Histology Procedure

Introduction 

Histopathology (compound of three Greek words: ἱστός histos "tissue", πάθος pathos "disease-suffering", and -λογία -logia) refers to the microscopic examination of tissue in order to study the manifestations of disease. Specifically, in clinical medicine, histopathology refers to the examination of a biopsy or surgical specimen by a pathologist, after the specimen has been processed and histological sections have been placed onto glass slides. In contrast, cytopathology examines free cells or tissue fragments.

Histological techniques provide a visual means for the examination and analysis of cell/tissue physiology and morphology at the microscopic level.  Histology represents a broad technology, invaluable for studying and understanding the microscopic three-dimensional organization, structure, and function of cells and tissues and is especially useful for the diagnosis and understanding of disease at the cellular level. The process first involves isolation and fixation of the cells/tissue of interest.  Fixation preserves the structure and morphology of the specimen throughout the harsh conditions of dehydration, clearing, embedding, sectioning and staining.

Tissue ProcessingTissues from the body taken for diagnosis of disease processes must be processed in the histology laboratory to produce microscopic slides that are viewed under the microscope by pathologists. The techniques for processing the tissues, whether biopsies, larger specimens removed at surgery, or tissues from autopsy, are described below. The persons who do the tissue processing and make the glass microscopic slides are histotechnologists.

Specimen AccessioningTissue specimens received in the surgical pathology laboratory have a request form that lists the patient information and history along with a description of the site of origin. The specimens are accessioned by giving them a number that will identify each specimen for each patient.

Gross ExaminationTissues removed from the body for diagnosis arrive in the Pathology Department and are examined by a pathologist, pathology assistant, or pathology resident. Gross examination consists of describing the specimen and placing all or parts of it into a small plastic cassette which holds the tissue while it is being processed to a paraffin block. Initially, the cassettes are placed into a fixative.

When a malignancy is suspected, then the specimen is often covered with ink in order to mark the margins of the specimen. Different colored inks can be used to identify different areas if needed. When sections are made and processed, the ink will mark the actual margin on the slide.

Gross specimen examination1

PREPARATION OF HISTOLOGICAL SPECIMENS

Preparation of histological specimens undergo the following steps:

1. fixation .

2. tissue processing :

a. dehydration .

b. clearing .

c. impregnation .

3. embedding .

4. trimming .

5. cutting .

6. staining .

(1) TISSUE FIXATION:

IntroductionOnce tissues are removed from the body, they undergo a process of self-destruction or autolysis, which is initiated soon after cell death by the action of intracellular enzymes causing the breakdown of protein and eventual liquefaction of the cell.

Autolysis is more severe in tissues which are rich in enzymes, such as the liver, brain and kidney, and is less rapid in tissues such as elastic fiber and collagen.The objective of fixation is to preserve cells and tissue constituents in as close a life-like state as possible and to allow them to undergo further preparative procedures without change. Fixation arrests autolysis and bacterial decomposition and stabilizes the cellular and tissue constituents so that they withstand the subsequent stages of tissue processing.

Fixation should also provide for the preservation of tissue substances and proteins. Fixation is, therefore, the first step and the foundation in a sequence of events that culminates in the final examination of a tissue section.

It is relevant to point out that fixation in itself constitutes a major artefact. The living cell is fluid or in a semi-fluid state, whereas fixation produces coagulation of tissue proteins and constituents, a necessary event to prevent their loss or diffusion during tissue processing; the passage through hypertonic and hypotonic solutions during tissue processing would otherwise disrupt the cells. For example, if fresh unfixed tissues were washed for prolonged periods in running water, severe and irreparable damage and cell lysis would result. In contrast, if the tissues were first fixed in formalin, subsequent immersion in water is generally harmless.

Tissue fixation The technique of using fixatives in the preparation of cytologic, histologic, or pathologic specimens for the purpose of maintaining the existing form and structure of all the constituent elements.

Fixatives

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Agents employed in the preparation of histologic or pathologic specimens for the purpose of maintaining the existing form and structure of all of the constituent elements. Great numbers of different agents are used; some are also decalcifying and hardening agents. They must quickly kill and coagulate living tissue. A large variety of fixatives is now available. Each fixative has advantages and disadvantages, some are restrictive while others are multipurpose.

Fixation - types of fixativesThe purpose of fixation is to preserve tissues permanently in as life-like a state as possible. Fixation should be carried out as soon as possible after removal of the tissues (in the case of surgical pathology) or soon after death (with autopsy) to prevent autolysis. There is no perfect fixative, though formaldehyde comes the closest. Therefore, a variety of fixatives are available for use, depending on the type of tissue present and features to be demonstrated.

There are five major groups of fixatives, classified according to mechanism of action:

Aldehydes Mercurials Alcohols Oxidizing agents Picrates

Over the years, various classifications of fixatives have been proposed, with major divisions according to function as coagulants and non-coagulants, or according to their chemical nature into three general categories which include alcoholic, aldehydic and heavy metal fixatives.

Factors involved in fixation Temperature Size of specimens and penetration of fixative Changes in volume pH and buffers Osmolality Concentration of fixatives Duration of fixation

Formulations for various fixativesThe details provided relate to commonly used fixatives. Many variations are available and more specialised fixative solutions are not provided.

1) Formaldehyde solutions

10% neutral buffer formalin (4% formaldehyde)

Reagents required 40% formaldehyde 100 ml Distilled water 900 ml Sodium dihydrogen orthophosphate 4 g Disodium hydrogen orthophosphate (anhydrous) 6.5 g

MethodPrepare, using quantities indicated. Fixation time: 24-72 hours.

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2) Baker's formol-calcium (modified)

Reagents required 40% formaldehyde 100 ml Distilled water 900 ml 10% calcium chloride 100 ml 7 g of cadmium chloride is sometimes added to the mixture

MethodPrepare, using quantities indicated. Fixation time: 16-24 hours.

3) Formol saline

Reagents required1 40% formaldehyde 100 ml2 Sodium chloride 9 g3 Tap water 900 ml

MethodPrepare, using quantities indicated.

4) Alcoholic formaldehyde

Reagents required1 40% formaldehyde 100 ml2 95% alcohol 900 ml3 0.5 g calcium acetate may be added to this mixture to ensure neutrality

MethodPrepare, using quantities indicated. Fixation time: 16-24 hours.

Paraformaldehyde

Reagents required Solution A26% sodium dihydrogen orthophosphate 41.5 ml52% sodium hydroxide 8.5 mlHeat to 60°C-80°C in a covered containerParaformaldehyde 2 g

MethodAdd paraformaldehyde to solution A, stirring until the mixture is clear. Filter and cool. Adjust pH to 7.2 - 7.4. Prepare fresh for use (duration of fixation depends on size of specimen and whether for light or electron microscopy).

Buffered formaldehyde-glutaraldehyde 200 mOsm38

REAGENTS REQUIRED1 Sodium dihydrogen orthophosphate 1.6 g2 Sodium hydroxide 0.27 g

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3 Distilled water 88 ml4 40% formaldehyde 10 ml5 50% glutaraldehyde 2 ml

METHODPrepare, using quantities indicated. Fixation time: 16-24 hours.

Alcoholic fixativesCarnoy's fixativeREAGENTS REQUIRED1 Absolute ethanol 60 ml2 Chloroform 30 ml3 Glacial acetic acid 10 ml

METHODPrepare, using quantities indicated. Fixation time: 1-5 minutes.

MethacarnREAGENTS REQUIRED1 Absolute methanol 60 ml2 Chloroform 30 ml3 Glacial acetic acid 10 ml

METHODPrepare, using quantities indicated. Fixation time: 5-6 hours.

Wolman's solutionREAGENTS REQUIRED1 Absolute ethanol 95 ml2 Glacial acetic acid 5 ml

METHODImmerse frozen section in solution and microwave at 650 watts for 15 seconds.

Acetic alcohol formalinREAGENTS REQUIRED1 40% formaldehyde 10 ml2 Acetic acid 5 ml3 Ethanol 85 ml

METHODPrepare, using quantities indicated. Fixation time: 24 hours at 4°C.

Picric acid fixativesRossman's fluidREAGENTS REQUIRED1 100% ethanol saturated with picric acid 90 ml2 Neutralised commercial formalin 10 ml

METHODPrepare, using quantities indicated. Fix for 12-24 hours and wash very well in 95% ethanol.

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Gendre's fluidREAGENTS REQUIRED1 90% ethanol saturated with picric acid 80 ml2 40% formaldehyde 15 ml3 Glacial acetic acid 5 ml

METHODPrepare, using quantities indicated. Fixation is normally for 4 hours, followed by washing in 80%, 95% and 100% ethanol.

Bouin's fluidREAGENTS REQUIRED1 Saturated aqueous picric acid solution 75 ml2 40% formaldehyde 25 ml3 Glacial acetic acid 5 ml

METHOD1 Prepare, using quantities indicated. Fixation may vary from a few hours to 18 hours.2 Washing with 70% ethanol after fixation will remove most of the yellow colour. Sections can also be washed after removal of paraffin wax.

MERCURIC FIXATIVES

1) Buffered formaldehyde sublimate

Reagents required1 Mercuric chloride 6 g2 Distilled water 90 ml3 Sodium acetate 1.25 g4 40% formaldehyde 10 ml

MethodPrepare, using quantities indicated. Fixation time: 16-18 hours.

2) Zenker's fluid

Reagents required1 Distilled water 950 ml2 Potassium dichromate 25 g3 Mercuric chloride 50 g4 Glacial acetic acid 50 g

MethodPrepare, using quantities indicated. Fixation is normally for 4-24 hours followed by an overnight wash.

3) Helly's fluid

Reagents required1) Solution ADistilled water 1000 mlPotassium dichromate 25 g

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Sodium sulphate 10 gMercuric chloride 50 g

2) Solution B40% formaldehyde 50 ml

MethodAdd solution A to solution B immediately before use.

4) B5 fixative

Reagents required1) Stock reagent AMercuric chloride 60 gSodium acetate 12.5 gDistilled water l

2) Stock reagent B10% buffered neutral formalin

MethodTo prepare a working solution mix 90 ml stock reagent A with 10 ml stock reagent B. Fixation time: 5-8 hours.

5) Susa fluid

Reagents required Distilled water 80 ml 40% formaldehyde 20 ml Glacial acetic acid 4 ml Trichloroacetic acid 2 g Mercuric chloride 4.5 g Sodium chloride 0.5 g

MethodPrepare, using quantities indicated. Fixation time: 12 hours.

(2)TISSUE PROCESSING :

Principles of tissue processingthe aim of tissue processing is to embed the tissue in a solid medium firm enough to support the tissue and give it sufficient rigidity to enable thin sections to be cut , and yet soft enough not to damage the knife or tissue .

The most satisfactory embedding material for routine histology is paraffin wax. Most fixatives are aqueous-based and these are not miscible with paraffin wax to enable impregnation with this medium, the tissue must be processed. Each stage must be of sufficient length to ensure completeness.

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The stages involved are:

Dehydration: to remove fixative and water from the tissue and replace them with dehydrating fluid. Clearing: replacing the dehydrating fluid with a fluid that is totally miscible with both the

dehydrating fluid and the embedding medium.

Impregnation: replacing the clearing agent with the embedding medium.

Embedding

DehydrationThe first step in processing is dehydration. Water is present in tissues in free and bound (molecular) forms. Tissues are processed to the embedding medium by removing some or all of the free water. During this procedure various cellular components are dissolved by dehydrating fluids. For example, certain lipids are extracted by anhydrous alcohols, and water soluble proteins are dissolved in the lower aqueous alcohols

To minimize tissue distortion from diffusion currents, delicate specimens are dehydrated in a graded ethanol series from water through 10%-20%-50%-95%-100% ethanol.

In the paraffin wax method, following any necessary post fixation treatment, dehydration from aqueous fixatives is usually initiated in 60%-70% ethanol, progressing through 90%-95% ethanol, then two or three changes of absolute ethanol before proceeding to the clearing stage.

Duration of dehydration should be kept to the minimum consistent with the tissues being processed. Tissue blocks 1 mm thick should receive up to 30 minutes in each alcohol, blocks 5 mm thick require up to 90 minutes or longer in each change. Tissues may be held and stored indefinitely in 70% ethanol without harm.

Dehydrating agents

Alcohols Ethanol is probably the most commonly used dehydrant in histology. Ethanol is a rapid, efficient

and widely applicable dehydrant. Methanol is a good ethanol substitute but rarely used for routine processing because of its

volatility, flammability and cost.

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Isopropanol was first suggested as an ethanol substitute during the prohibition era in the United States.

Glycol-ethersUnlike the alcohols, these reagents do not act as secondary fixatives, and apart from solvent effects do not appear to alter tissue reactivity.

Ethoxyethanol Dioxane

Polyethylene glycols

Other dehydrants

Acetone Tetrahydrofuran

Phenol

ClearingClearing is the transition step between dehydration and infiltration with the embedding medium. Many dehydrants are immiscible with paraffin wax, and a solvent (transition solvent, ante medium, or clearant) miscible with both the dehydrant and the embedding medium is used to facilitate the transition between dehydration and infiltration steps. Shrinkage occurs when tissues are transferred from the dehydrant to the transition solvent, and from transition solvent to wax

Choice of a clearing agent depends upon the following:

The type of tissues to be processed, and the type of processing to be undertaken The processor system to be used Intended processing conditions such as temperature, vacuum and pressure Safety factors Cost and convenience. Speedy removal of dehydrating agent. Ease of removal by molten paraffin wax. Minimal tissue damage.

Transition solvents

HydrocarbonsToluene and xylene clear rapidly and tissues are rendered transparent, facilitating clearing endpoint determination.

Petroleum solvents Chlorinated hydrocarbons

Chloroform

Carbon tetrachloride

Trichloroethane

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EstersThese are colourless flammable solvents miscible with most organic solvents and with paraffin wax.

n-Butyl acetate Amyl acetate, methyl benzoate and methyl salicylate

TerpenesTerpenes are isoprene polymers found in essential oils originally derived from plants,77 though some are now synthesised.

Cedarwood oil Limonene

Terpineol

Infiltration

It is the saturation of tissue cavities and cells by a supporting substance which is generally, but not always, the medium in which they are finally embedded. Tissues are infiltrated by immersion in a substance such as a wax, which is fluid when hot and solid when cold. Alternatively, tissues can be infiltrated with a solution of a substance dissolved in a solvent, for example nitrocellulose in alcohol-ether, which solidifies on evaporation of the solvent to provide a firm mass suitable for sectioning.

Parffin waxProperties of paraffin wax are as follows:

Paraffin wax is a polycrystalline mixture of solid hydrocarbons produced during the refining of coal and mineral oils. It is about two thirds the density and slightly more elastic than dried protein

The properties of paraffin wax are improved for histological purposes by the inclusion of substances added alone or in combination to the wax:

improve ribboning increase hardness decrease melting point improve adhesion between specimen and wax

Embedding

Embedding is the process by which tissues are surrounded by a medium such as agar, gelatine, or wax which when solidified will provide sufficient external support during sectioning.

Embedding tissues in paraffin waxTissues are embedded by placing them in a

mould filled with molten embedding medium

which is then allowed to solidify. Embedding requirements and procedures are essentially the same for all waxes, and only the technique for paraffin wax is provided here in detail. At the completion of processing, tissues are held in clean paraffin wax which is free of solvent and particulate matter.

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Tissue processingA modular tissue embedding centre

There are four main mould systems and associated embedding protocols presently in use :

traditional methods using paper boats; Leuckart or Dimmock embedding irons or metal containers; the Peel-a-way system using disposable plastic moulds and; systems using embedding rings or cassette-bases which become an integral part of the block and serve as the block holder in the microtome.

Tissue processingEmbedding moulds:

(A) paper boat;

(B) metal bot mould;

(C) Dimmock embedding mould;

(D) Peel-a-way disposable mould;

(E) base mould used with embedding ring

(F) or cassette bases (G)

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General Embedding ProcedureMETHOD1 Open the tissue cassette, check against worksheet entry to ensure the correct number of tissue pieces are present.2 Select the mould, there should be sufficient room for the tissue with allowance for at least a 2 mm surrounding margin of wax.3 Fill the mould with paraffin wax.4 Using warm forceps select the tissue, taking care that it does not cool in the air; at the same time.5 Chill the mould on the cold plate, orienting the tissue and firming it into the wax with warmed forceps. This ensures that the correct orientation is maintained and the tissue surface to be sectioned is kept flat.6 Insert the identifying label or place the labelled embedding ring or cassette base onto the mould.7 Cool the block on the cold plate, or carefully submerge it under water when a thin skin has formed over the wax surface.8 Remove the block from the mould.9 Cross check block, label and worksheet.

ORIENTATION OF TISSUE IN THE BLOCKCorrect orientation of tissue in a mould is the most important step in embedding. Incorrect placement of tissues may result in diagnostically important tissue elements being missed or damaged during microtomy. In circumstances where precise orientation is essential tissue should be marked or agar double embedded. Usually tissues are embedded with the surface to be cut facing down in the mould. Some general considerations are as follows:

elongate tissues are placed diagonally across the block tubular and walled specimens such as vas deferens, cysts and gastrointestinal tissues are embedded

so as to provide transverse sections showing all tissue layers tissues with an epithelial surface such as skin, are embedded to provide sections in a plane at right

angles to the surface (hairy or keratinised epithelia are oriented to face the knife diagonally) multiple tissue pieces are aligned across the long axis of the mould, and not placed at random.

Tissue orientation in the block.

(A) elongate tissues;

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(B) tubular or cystic specimens;

(C) hairy skin;

(D) multiple tissue fragments.

The knife is at the lower block margin.

Processing methods and routine schedulesTissues are usually more rapidly processed by machine than by manual methods, although the latter can be accelerated by using microwave or ultrasonic stimulation. For routine purposes tissues are most conveniently processed through dehydration, clearing and infiltration stages automatically by machine. There are two broad types of automatic tissue processors - tissue-transfer and fluid-transfer types.

Automated tissue processingTissue-transfer processors : These processors are characterised by the transfer of tissues, contained within a basket, through a series of stationary reagents arranged in-line or in a circular carousel plan. The rotary or carousel is the most common model of automatic tissue processor, and was invented by Arendt in 1909.79 It is provided with 9-10 reagent and 2-3 wax positions, with a capacity of 30-110 cassettes depending upon the model. Fluid agitation is achieved by vertical oscillation or rotary motion of the tissue basket. Processing schedules are card-notched, pin or touch pad programmed.

Processing schedules for a tissue-transfer processor. Day schedule for urgent specimens, tissues 2 mm, fixed in Carnoy's fluid. Overnight schedule for routine processing. Tissue blocks 2-3 mm, single load. For a double load, immersion times should be equal. Weekend processing: tissues are held in fixative, or preferably 70% ethanol until Sunday.

STEP TEMPERETURE TIME

Fixative(10%buffered formalin)

40 120 minutes

Fixative(10%buffered formalin)

40 120 minutes

70% alcohol(ethanol) 40 60 minutes

95% alcohol (ethanol) 40 60 minutes

Absolute alcohol 40 60 minutes

Absolute alcohol 40 60 minutes

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Absolute alcohol 40 60 minutes

Xylene 40 60 minutes

Xylene 40 60 minutes

paraffin wax 60 60 minutes

paraffin wax 60 60 minutes

paraffin wax 60 60 minutes

Embed

TOTAL TIME 14 hours

Tissue-transfer processors allow maximum flexibility in the choice of reagents and schedules that can be run on them, in particular, metal-corrosive fixatives, a wide range of solvents, and relatively viscous nitrocellulose solutions can all be accommodated. These machines have a rapid turn-around time for day/night processing. In more recent models the tissue basket is enclosed within an integrated fume hood during agitation and transfer cycles thus overcoming the disadvantages of earlier styles.

A tissue-transfer tissue processor with an integrated fume hood (Shandon Citadel). Tissue cassettes are loaded into the basket on the rotating head, which transfers tissues around the series of reagent containers

AUTOMATIC TISSUE PROCESSOR

Features:

1) Glass beakers / aluminum reagent containers

2) 2 wax baths Standard tissue basket with

capacity of 80 cassettes

3) Possibility of interrupting an automatic

process for reloading or removing cassettes for

special applications before the end of a run

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GENERAL CONSIDERATIONSBaskets and metal cassettes should be clean and wax-free.Tissues should not be packed too tightly in baskets so as to impede fluid exchange.Processors must be free of spilt fluids and wax accumulations to reduce hazards and to ensure mechanical reliability.Fluid levels must be higher than the specimen containers.Timing and delay mechanism must be correctly set and checked against the appropriate processing schedule.A processor log should be kept in which the number of specimens processed, processing reagent changes, temperature checks on the wax baths and the completion of the routine maintenance schedule, is recorded as an integral part of the laboratory quality assurance program.

Frozen Sections

At times during performance of surgical procedures, it is necessary to get a rapid diagnosis of a pathologic process. The surgeon may want to know if the margins of his resection for a malignant neoplasm are clear before closing, or an unexpected disease process may be found and require diagnosis to decide what to do next, or it may be necessary to determine if the appropriate tissue has been obtained for further workup of a disease process. This is accomplished through use of a frozen section. The piece(s) of tissue to be studied are snap frozen in a cold liquid or cold environment (-20 to -70 Celsius). Freezing makes the tissue solid enough to section with a microtome.

Frozen sections are performed with an instrument called a cryostat. The cryostat is just a refrigerated box containing a microtome. The temperature inside the cryostat is about -20 to -30 Celsius. The tissue sections are cut and picked up on a glass slide. The sections are then ready for staining.

Cutting a frozen section

(3)TISSUE-TRIMMING ( border molding )

The shaping of an impression material by the manipulation or action of the tissues adjacent to the borders of an impression.

(4) CUTTING :

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Sectioning with microtome

(6) STAINING :

H&E  and PAS are the two most commonly used stains in histopathology.

Haematoxylin and eosin (H & E)

This is the most commonly used stain in routine pathology. Haematoxylin, a basic dye stains acidic structures a purplish blue. Nuclei (DNA), ribosomes and rough endoplasmic reticulum (with their RNA) are therefore stained blue with H&E. Eosin, in contrast is an acidic dye which stains basic structures red or pink. Most cytoplasmic proteins are basic and therefore stained pink or pinkish red. In summary, H&E stains nuclei blue and cytoplasm pink or red. The following specimen shows a normal skin stained with H&E.

