His to Pathology Department

TARIQ AZIZ / LAB APP OF HISTOPATHOLOGY DEPARTMENT

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HISTOPATHOLOGY DEPARTMENT

I - INTRODUCTION:

Cells are the building blocks of all living things. Groups of these cells unite to perform a specific function, these are called tissues. Microscopic study of individual cells in smears is called cytology and study of tissue is called histology. Histology is a branch of anatomy that deals with minute structure, composition, and function of tissues. Histopathology means study of diseased tissues. The pathologist responsible for the diagnosis of diseased tissues is dependent on the technical skill of a histotechnologist / cytotechnologist, who prepares microscopic slides of the specimen. These specimen slides should be well sectioned and properly stained to provide detailed information of the tissues under examination. Histopathological studies have proved to be one of the most effective in diagnosing tissue abnormalities, benign and malignant conditions.

Histopathological specimens are mostly collected by a surgeon in an operation room. The specimens in the form of pieces of tissues are submitted to the histopathology section of pathology department. Each specimen is immediately placed in a proper fixative and then it is entered in a log book (logging), labeled and given identification number. Histopathology request (Form No. 37) is completed in all respect by the consultant / physician and the nurse concerned. A morphological description of the tissue is noted by the pathologist and the gross examination of the tissue, a portion of tissue is trimmed into suitable sized blocks and handed over to the histotechnologist for further processing.


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II – RESPONSIBILITIES OF A TECHNICIAN: a.) Specimen preservation b.) Specimen logging c.) Preparation of specimen to facilitate their gross and

microscopic examinations to be performed by histopathologist

d.) Filing and preservation of records for future reference. III – BASIC STEPS FOR TISSUE PROCESSING:

a.) Fixation b.) Embedding c.) Microtomy d.) Staining e.) Mounting

IV – LABORATORY REQUIREMENTS:

a.) General glassware – pipettes, flasks, reagent bottles, etc. b.) Specimen containers – various sizes. c.) Couplin staining jars d.) Microscope slides and coverslips e.) Reagent bottles f.) Fixatives g.) Various organic solvents h.) Decalcifying solutions i.) Embedding materials j.) Various staining solutions k.) Various dilutions of ethyl alcohol l.) Mounting media etc.

V – EQUIPMENT AND INSTRUMENTS:

a.) Microscopes h.) Slide washing tray b.) Microtomes i.) Balance c.) Manual/Automated tissue processor d.) Paraffin oven j.) Automated staining e.) Tissue floating bath machine f.) Embedding oven g.) Slide warmer


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HISTOPATHOLOGY TECHNIQUES AND PROCEDURES FIXATION: Good fixation is the most important factor in the production of satisfactory slides. If the tissue is well fixed one rarely has difficulty with the later stages; if it is badly fixed, good sections are absolutely impossible. The three essentials are: fresh tissue, proper penetration of fixative and the correct choice of fixative. Surgical specimens should be placed in fixative immediately after they are taken and should be sent to the laboratory as soon as possible so that the pathologist can supervise their proper fixation. In postmortem cases, proper refrigeration of the body will do much to retard autolysis and permit reasonably good sections if delay is unavoidable. Even very early autolysis renders tissue abnormally fragile and such tissues should be handled as gently as possible to avoid artifact. Inadequate penetration of the fixative is one of the commonest causes of bad results. The maximum thickness that can be penetrated is about 10mm for loose tissues and 5mm for compact or cellular tissues (e.g. lymph nodes or spleen). For dealing with specimens bigger than this the following methods are recommended:

• Solid organs: Cut slices as big as necessary but not thicker than5mm. • Hollow organs: Either open out or fill with fixative or pack lightly with wool

soaked in fixative. • Large Specimens that require dissection: Inject fixative along vessels (or

bronchi, in case of lungs). CHOICE OF FIXATIVE The purposes of fixation are: 1. To inhibit autolytic enzymes and to kill the organism of

decompositions. 2. To preserve tissue as nearly as possible in its original form. 3. To protect the tissue against subsequent damage during

embedding. 4. To render the various constituents receptive to the proposed

stains. The choice of fixative will depend on the nature of the tissue and the type of staining to be employed.


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The following substances are commonly used in fixation: 1. Ethyl Alcohol

Used as 90-100% alcohol. Precipitates albumin and globulin but not nucleoprotein. Shrinks and hardens tissue and displaces cytoplasmic contents. Destroys mitochondria. A reducing agent, and therefore incompatible with chromic acid, chromates or osmium tetroxide. Preserves glycogen. Useful for histochemistry.

2. Formaldehyde Sold as formalin (a 40% w/w solution of gas in water). Used as 10% or better 15% of formalin in normal saline, or calcium chloride solution. Penetrates rapidly but fixes slowly. Does not precipitate protein but combine with NH2 groups to form an insoluble gel. Preserves practically all elements including fats. Renders phospholipids insoluble in fat solvents. Does not shrink but allows shrinkage during embedding.

3. Mercuric Chloride Used as saturated (about 70%) or half saturated aqueous solution. Oxidizing agent. Penetrates rapidly. Penetrates protein by combining with it. Causes no shrinkage by itself and subsequent embedding causes only minimal shrinkage. Fixes chromatin well and enhances its subsequent staining. Preserve nearly all elements. Forms precipitates in tissues. Corrodes metal containers. Rarely used along but it is valuable constituent of many other fixatives.

4. Acetic Acid Used as 1-5% aqueous solution. Precipitates and fixes nucleoprotein but not cytoplasm and is, therefore, valuable for nuclear fixation. Penetrates extremely rapidly. Tends to swell tissue and does not harden. Destroys mitochondria and cytoplasmic granules. Rarely used alone.


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5. Picric acid Used as saturated aqueous solution (about 1%). Penetrates poorly and causes shrinkage but does not harden. Precipitates protein by combining with them. Preserves glycogen and nearly all other elements. Permits nearly all stains. Not used alone.

6. Chromic acid Used either as a pure chemical or as a mixture of dichromate and acetic acid (e.g. Zenker). An oxidizing agent and therefore, incompatible with formalin or alcohol. Fixes by both denaturing and precipitating proteins. Causes moderate shrinkage but does not harden badly. Penetrates slowly. Preserves most elements. Tends to weaken nuclear staining by dissolving nucleoprotein.

7. Potassium dichromate Used as 2-3% aqueous solution. A weak oxidizing agent and cannot therefore, be kept with formalin for long. Fixes by making proteins insoluble in water but without actually precipitating them. Tends to dissolve chromatin and therefore weakens nuclear staining. A good cytoplasmic and bad nuclear fixative. Fixes lipids and mitochondria well. Causes no shrinkage but allows no shrinkage during embedding. Gives chromaffin reaction.

8. Osmium tetroxide Used as 1% or 2% aqueous solution. Expensive and unstable with an irritating vapor. Powerful oxidizing agent and therefore incompatible with formalin or alcohol. Penetrates very badly. Fixes by setting proteins in an insoluble gel without precipitating them. Preserves fats and gives black precipitate of osmium dioxide with unsaturated fats. Also preserves very fine cell detail e.g. Golgi bodies. Used mainly for electron microscopy. (Store in dark bottle with 1 drop of saturated aqeous HgCl2 to each 10cc to prevent oxidation).


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TYPES OF FIXATIVE: The number of features available is almost infinite. The following list gives only those in regular use in this department but is sufficient for all ordinary purposes . Note: If dichromate is used it is better to wash the tissue in water before embedding to prevent the mercury corroding instruments. Mercury must be removed from the cut sections with iodine before staining but there is no need to add iodine to the dehydrating alcohol, it does not help.

1. Formol Saline

• Commercial formalin 15ml • 0.85% aqueous solution of NaCl 85 ml (Keep over marble chips or calcium hydroxide chalk to absorb acid). The acid only affects the staining and adds to quantity of formalin pigment formed with fre Ebgin tissues. Formalin is a very cheap and very popular fixative but it has certain definite disadvantages. Its advantages are as follows: 1. It is the only fixative foe specimens destined for tissue

mounting photography. 2. It is the only fixative which leaves tissues suitable for dissection. 3. By virtue of its cheapiness it is desirable for big specimens. 4. It is the best fixative for frozen sections. 5. It is the only fixative for many of the silver impregnation methods.

e.g., For neurologia or for spirochaetes, though silver reticulin and Bodian can be done in other good fixatives, very good for myelin.

Its Disadvantages are as follows: 1. It acts slowly and requires several days to achieve complete fixation. 2. It allows considerable shrinkage during subsequent embedding. 3. It forms a black pigment with laked blood if the pH is less than 6.8

(e.g. in stale or autolysing tissue). Note: The shiny pale blue glittering nuclei indicate poor fixation.


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2. Formol Calcium • Commercial formalin 15ml • 1% aqueous solution of calcium chloride 85ml Used where the preservation of small quantities of fat is important. Not as good as formol-saline. Used only for analytical histochemistry.

3. Formol-Mercury

(Formalin prevents protein flocculation caused by Hg). • Mercuric chloride 70g (3.5%) • Commercial formalin 350ml (17.5%) • Water 2 liters (We add Edicol pea green to aid recognition) The color also helps keep track of tiny biopsies. The original formula (Lendrum,1941) gave saturated mercuric chloride (7% approximately) and 10% formalin. Our modification causes less hardening and is cheaper. This is probably the best routine fixative. It is rapidly acting and causes less shrinkage and gives much more brilliant staining and sharper detail than formol saline. It is equally useful for fresh biopsies or stable autopsies. It permits all ordinary stains including the silver reticulin methods. Time of fixation: 6-24 hours depending upon size. Prolonged time does no harm.

4. Helly’s fluid (Modified by Maximow) Also known in American literature as Formol-Zenker

• Potassium dichromate 2.5g • Mercuric chloride 5.0g • Sodium sulphate 1.0g • Water 100 ml (Add 10ml of commercial formalin before use)

This is an excellent fixative and unlike Zenker’s, it preserves cytoplasmic granules well. We use it for all hemopoietic tissues or other tissue which needs to be stained by Giemsa, Leishman, etc. Improved in routines. Time of fixation: 3-18 hours, according to thickness. More than 24 hours is harmful. Zenker contains acetic acid. This produces chromic acid which is not a good fixative and is therefore not recommended. The formalin in Helly’s allows chromate to remain. Potassium dichromate if applied for long periods interferes with nuclear stain. For human tissues optimum time is 18-20 hours while for rat tissues 6 hours.


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5. Bouin’s fluid • Saturated aqueous picric acid 75ml • Commercial formalin 25ml(25%) • Glacial Acetic acid 5ml

This is an excellent cytological fixative which preserves all detail well. It is useful for small biopsies of cellular tissue but is not satisfactory for large specimens or as a general fixative because it penetrates badly and ruins red blood cells. It is a poor fixative for autopsy tissue. Bad for brain tissues. Blocks have to be thin. Preliminary fixation formol. Time of fixation: 6-24 hours. Unduly long fixation (several days) is harmful.

Transfer to alcohol not water). Saline prevents the bad effects of Bouin and this gives

excellent results. 6. Heidenhain’s Susa

• Mercuric chloride 45g 4.5% • Sodium chloride 5g 0.5% • Trichloracetic acid 20ml 2% • Glacial acetic acid 4ml 4% • Commercial formalin 200ml 20% • Water 800ml

This is a very good general fixative. It gives brilliant staining and very sharp nuclear detail. It tends to dissolve cytoplasmic granules and makes the cytoplasm rather transparent. It penetrates and fixes rapidly and neither shrinks nor hardens. It permits nearly all stain, including silver reticulin methods. It preserves red cells badly and is rather expensive. Useful when space is essential. Nuclear stain is excellent and is good for mitotic fig. Studies and hence ideal for tumors. Time of fixation: 6-12 hours. Transfer to alcohol 90-96% which further saves time 7. Carooy

• Alcohol 60ml • Chloroform 30ml • Glacial Acetic acid 10ml

This fixative gives sharp clear staining but not fine detail. It preserves glycogen and is better general fixative than alcohol but not as good as Bouin, It hemolyzes red cells but leaves the empty envelops intact. It dehydrates as well as fixing and is useful if speed is essential. Very poor fixative but contains no water. Good for Mucin MPS. Time of fixation: 12 hours depending on size. Longer does no harm. Transfer to

alcohol only. 8. Muller’s fluid

• Potassium dichromate 2.5g • Sodium sulphate 1.0g • Water 100ml

Not now used as a fixative. See under Marchi method.


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SPECIAL HISTOCHEMICAL FIXATIVES Modified Gendre fluid (for glycogen)

• Saturated picric acid in 96% alcohol 85 parts • 40% formaldehyde 100parts • Glacial acetic acid 5parts

Time of fixation: (Small blocks only) 18 hours at 73o ± 12 hours at –20oC.

The lower temperature is preferable if diffusion artifacts is to be avoided.

Cold Formalin/ Acetone (for phosphatases and esterases) 1. Fix thin slices of tissue in 10% neutral formalin (buffered to ±

pH 7.0 with either sodium acetate or diphasic sodium phosphate) for 12-16 hours at 0 to –4 oC

2. Wash for 6-24 hours in running cold water. 3. Dehydrate in 2 changes of absolute acetone at 0 to

–4oC in 1 hour in each. 4. Continue dehydration in absolute alcohol at room temperature

for 1 hour. 5. Clear in toluene. 6. Embed in paraffin wax, preferably at 52-54oC in vacuo. Time 1

hour or 10 minutes in vacuo. The following fixatives are recommended for fixation of cryostat sections before carrying out histological or histochemical procedures.

Fixative Temperature Time Methods Acetone 4oC 1 hr Phosphatases

Formol Saline 4oC 1 hr Esterases

Formol Saline 20oC 5-60 min Histological Methods

Formol Calcium 4oC Block 18 hrs

1-3 hours

A good block fixative for histochemistry (especially lipids) also good for phosphatases and esterases.

Formol Alcohol 1 part in 9 parts

20oC 3-6 hours Glycogen and mucins (also can be used as a rapid fixative for histology).

Wolman 5% Acetic

20oC 5-30 min Nucleic Acid


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PRACTICAL GUIDE TO CHOICE OF FIXATIVES It is difficult to give general rules because the choice depends on the nature of elements to be demonstrated and the stains to be employed. The following list of method in use in this Department is meant to be a guide. • Autopsy tissues (do not allow sandwiching of blocks) Thin slices ( not more than 5mm thick) are dropped into an

excess of formal saline. Blocks are trimmed to shape and refixed in formol mercury (F.M.) in the tissue processor.

• Small cellular biopsies (Tumour biopsies, cytology endometrium).

F.M. • Large Surgical Specimens

Place the whole in formol saline in theatre. Dissect and select blocks. Refix in F.M. in tissue processor.

• Tissue for frozen sections Formol Saline (F.M. is possible but difficult) Freeze unfixed: block cut cryostat section. Take one direct

to stain, 2nd fix it in an alcohol other mixture for 1 minute. To fix as the permanent section. Unfix will decompose.

• Nervous tissue Formol Saline

• Specimens for museum mounting Kaiserling’s solution

• Haemopoietic tissue Helly

• Histochemical tests Look up appropriate fixative

Formol Saline followed by Bouin’s or by Helly’s is best on formol mercury. Gelatin blocks can be used to serve as a base for very tiny tissues on frozen section. Gelatin molten is poured in dish allowed to set and fixed in formalin. Cut into blocks and stored in the fridge.


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PARAFFIN EMBEDDING

Blocks of tissues are trimmed to suitable size after fixation. This should be 3-5mm thick and preferably small enough to fit under a standard (7/8” x 7/8”) cover-slip. There is no advantage and much disadvantage un using unduly small blocks since they give an inadequate sample of tissue. If the tissue has been fixed in bichromate and has not already been washed, the blocks should be washed overnight in running water. • Dehydration: The usual method of dehydration is the use of ascending

concentration of ethyl, isopropyl, or butyl alcohol; we use the first. It is important to avoid sudden changes from water to high concentrations alcohol because this causes the tissue to shrink and harden and consequently makes it difficult to cut. Dehydration should be in graduated steps through multiple baths.This can be done easily and cheaply by using old used alcohol for the earlier baths.

• Clearing: The more useful agents are xylene, benzene, kerosene, and chloroform. Xylene and Benzene are cheap and rapid but tend to harden the tissue badly (benzene is the better of the two). Chloroform is slow and more expensive but does not harden tissue nearly so badly. It can be used several times. Kerosene is cheap and does no harden but will not mix with alcohol unless it is over 99%. We use a mixture of 3 parts chloroform to 7 parts kerosene.

• Wax impregnation: Paraffin wax with a melting point of about 56oC is generally used and kept in an oven at 58-59 oC. It is important to see that the oven temperature does not rise above this because temperature of much over 60 oC will harden tissues badly. It is an advantage to keep the tissues in the paraffin oven for the minimum time consistent with complete impregnation. This is best achieved by passing the tissue through a series of baths of wax; the clearing agent diffuses out into the wax quite rapidly but takes so many hours to evaporate. A vacuum embedding oven, if available, is very rapid.

• Casting: When the tissues are impregnated they are cast in fresh wax. They can be cast in boxes of folded paper metal containers or boxes made with a metal or glass plate and brass angle pieces. The tissue is carefully oriented sot that the surface to be cut is downwards and the wax is then rapidly cooled to prevent it s crystalling. If, inspite of rapid cooling the wax crystallizes, it can be mixed with a small quantity of beeswax (about 2g in a pint of molten wax); this will usually prevent crystallization.

• Labeling: The most serious error which can occur in biopsy diagnosis is the muddling of specimens. It is, therefore, essential to use a labeling technique which is as nearly as possible foolproof. The best method is to use slips of cardboard or cartridge paper (1 x 4cm) with the number written in black lead and carry these through the whole process with the specimen.


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Embedding Technique The embedding technique used in this department is as follows: The fixed specimens are examined, their morbid anatomy is described and blocks are selected about tea time (this allows for specimens arriving late in the day). The blocks and their numbered labels are placed in the automatic tissue processor. This is set to run from 5:30pm to 9:15am giving the following changes of reagent: • Formol Mercury for further fixation ………...2½ hrs • Alcohol 4 times used…………………………¾ hrs • Alcohol 3 times used………………………...1 hr • Alcohol twice………………………………..1 ¼ hrs • Alcohol once used…………………………...1 ¼ hrs • Fresh alcohol (74 O.P.)……………………...1¾ hrs • Kerosene-chloroform mixture(7:3 parts)……1 hr • Kerosene-chloroform mixture……………….2½ hrs • Pure chloroform……………………………..1 hr • Paraffin wax (M.P. 54oC)……………………1 hr}*All 3

baths • Paraffin wax (M.P. 56oC)……………………1 hr}*kept

at • Paraffin wax (M.P. 56oC)……………………1 hr}*59-

60oC This tissues are cased in the morning and cut in the afternoon. Autopsy tissues are processed by hand if the machine is too full of biopsy specimens. The technique is as follows: The blocks together with their numbered labels are placed in old (4 times used) alcohol (about 70% alcohol) overnight. Next morning they are transferred to three times, used alcohol and subsequently to twice used, once used, and finally fresh alcohol (74 O.P.) These 4 baths take about 6 hrs and in the afternoon the tissues are transferred to used kerosene chloroform (i.e. kerosene-chloroform containing some alcohol). At the end of the day they are transferred to fresh kerosene chloroform where they remain


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overnight. The following mornings they are put through 4 baths of wax in three hours and then cast in fresh wax. Common Technical errors: • Incomplete fixation: This usually affects the center of the block. If

severe, the tissues crumble on cutting and will not adhere to slide. With less severe degree the nuclei look homogenous and take only a pale blue stain.

• Incomplete dehydration: If xylene or benzene are used for the clearing, this error can be recognized because the center of the block fails to become translucent. With chloroform, which does not render tissue transparent, it cannot be recognized until the block is cut, when it will be found that the middle of the block is soft and not impregnated with wax.

• Incomplete impregnation: the middle of the block is soft and smells of the clearing agent.

• Crystallization of the wax: The wax block should be homogenous and translucent. It is streaky and opaque the wax may have crystallized due to slow cooling or unsuitable wax.

• Unduly hard blocks: The block may be so hard that it cannot be cut properly. The tissues commonly affected are dense collagen, bone, skin, eye and colloid goiters. The commonest causes are: incomplete decalcification, bad fixation, sudden jumps from water to high grade alcohol, too long in xylene or benzene (not chloroform), too long in paraffin oven, too high temperature in the paraffin oven. It is sometimes found the very fatty tissues (e.g. lipoma) crumble and fail to cut properly. This may be due to failure to remove all the fat. If this occurs, give the tissue an extra bath of clearing agent.

The nomenclature of the different forms of alcohol is muddling. The following data may be helpful.

Proof spirit: Originally the mixture of alcohol and water when used to wet gun powder; would just permit it to burn. Now defined as that mixture which at a room temperature of 60oF will weigh 12/13 of the weight of an equal volume of water. In practice this is 49% of alcohol by weight and 60% by volume.

Under-proof and over-proof: A spirit that is X under proof will become proof spirit


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when X parts of pure alcohol are added to 100 parts of it. A spirit that is X over proof becomes proof spirit when X parts of water are added to 100 parts of it.

Methylated spirit: (64 O.P.) consists of rectified spirit (approximately 95% ethyl alcohol) to which 10% of wood spirit (about 95% methyl alcohol) has been added. To this pyridine and a dye are added before sale to the public.

Absolute Methylated spirit: (74 O.P.) consists of a 10:1 mixture of ethyl and methyl alcohols with about 1% of water.

Absolute Ethyl alcohol: is pure ethyl alcohol and is nearly anhydrous (about 0.3 to 0.5% of water).

CALCIFIED TISSUES Bone

For ordinary specimens of bone the following technique gives reliable results in a reasonable length of time. • Fix in formol saline or formol mercury. • Trim the pieces of tissue to a reasonable size and a

thickness of about 3mm (a fret-saw is useful for this). Refix if necessary.

Decalcifying solution: Formic acid 15 ml Sodium citrate 5 g Water 100 ml

• Place tissue in an excess of this solution and keep it agitated. (We use the tissue processor, this is essential to shorten the time). Test for complete decalcification either by testing for calcium in the solution or by x-rays.

• When completely decalcified wash in running water overnight.

• Embed in wax in the ordinary way. • Stain as desired.


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Calcified tissues Calcified tissues, e.g. heart valves, are treated as above. The time needed in acid varies with the tissue.. Undecalcified bone To distinguish between bone and osteoid the most reliable method is to cut undecalcified sections and stain the calcium by Kossa’s method. This is not unduly difficult and the distinction between bone and osteoid. • Fix in formol saline. • Cut out a small piece of cancellous bone about 5 x 5 x 2 mm. (This

can be done with a sharp knife). • Dehydrate in the ordinary way as far as absolute alcohol. • Immerse in 3% nitrocellulose in methyl benzoate (8 hrs) • Transfer to 6% nitrocellulose in methyl benzoate (8 hrs) • Transfer to 12% nitrocellulose in methyl benzoate (8 hrs) • Transfer to 20% nitrocellulose in methyl benzoate (overnight) • Drain off the excess nitrocellulose and immerse the tissue with its

adherent nitrocellulose in Benzene (2 changes of ½ hour each). This doubly embedded block can be cut on a sledge-type microtome using a heavy razor without damaging the latter unduly but the sections break to pieces in the process of cutting and the fragments can not be picked up. To prevent this a strip of “Sellotape” is pressed on to the top of the block so that when the next section is cut it remains stuck to the “Sellotape”. The strip of “Sellotape” can be moved fprward with each cut until sufficient sections have been obtained. The individual sections are cut with scissors and attach to slides by pressing them on to a warmed, albuminised slide (sticky side to glass), The tape can be floated off by wetting it with wet blotting paper for 10 minutes. The adhesive can then be dissolved off in warm benzene leaving the section of bone attached to the slide. • Stain the calcium by Von Kossa’s method and counterstain with Van

Gieson’s stain for 30 seconds only. • Dehydrate, clear and mount. Result : Bone - Dark brown to black Osteoid - Red ADHESION OF SECTIONS TO SLIDE PARAFFIN SECTIONS

Sections that are well fixed adhere well. Badly fixed sections tend to float off. Thick sections tend to float off more easily than thin ones. All sections float off dirty or greasy slides. Prolonged


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staining methods or any method involving ammoniacal solutions (e.g. silver) tend to loosen sections.

While well fixed sections will generally adhere to clean slides without aid, it is a wise precaution to use some adhesive. The following two methods are in use in this department and work well.

1. Glycerine Albumin: This is a mixture of glycerine and egg white

and can be made or bought. A very tiny drop (pinhead or less) is rubbed over the surface of the slide before floating the section on to it from the hot water bowl. If you are too slow in picking up the , the solution gets washed away. This method depends on the albumin being precipitated by alcohol when bringing the section down to water.

2. Chromate-Gelatin: Dissolve a fragment of leaf gelatin about ½”

square and a crystal of potassium dichromate about as big as a split pea in the hot water on to which the sections are floated. The gelatin dissolved in the water is denatured by the chromate in the presence of light and becomes insoluble in water after the section has been dried.

FROZEN SECTIONS:

Gelatin 0.5g Merthiolate 15mg (preservative) Distilled water 100ml

• Warm to 56oC and dip clean slides in the solution to coat them • Dry coated slides at room temperature under cover to avoid dust.

Store in boxes for use. • Float frozen sections on to slide and flatten out. • Blot carefully. • Place slide in Couplin jar with a few ml of commercial formalin for 1-2

minutes for vapor to denature gelatin. • Wash in tap water to remove formalin. • Stain sections in the ordinary way. MOUNTING MEDIA The choice of a suitable mountant is important to avoid fading of sections. 1. D.P.X. Dibutyl phthalate 5ml Xylene 35ml Mix and add distrene 80


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• D.P.X. can be bought ready made. • A good general mountant. Does not cause fading as

badly as balsam. Shrinks a good deal. Must be in a thin layer or it distorts the microscopic image. Dries very rapidly. Used as a routine in this department.

2. Canada Balsam: A very old mountant. Easily becomes acid and causes

fading of sections. 3. Chrome Glycerine Jelly: Distilled water 80ml Glycerine 20ml Powdered gelatin 3g Chrome alum 0.2g

• Dissolve the chrome alum (chromium potassium sulphate) in 30ml of water. Dissolve the gelatin (edible gelatin from the grocer’s) in 50ml of water. Mix and add the glycerine. Test pH with indicator and neutralize with NaHCO3 if acid.

• This mountant does not cause fading like ordinary glycerine jelly and also sets much harder, obviating the need to ring sections.

4. Apathy’s aqueous medium Gum acacia (gum Arabic) 50g Sucrose (cane sugar) 50g Distilled water 100ml

• Dissolve at 55oC – 60oC with shaking. Top up to 100ml. If necessary, add 15mg of merthiolate. Place in vacuum oven while warm to get rid of bubbles.

• Add: Potassium acetate 50g or NaCl 10g

- to prevent violet dye leaching out of amyloid. (This medium sets hard and does not need ringing).


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STAINS FOR NUCLEI Three groups of stains are available: haematoxylin, celestine blue and a group of red dyes.

1. HAEMATOXYLIN This is an extract of a tree (log wood) and is bought as a brown powder. Used alone it is quite useless but it has the property of combining with various heavy metals to form dyes known technically as “lakes”. The metals most commonly used are iron, aluminum, tungsten, and lithium but others have been used. As a general rule the iron lakes give intense grey-black or blue-black nuclear staining which is resistant to subsequent decolorization; these dyes are used when the counterstain chosen is liable to decolorize more labile haematoxylin or where very sharp definition of the nuclear chromatin is required, as in the study of chromosomes. Examples are Weigart’s and Haidenhan’s hematoxylin. The aluminum “lakes” (known as haemalum) give a lighter, more transparent blue color but are labile and change color to red in the presence of acids. They are very simple to use and are, in fact, the common routine stains for nuclei. Examples are Erlich’s, Harris’ and Delafield’s haemalum. Tungsten is used as phosphotungstic acid and the lake produced is used to stain tissue components as well as nuclei. Lithium-hematoxylin lake can be used to stain myelin sheaths.

2. CELESTINE BLUE: Celestine blue ( a synthetic oxazin dye) combines with iron to form a lake which stains nuclei blue. If, however, sections are stained for a few minutes in this lake and then restained in haemalum the resulting stain is as resistant to subsequent decolorization as an iron-hematoxylin stain. This method,


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being rapid and easy, has become a popular substitute for Weigart’s iron-haematoxylin.

3. RED NUCLEAR STAINS:

Red nuclear stains are sometimes necessary as a contrast to some other blue or black dye such as Perl’s stain for iron or Masson’s method for melanin. The three common ones are Alum-Carmine, