study morphology: hyphal septation, spore arrangement.
2. Calcofluor White Staining	Fluorescent dye binds to chitin and cellulose in fungal walls; fluoresces under UV light.	Mix fungal material with Calcofluor White, observe under fluorescence microscope.	Fungal structures bright blue/white; dark background.	- Rapid detection of fungi in clinical samples (e.g., skin scrapings, sputum). - Visualization of yeast and filamentous forms (e.g., Candida, Trichophyton).
3. Periodic Acid-Schiff (PAS) Staining	Periodic acid oxidizes polysaccharides to aldehydes, which react with Schiff reagent; fungal cell walls stain magenta.	Treat tissue section with periodic acid (Schiff reagent). Observe under bright-field microscope.	Fungal structures appear magenta/red; tissue lightly stained.	- Detection of fungi in tissue sections or biopsies (e.g., Candida albicans, Histoplasma capsulatum). - Diagnosis of invasive fungal infections.

In short

• LPCB → culture/fungi morphology
• Calcofluor White → rapid clinical detection, fluorescence
• PAS → tissue biopsy, invasive fungal infections