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PAS (periodic acid-Schiff)

This stain is versatile and has been used to stain many structures including glycogen, mucin, mucoprotein, glycoprotein, as well as fungi. PAS is useful for outlining tissue structures--basement membranes, capsules, blood vessels, etc. As it stains many structure; this can give rise to a high background. It is very sensitive, but specificity depends upon interpretation. The following specimen is a normal cornea that has been stained with PAS and the basement membrane of the epithelium is highlighted.

Special Stains in Histology

Mucin stains

There are a variety of mucin stains, all attempting to demonstrate one or more types of mucopolysaccharide substances in tissues. The types of mucopolysaccharides are as follows:

Neutral - These can be found in glands of the GI tract and in prostate. They stain with PAS but not with Alcian blue, colloidal iron, mucicarmine, or metachromatic dyes.

Acid (simple, or non-sulfated) - Are the typical mucins of epithelial cells containing sialic acid. They stain with PAS, Alcin blue at pH 2.5, colloidal iron, and metachromatic dyes. They resist hyaluronidase digestion.

Acid (simple, mesenchymal) - These contain hyaluronic acid and are found in tissue stroma. They do not stain with PAS, but do stain with Alcian blue at pH 2.5, colloidal iron, and metachromatic dyes. They digest with hyaluronic acid. They can be found in sarcomas.

Acid (complex, or sulfated, epithelial) - These are found in adenocarcinomas. PAS is usually positive. Alcian blue is positive at pH 1, and colloidal iron, mucicarmine, and metachromatic stains are also positive. They resist digestion with hyaluronidase.

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Acid (complex, connective tissue) - Found in tissue stroma, cartilage, and bone and include substances such as chondroitin sulfate or keratan sulfate. They are PAS negative but do stain selectively with Alcian blue at pH 0.5.

There are a variety of stains for mucin:

Colloidal iron ("AMP") - Iron particles are stabilized in ammonia and glycerin and are attracted to acid mucopolysaccharides. It requires formalin fixation. Phospholipids and free nucleic acids may also stain. The actual blue color comes from a Prussian blue reaction. Tissue can be pre-digested with hyaluronidase to provide more specificity.

Alcian blue - The pH of this stain can be adjusted to give more specificity. PAS (peroidic acid-Schiff) - Stains glycogen as well as mucins, but tissue can be pre-digested with

diastase to remove glycogen. Mucicarmine - Very specific for epithelial mucins.

The mucin stain with the most specificity is mucicarmine, but it is very insensitive, so it is not really very useful. The stain that is the most sensitive is PAS, but you must learn how to interpret it in order to gain specificity. Colloidal iron stains are unpredictable. Alcian blue stains are simple, but have a lot of background staining.

Stains for biogenic amines

Cells that produce polypeptide hormones, active amines, or amine precursors (epinephrine, norepinephrine) can be found individually (Kulchitsky cell of GI tract) or as a group (adrenal medulla). The following is a traditional classification of the staining patterns based upon the ability of the cells to reduce ammoniacal silver nitrate to metallic silver (black deposit in tissue section):

Chromaffin Argentaffin Argyrophil (pre-reduction step necessary)

The distinction between chromaffin and argentaffin is artificial, since this depends upon the fixative used. "Chromaffin" cells have cytoplasmic granules that appear brown when fixed with a dichromate solution. "Argentaffin" cells reduce a silver solution to metallic silver after formalin fixation. Either reaction can be produced depending upon which fixative was used. Traditionally, chromaffin reaction is associated with adrenal medulla or extraadrenal paraganglion tissues (pheochromocytomas) whereas argentaffin reaction is associated with carcinoid tumors of the gut. Using a pre-reduction step may get more cells to stain, but they are called "argyrophil" then.

Types of stains for argentaffin include:

1. Diazo (diazonium salts)2. Fontana-Masson3. Schmorl's4. Autofluorescence

Types of stains for chromaffin include:

1. Modified Giemsa2. Schmorl's3. Wiesel's

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Types of stains for argyrophil include:

1. Grimelius (Bouin's fixative preferred)2. Pascual's

Melanin stains

Melanin is normally found in the skin, eye, and substantia nigra. It may also be found in melanomas.

The commonly used Fontana-Masson ("melanin stain") method relies upon the melanin granules to reduce ammoniacal silver nitrate (but argentaffin, chromaffin, and some lipochrome pigments also will stain black as well).

Schmorl's method uses the reducing properties of melanin to stain granules blue-green.

The most specific method of all is an enzyme histochemical method called DOPA-oxidase. It requires frozen sections for best results, but paraffin sections of well-fixed tissues may be used. The stain works because the DOPA substrate is acted upon by DOPA-oxidase in the melanin-producing cells to produce a brownish black deposit.

Bleaching techniques remove melanin in order to get a good look at cellular morphology. They make use of a strong oxidizing agent such as potassium permanganate or hydrogen peroxide. Ocular melanin takes hours to bleach, while that from skin takes minutes.

Formaldehyde-induced fluorescence can be used to highlight biogenic amines (chromaffin, argentaffin) and melanin in tissues. Formalin fixation imparts a strong yellow autofluorescence to unstained tissues with these substances.

The pseudomelanin of melanosis coli is PAS positive whereas true melanin is not. Moreover, pseudomelanin pigment is usually found in macrophages.

Lipochrome (lipofuschin) pigments

These are the breakdown products within cells from oxidation of lipids and lipoproteins. They are the wear-and-tear pigments found most commonly in heart, liver, CNS, and adrenal cortex (zona reticularis). The less highly oxidized "ceroid" pigment of testis interstitium and seminal vesicle is another form of lipochrome.

Lipochrome can be stained by Sudan black B, long Ziehl-Neelson acid fast, and Schmorl's methods. Lipochrome may also exihibit a strong orange autofluorescence in formalin-fixed, unstained paraffin sections.

Iron (hemosiderin)

Hemosiderin (storage iron granules) may be present in areas of old hemorrhage or be deposited in tissues with iron overload (hemosiderosis is the term used if the iron does not interfere with organ function; hemochromatosis refers to a condition of iron overload associated with organ failure).

Perl's iron stain is the classic method for demonstrating iron in tissues. The section is treated with dilute hydrochloric acid to release ferric ions from binding proteins. These ions then react with potassium

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ferrocyanide to produce an insoluble blue compound (the Prussian blue reaction). Mercurial fixatives seem to do a better job of preserving iron in bone marrow than formalin.

Calcium

Only calcium that is bound to an anion (such as PO4 or CO3) can be demonstrated. Calcium forms a blue-black lake with hematoxylin to give a blue color on H&E stain, usually with sharp edges.

VonKossa stain is a silver reduction method that demonstrates phosphates and carbonates, but these are usually present along with calcium. This stain is most useful when large amounts are present, as in bone.

Alizarin red S forms an orange-red lake with calcium at a pH of 4.2. It works best with small amounts of calcium (such as in Michaelis-Gutman bodies). The alizarin method is also used on the Dupont ACA analyzer to measure serum calcium photometrically.

Azan stain can be used to differentiate osteoid from mineralized bone.

Urates

Uric acid crystals are seen in acid urine. In tissue, urates are present as sodium urate. They are soluble in aqueous solutions and slightly soluble in weak alcoholic solutions. Therefore, tissues must be fixed in 95% or absolute alcohol to prevent leaching of urates.

Methenamine silver stains urates black. Sodium urate crystals are also birefringent on polarization. Using a red plate, the crystals show negative birefringence (yellow color) when the crystal's long axis is aligned in the direction of the slow wave. At 90 degrees to this, the crystals will be blue.

Exogenous pigments and minerals

These come from industrial or environmental exposure by inhalation, ingestion, or contact. Sometimes exposure comes from work-related activities (miners). Sometimes they are planned (tattoo).

Carbon appears as anthracotic pigment in the lungs. It can be distinguished from melanin by doing a melanin bleach. Poorly fixed tissues may contain formalin-heme pigment, which is black and finely granular, but this is widely scattered in the tissues without regard to cellular detail. Formalin-heme pigment is also birefringent on polarization.

Asbestos is a special type of long-thin silica crystal, usually of the mineral group chrysotile. In tissue, these crystals are highly irritative and highly fibrogenic. The fibers become coated with a protein-iron-calcium matrix, giving them a shish-kebab appearance. These are called "ferruginous bodies" because they are highlighted with an iron stain.

Silica is present in many minerals and building materials. Most forms are very inert and cannot be stained in tissue but can be demonstrated by white birefringence on polarization. It is most often present in lung, but can make its way into lymph node.

Street drugs for injection often are diluted with compounds containing minerals such as silica or talc. These crystals can be found throughout the body, but especially in lymphoreticular tissues.

Tattoo pigment is usually black and is inert and non-polarizable. Red tattoo pigment often contains cinnabar (which has mercury in it).

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In general, minerals are best demonstrated by microincineration techniques or by scanning electron microscopy with energy dispersive analysis (SEM-EDA).

Another use for SEM-EDA in forensic pathology is for analysis of gunshot residue. The primer residue has a characteristic pattern because of the elemental composition which contains antimony, barium, and lead.

Fat stains

The oil red O (ORO) stain can identify neutral lipids and fatty acids in smears and tissues. Fresh smears or cryostat sections of tissue are necessary because fixatives containing alcohols, or routine tissue processing with clearing, will remove lipids. The ORO is a rapid and simple stain. It can be useful in identifying fat emboli in lung tissue or clot sections of peripheral blood.

Connective tissue stains

The trichrome stain helps to highlight the supporting collagenous stroma in sections from a variety of organs. This helps to determine the pattern of tissue injury. Trichrome will also aid in identifying normal structures, such as connective tissue capsules of organs, the lamina propria of gastrointestinal tract, and the bronchovascular structures in lung.

The reticulin stain is useful in parenchymal organs such as liver and spleen to outline the architecture. Delicate reticular fibers, which are argyrophilic, can be seen. A reticulin stain occasionally helps to highlight the growth pattern of neoplasms.

An elastic tissue stain helps to outline arteries, because the elastic lamina of muscular arteries, and the media of the aorta, contain elastic fibers. The van Gieson method for elastic fibers provides good contrast.

Giemsa stain

There are a variety of "Romanowsky-type" stains with mixtures of methylene blue, azure, and eosin compounds. Among these are the giemsa stain and the Wright's stain (or Wright-Giemsa stain). The latter is utilized to stain peripheral blood smears. The giemsa stain can be helpful for identifying components in a variety of tissues.

One property of methylene blue and toluidine blue dyes is metachromasia. This means that a tissue component stains a different color than the dye itself. For example, mast cell graules, cartilage, mucin, and amyloid will stain purple and not blue, which is helpful in identifying these components.

Microorganisms

Bacteria appear on H and E as blue rods or cocci regardless of gram reaction. Colonies appear as fuzzy blue clusters. Tissue gram stains are all basically the same as that used in the microbiology lab except that neutral red is used instead of safranin. Gram positive organisms usually stain well, but gram negatives do not (because the lipid of the bacterial walls is removed in tissue processing). Brown and Brenn (or the Brown and Hopps modification) is the method most commonly used.

Fungi stain blue with H and E and red with PAS. The most sensitive method for demonstrating them is Methenamine silver.

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Spirochetes are very difficult to stain. The best method is the Warthin-Starry. A Giemsa stain may help demonstrate donovan bodies and leishmania.

AFB (acid fast bacilli) stain

This stain uses carbol-fuchsin to stain the lipid walls of acid fast organisms such as M. tuberculosis. The most commonly used method is the Ziehl-Neelsen method, though there is also a Kinyoun's method. A modification of this stain is known as the Fite stain and has a weaker acid for supposedly more delicate M. leprae bacilli. However, much of the lipid in mycobacteria is removed by tissue processing, so this stain can, at times, be very frustrating and cause you to search extensively for organsisms you are sure are in a big granuloma. The most sensitive stain for mycobacteria is the auramine stain which requires a fluorescence microscope for viewing.

There are things other than mycobacteria that are acid fast. Included are cryptosporidium, isospora, and the hooklets of cysticerci.

Gomori methenamine silver stain

This stain, often abbreviated as "GMS", is used to stain for fungi and for Pneumocystis carinii. The cell walls of these organisms are stained, so the organisms are outlined by the brown to black stain. There is a tendency for this stain to produce a lot of artefact from background staining, so it is essential to be sure of the morphology of the organism being sought.

PAS (periodic acid-Schiff)

This an all-around useful stain for many things. It stains glycogen, mucin, mucoprotein, glycoprotein, as well as fungi. A predigestion step with amylase will remove staining for glycogen. PAS is useful for outlining tissue structures--basement membranes, capsules, blood vessels, etc. It does stain a lot of things and, therefore, can have a high background. It is very sensitive, but specificity depends upon interpretation.

HISTOPATHOLOGYSTAINING TECHNIQUES

Stains used to identify ocular microbesq

a

Microbes Stain Colour

Bacteria Gram staining Positive blue/black a 

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Negative red a 

Chlamydia Giemsa stain blue-purple inclusion bodies a 

Mycobacteria Ziehl-Neelsen (ZN) 

red a 

Fungi Grocott hexamine (methenamine) silver

black a 

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Acanthamoeba cysts

PAS purple a 

Grocott hexamine  (methenamine) silver

black a 

calcofluor white

green fluoresence a 

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Stains used to highlight special substanceq

StainSubstance highlighted Colour

Alcian blue mucopolysaccharide blue a 

 macular dystrophy 

a

Alizarian red calcium red a 

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aband keratopathy a

Colloidal iron mucopolysaccharide blue a 

 macular dystrophy 

a

Congo red amyloid red a 

 lattice dystrophy 

a

Masson trichrome

hyaline materials red a 

 granular dystrophy 

a

oil red O lipid red 1 

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 fat within the cells 

a

Perl's prussian blue

iron blue a 

 iron in the epithelium in keratoconus 

a

TECHNIQUE PAGETaking Paraffin Sections to Water 2Dehydration and Clearing of Sections in Xylene for Mounting 2Blotting Sections Dry Before Mounting 3Mounting of Sections in DPX 3Ziehl-Neelsen Technique for Acid-Fast Bacilli 4-5Gram-Twort Modification for Bacteria in Paraffin Sections 6-7Periodic Acid Schiff Technique 8Periodic Acid Schiff / Alcian Blue Technique 9Buffered Congo Red Method for Amyloid 10Perls’ Prussian Blue Method for Haemosiderin 11

TAKING PARAFFIN SECTIONS TO WATERSAFETY NOTE: Turn on the exhaust system before commencing.Wear protective clothing, gloves and safety glasses during the procedure.xylene 2 minutesxylene 2 minutesabsolute alcohol 1 minuteabsolute alcohol 2 minutes70% alcohol 30 secondswater

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Take the slides through the various solutions as follows:1. Place slides in slide holder.2. Lift lid off staining dish, and immerse the slides in the first solution with agitation.3. Replace lid and allow slides to remain in solution for the specified time with periodic agitation.4. Remove slides from the solution (with agitation) and tilt slide holder to allow excess solution to drainbefore transferring it to the next solution.5. Continue in this manner through the remaining solutions for the specified timesTimes can be reduced if slides are agitated constantly.

DEHYDRATION AND CLEARING OF SECTIONS IN XYLENE BEFORE MOUNTINGSAFETY NOTE: Turn on the exhaust system before commencing.Wear protective clothing, gloves and safety glasses during the procedure.Dehydration 70% alcohol 15 secondsabsolute alcohol 1 minuteabsolute alcohol 2 minutesClearing xylene 15 secondsxylene 15 seconds

Take the slides through the various solutions as follows:1. Place slides in slide holder.2. Lift lid off staining dish, and immerse the slides in the first solution with agitation.3. Replace lid and allow slides to remain in solution for the specified time with periodic agitation.4. Remove slides from the solution (with agitation) and tilt slide holder to allow excess solution to drainbefore transferring it to the next solution.5. Continue in this manner through the remaining solutions for the specified times.Times can be reduced if slides are agitated constantly.

BLOTTING DRY AND CLEARING OF SECTIONSSAFETY NOTE: Turn on the exhaust system before commencing.Wear protective clothing, gloves and safety glasses during the procedure.Blotting Dry1. Place slide face down carefully on blotting paper.2. Fold blotting paper over slide over and apply gentle pressure to dry slide.3. Lift slide and move to dry section of blotting paper and repeat until section is completely dry.4. Allow section to air dry if necessary before clearing.Clearing xylene 15 secondsXylene 15 secondsClear dried section by dipping the slide in xylene for specified times and mount in DPX.

MOUNTING OF SECTIONSSAFETY NOTE: Turn on the exhaust system before commencing.Wear protective clothing, gloves and safety glasses during the procedure.If section is dry, dip in xylene before mounting the coverslip1. Remove slide from slide holder.2. Carefully dry the BACK of the slide with a tissue.3. Lay the slide on blotting paper or bench-coat on the fume bench.4. Place a drop of DPX mounting medium on the section.5. Carefully place a cover slip over the DPX ensuring that it covers the section.6. Remove any bubbles by jiggling the cover slip gently.7. Place mounted slide on small cardboard slide holder and place in 50oC oven for a few hours.

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8. Allow mounted slide to dry lying flat for at least 1 week.

ZIEHL-NEELSEN TECHNIQUE FOR ACID-FAST BACILLI (ZN)(Ziehl, 1882; Neelsen, 1883)The histological method is similar to the classical bacteriological technique that depends upon theresistance of certain bacilli to decolourisation by acid alcohol after being stained with hot carbol-fuchsin.

FixationMost fixatives can be used. Avoid Carnoy which removes lipid from the bacilli which makes them lessacid-fast. Formalin, especially when prolonged, is said to reduce acid-fastness, but specimens usually stainperfectly satisfactorily. The treatment of formalin fixed sections with 0.5% ammonium hydroxide beforestaining may improve the brightness of colour of acid-fast bacilli, but this is seldom necessary and maydetach the sections from the slide.SectionsThin (3-5 μ) sections.SAFETY NOTE: Turn on the exhaust system before staining.Wear protective clothing, gloves and safety glasses during the staining procedure.

ZN Staining ProcedureZN Procedure - Slide Method1. Place a rectangle of filter paper over the section (to prevent precipitation of the stain) and flood theslide with carbol-fuchsin.2. Warm the slide until the stain begins to steam; this can be conveniently done by the flame from athroat swab soaked in alcohol.3. Leave for 5 minutes.4. Wash in tap water for 2 minutes.5. Differentiate in acid alcohol (3% HCl in 95% ethanol) until no more colour runs from the slide.6. Rinse in water to remove acid alcohol.7. Counterstain in acidified methylene blue for 30 seconds.8. Wash in water, dehydrate, clear and mount in synthetic resin eg DPX.ORZN Procedure - Coplin Jar Method1. Take sections to water.2. Place the working solution in a coplin jar and pre-heat in 58 - 60oC waterbath for 10 mins3. Place the slide in the warmed vessel of carbol-fuchsin for at least 30 minutes at 58 - 60oC.4. Remove slide from coplin jar and wash in tap water for 2 minutes.5. Differentiate in acid alcohol (3% HCl in 95% ethanol) until no more colour runs from the slide.6. Rinse in water to remove acid alcohol.7. Counterstain in acidified methylene blue for 30 seconds.8. Wash in water, dehydrate, clear and mount in synthetic resin eg DPX.

ResultsAcid-fast bacilli: redOther bacteria: blueCells and their nuclei: blueRed blood cells should retain a slight red colour.

Notes1 Since this method involves the use of both acid and alcohol decolourisation, the risk of mistaking thenon-pathogenic acid-fast bacilli for the tubercle bacillus is decreased. In sections the tubercle bacilliwill be found in the abnormal areas (tubercles) and this risk is slight. The non-pathogenic acid-fast

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bacilli that are found in butter, milk, or cerumen are seldom acid-fast, but the smegma bacillus mayrequire prolonged alcohol decolourisation.2 Counterstaining should be light, especially in sections containing nuclear material, such as lymphoidtissue. Heavy counterstaining makes the identification of acid-fast bacilli difficult and may colourthem purple.3 Basic fuchsin specified for Schiff’s reagent may not give satisfactory results.4 Control sections of known positive tuberculous material with abundant acid-fast bacilli are valuablein that they indicate that the staining technique is satisfactory when several negative sections arebeing examined. They also give an idea of the colour of the acid-fast bacilli; this may vary from alight to a dark red.The leprosy bacillus is more easily decolourised than the tubercle bacillus, and differentiation must becarefully controlled. A faint residual red colour in the tissues is especially important, and sections that donot show this may be unreliable for the exclusion of leprosy.

Reagent Preparation1. Carbol FuchsinBasic Fuchsin 1.0 gAbsolute Ethanol 10 mL5% phenol in distilled water 100 mLDissolve the basic fuchsin in the alcohol, then mix with the phenol solution. Filter.2. Acid Alcohol (3% HCl)3% hydrochloric acid in 95% alcohol.3. Acidified Methylene Blue CounterstainMethylene Blue 0.25 gGlacial Acetic Acid 1 mLDistilled Water 99 mL

THE GRAM-TWORT MODIFICATION FOR BACTERIA IN PARAFFIN SECTIONSThe following modification of the Twort (1924) method for bacteria has the advantages of easierdifferentiation and a better colour contrast compared with the other Gram techniques (Ollett, 1947, 1951).The sections are easy to examine for long periods without eye strain.

FixationFormalin; other fixative can be used.SectionsThin (3-5 μm) paraffin sections.Safety Note: Turn on the exhaust system before staining.Wear protective clothing, gloves and safety glasses during the staining procedure.

Procedure1 Stain in 1% crystal violet for 3-4 minutes.2 Wash quickly in distilled water.3 Treat with Gram’s iodine for 3 minutes.4 Wash quickly in distilled water and blot dry.5 Decolourise briefly with 2% acetic acid in absolute alcohol until no more colour comes away the section should be a dirty straw colour at this stage.6 Wash quickly in distilled water.7 Counterstain in Modified Twort Stain in closed coplin jar for 5 minutes.8 Wash quickly in distilled water.9 Decolourise quickly and carefully in 2% acetic acid in absolute alcohol until no more red colour comes away (a few seconds).

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10 Clear in xylene and mount in a synthetic resin medium eg DPX.

Results Gram positive bacteria: blue-blackGram negative bacteria: pinkNuclei: redCytoplasm: light greenRed blood cells: greenPrinciple of Gram StainStain Gram-Pos Bacteria Gram-Neg BacteriaCrystal Violet with Iodine Mordant Blue-Black BlackAcid Alcohol Blue-Black ColourlessMethyl Red Blue-Black PinkIf sections are exposed too long to alcohol, the Gram-positive bacteria will also decolourise.Most bacteria, especially in large numbers stain a pale grey colour with haematoxylin.With all stains for microbes, it is essential that known positive control slides are stained along with the section.

Reagents1. Crystal VioletCrystal violet (CI 42555 ) 2.0 g; 95% alcohol 20.0 ml; ammonium oxalate 0.8 g; distilled water 80.0 mL. Dissolve dye in the alcohol & the ammonium oxalate in the dH2O, mix together. Mixture stable 2-3 years.2. Grams Iodine1.0 g Iodine crystals (harmful); 2.0 g potassium iodide (harmful); 300.0 mL distilled water. Dissolve KI in 2-3 mL only dH2O - the crystals will dissolve and the solution will become very cold. Dissolve the iodine crystals in the conc KI soln. Dilute mixture with the remainder of the dH2O3. Modified Twort’s Stain - Stock Solution (stable 1 year)0.2% Neutral Red in 95% Ethanol 90 mL0.2% Fast Green FCF in 95% Ethanol 10 mL4. Modified Twort’s Stain - Working Solution (prepare fresh)Dilute 1 volume of the stock solution with 3 volumes of distilled water.5. Acid Alcohol (2% Acetic)2% acetic acid in absolute alcohol.

PERIODIC ACID SCHIFF TECHNIQUE (PAS)Sections are oxidised by the periodic acid resulting in the formation of aldehyde groups. These then react with Schiff’s reagent (a leucofuchsin) to restore the quinoid chromophoric grouping, giving a magenta coloured final product to the PAS positive substances.

Procedure1 Take sections to water2 Remove mercuric deposit (if present) with iodine thiosulphate3 Oxidize with Periodic Acid 5 mins4 Wash well in running water for 5 minutes5 Rinse with distilled water6 Stain with Schiff's Reagent 15-20 mins7 Wash well in running tap water 5 mins8 Stain nuclei with Harris Haematoxylin 1 min9 Wash in running tap water 2 mins10 Differentiate briefly (1-2 seconds) with acid alcohol11 Wash and blue nuclei in ammonia water

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12 Wash briefly in running tap water13 Blue in ammonia water and running tap water14 Dehydrate quickly in alcohol, clear in xylene and mount in DPX

ResultsSimple polysaccharides, neutral mucosubstances, some macro mucosubstances and basement membranes are PAS positive (Magenta in colour)

Reagents1. 1% Aqueous Periodic Acid2. Schiff’s Reagent3. Harris’s Haematoxylin4. Ammonia Water5. Acid Alcohol - 1% HCl in 70% ethanol

PERIODIC ACID SCHIFF/ALCIAN BLUE TECHNIQUE (PAS/AB)Sections are oxidised by the periodic acid resulting in the formation of aldehyde groups. These then react with Schiff’s reagent (a leucofuchsin) to restore the quinoid chromophoric grouping, giving a magenta coloured final product to the PAS positive substances. By first treating the section with Alcian Blue the acid mucins will stain and therefore will not react when the section is subsequently stained with the PAS method. The PAS will stain neutral mucins and carbohydrates, red.

Alcian Blue/PAS Staining Procedure1. Take sections to water.2. Stain in 1% Alcian Blue solution for 10-15 minutes.3. Wash in running water for 2 minutes.4. Rinse in distilled water.5. Oxidize with Periodic Acid 5 mins6. Wash in running water for 5 minutes7. Rinse with distilled water8. Stain with Schiff's Reagent 15-20 mins9. Wash well in running tap water 5 mins10. Stain nuclei with Harris Haematoxylin 1 min11. Wash in running tap water 2 mins12. Differentiate briefly (1-2 seconds) with acid alcohol13. Wash briefly in running tap water14. Blue in ammonia water and running tap water15. Dehydrate quickly in alcohol, clear in xylene and mount in DPX

ResultsNeutral mucins magentaAcid mucins blueMixtures of above blue/purpleNuclei deep blueBasement membranes magenta

Reagents1 1% Alcian Blue in 3% Acetic Acid (pH 2.5)2 0.5 % Aqueous Periodic Acid3 Schiff’s Reagent4. Harris’s Haematoxylin

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5. Ammonia Water6. Acid Alcohol - 1% HCl in 70% ethanol

BUFFERED CONGO RED METHOD FOR AMYLOID (EASTWOOD AND COLE 1971)

FixativeBuffered formal-salineSafety Note: Turn on the exhaust system before staining.Wear protective clothing, gloves and safety glasses during the staining procedure.

Procedure1. Take sections to water.2. Stain in Harris’s Haematoxylin for 30 seconds.3. Rinse in tap water.4. Differentiate in acid alcohol for a few seconds (2 - 3dips)5. Rinse in tap water.6. Blue in ammonia water followed by running tap water.7. Stain in 0.5% Congo Red for 10-20 minutes.8. Differentiate in 70% alcohol for a few seconds (2 - 3 dips).9. Blot dry.10. Clear in xylene and mount in DPX.

ResultsAmyloid orange to redElastic tissue, eosinophils orange to redNuclei: blue

Notes1. Dichroism is pronounced and can assist in distinguishing amyloid from other tissue components.2. The stain has a permanent shelf life but may require occasional filtering.

Reagents1. pH 10.0 Sorensen-Walbum buffer0.1M Glycine (MW 72.07) 30 mL0.1M Sodium Chloride (MW 58.5) 30 mL0.1M Sodium Hydroxide (MW40.0) 40 mLMix together and check pH.2. 0.5% Congo Red Stain Congo Red 0.5gAbsolute alcohol 50mlpH 10.0 Sorensen-Walbum Buffer 50ml3. Harris’s Haematoxylin4. Ammonia Water5. Acid Alcohol - 1% HCl in 70% ethanol6. 70% Alcohol - 70% ethanol

PERLS’ PRUSSIAN BLUE METHOD FOR HAEMOSIDERIN (Perls)Ferric iron combines with potassium ferrocyanide to form the insoluble Prussian blue precipitate as follows:FeCl3 + K4Fe(CN)6 = KFeFe(CN)6¯ + 3KCl

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FixationNeutral formalin (acid fixatives and potassium dichromate should be avoided).

SectionsThin (3-5 μ) paraffin sections.SAFETY NOTE: Turn on the exhaust system before staining.Wear protective clothing, gloves and safety glasses during the staining procedure.

Procedure1. Take sections to water2. Rinse well in distilled water.3. Transfer sections to a mixture of equal parts of 2% Potassium Ferrocyanide and 2% Hydrochloric Acid for 20-30 min.4. Wash in tap water and then rinse in distilled water.5. Counterstain in filtered 1% neutral red for 1 minute.6. Rinse in tap water.7. Rapidly dehydrate in absolute alcohol, clear and mount.

ResultsHaemosiderin and ferric salts: deep blueTissues and nuclei: redCytoplasm pinkErythrocytes yellow

Notes1 More pronounced staining is obtained by heating the ferrocyanide to 37oC.2 If the stain fades it may be revived by treating with 10 vol water.3 Tap water must be avoided at all times. The distilled water must be iron-free, and the hydrochloric acid must be of analytical grade, or it will contain iron.

Reagents1. 2% Aqueous Solution of Hydrochloric Acid2. 2% Aqueous Solution of Potassium Ferrocyanide3. Perls Working Solution: Mix equal parts of 2% hydrochloric acid and 2% potassium ferrocyanide solution just before use.4. 1% Neutral RedNeutral red (CI 50040) 1.0 gDistilled water 99.0 mL Glacial acetic acid 1.0 mL

Coverslipping

The stained section on the slide must be covered with a thin piece plastic or glass to protect the tissue from being scratched, to provide better optical quality for viewing under the microscope, and to preserve the tissue section for years to come. The stained slide must go through the reverse process that it went through from paraffin section to water. The stained slide is taken through a series of alcohol solutions to remove the water, then through clearing agents to a point at which a permanent resinous substance beneath the glass coverslip, or a plastic film, can be placed over the section

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Decalcification

Some tissues contain calcium deposits which are extremely firm and which will not section properly with paraffin embedding owing to the difference in densities between calcium and parffin. Bone specimens are the most likely type here, but other tissues may contain calcified areas as well. This calcium must be removed prior to embedding to allow sectioning. A variety of agents or techniques have been used to decalcify tissue and none of them work perfectly. Mineral acids, organic acids, EDTA, and electrolysis have all been used.

Strong mineral acids such as nitric and hydrochloric acids are used with dense cortical bone because they will remove large quantities of calcium at a rapid rate. Unfortunately, these strong acids also damage cellular morphology, so are not recommended for delicate tissues such as bone marrow.

Organic acids such as acetic and formic acid are better suited to bone marrow, since they are not as harsh. However, they act more slowly on dense cortical bone. Formic acid in a 10% concentration is the best all-around decalcifier. Some commercial solutions are available that combine formic acid with formalin to fix and decalcify tissues at the same time.

EDTA can remove calcium and is not harsh (it is not an acid) but it penetrates tissue poorly and works slowly and is expensive in large amounts.

Electrolysis has been tried in experimental situations where calcium had to be removed with the least tissue damage. It is slow and not suited for routine daily use.

Artefacts in Histologic Sections

A number of artefacts that appear in stained slides may result from improper fixation, from the type of fixative, from poor dehydration and paraffin infiltration, improper reagents, and poor microtome sectioning.

The presence of a fine black precipitate on the slides, often with no relationship to the tissue (i.e., the precipitate appears adjacent to tissues or within interstices or vessels) suggests formalin-heme pigment has formed. This can be confirmed by polarized light microscopy, because this pigment will polarize a bright white (and the slide will look like many stars in the sky). Formalin-heme pigment is most often seen in very cellular or bloody tissues, or in autopsy tissues, because this pigment forms when the formalin buffer is exhausted and the tissue becomes acidic, promoting the formation of a complex of heme (from red blood cells) and formalin. Tissues such as spleen and lymph node are particularly prone to this artefact. Making thin sections and using enough neutral-buffered formalin (10 to 1 ratio of fixative to tissue) will help. If the fixative solution in which the tissues are sitting is grossly murky brown to red, then place the tissues in new fixative.

The presence of large irregular clumps of black precipitate on slides of tissues fixed in a mercurial fixative such as B-5 suggests that the tissues were not "dezenkerized" prior to staining. These black precipitates will also appear white with polarized light microscopy.

Tissues that are insufficiently dehydrated prior to clearing and infiltration with paraffin wax will be hard to section on the microtome, with tearing artefacts and holes in the sections. Tissue processor cycles should allow sufficient time for dehydration, and final ethanol dehydrant solution should be at 100% concentration. In humid climates, this is difficult to achieve. Covering or sealing the solutions from

35

ambient air will help. Air conditioning (with refrigerants, not with evaporative coolers) will also reduce humidity in the laboratory. Toluene as a clearing agent is more forgiving of poorly dehydrated tissues, but it is more expensive and presents more of a health hazard than other non-xylene clearing agents

Though alcohols such as ethanol make excellent fixatives for cytologic smears, they tend to make tissue sections brittle, resulting in microtome sectioning artefacts with chattering and a "venetian blind" appearance.

Bubbles under the coverslip may form when the mounting media is too thin, and as it dries air is sucked in under the coverslip. Contamination of clearing agents or coverslipping media may also produce a bubbled appearance under the microscope.

Problems in Tissue Processing

"Floaters" are small pieces of tissue that appear on a slide that do not belong there--they have floated in during processing. Floaters may arise from sloppy procedure on the cutting bench-- dirty towels, instruments, or gloves can have tissue that is carried over to the next case. Therefore, it is essential that you do only one specimen at a time and clean thoroughly before opening the container of the next case.

The best way to guard against unrecognized floaters is to always separate like specimens in the numbering sequence. For example, if you have three cases with prostate chips, separate them in accessioning with totally different specimens such as uterus or stomach. That way, if numbers are transposed or labels written wrong or tissue carried over, then you will have an obvious mismatch. Carrying over one prostate to another, or transposing the numbers of identical tissues may never be recognized.

If reusable cassettes are employed, you must be aware that tissue may potentially be carried over and appear as "floaters" even several days later, when the cassette is re-used. The problem arises when, during embedding, not all the tissue is removed from the cassette. Then, in the cleaning process, not all of the wax is removed. Then, the next person using the cassette does not pay attention to the fact that there is tissue already in the cassette and puts his specimen in it. The floater that appears on the slide will look well-preserved--it should, because it was processed to paraffin.

Always be sure that you properly identify the tissue! This means that you make sure that the patient label on the specimen container matches that of the request slip. An accession number is given to the specimen. This number must appear with the tissue at all times. You must never submit a cassette of tissue without a label. You must never submit a cassette of tissue with the wrong label. Mislabelling or unlabelling of tissues is courting disaster.

Safety in the Lab

The lab should be well-ventilated. There are regulations governing formalin and hydrocarbonds such as xylene and toluene. There are limits set by the Occupational Safety and Health Administration (OSHA) that should not be exceeded. These limits have recently been revised to reduced levels.

Every chemical compound used in the laboratory should have a materials safety data sheet on file that specifies the nature, toxicity, and safety precautions to be taken when handling the compound.

The laboratory must have a method for disposal of hazardous wastes. Health care facilities processing tissues often contract this to a waste management company. Tissues that are collected should be stored in

36

formalin and may be disposed by incineration or by putting them through a "tissue grinder" attached to a large sink (similar to a large garbage disposal unit).

Every instrument used in the laboratory should meet electrical safety specifications (be U.L. approved) and have written instructions regarding its use.

Flammable materials may only be stored in approved rooms and only in storage cabinets that are designed for this purpose.

Fire safety procedures are to be posted. Safety equipment including fire extinguishers, fire blankets, and fire alarms should be within easy access. A shower and eyewash should be readily available.

Laboratory accidents must be documented and investigated with incident reports and industrial accident reports.

Specific hazards that you should know about include:

• Bouin's solution is made with picric acid. This acid is only sold in the aqueous state. When it dries out, it becomes explosive.

• Many reagent kits have sodium azide as a preservative. You are supposed to flush solutions containing sodium azide down the drain with lots of water, or there is a tendency for the azide to form metal azides in the plumbing. These are also explosive.

• Benzidine, benzene, anthracene, and napthol containing compounds are carcinogens and should not be used.

• Mercury-containing solutions (Zenker's or B-5) should always be discarded into proper containers. Mercury, if poured down a drain, will form amalgams with the metal that build up and cannot be removed.

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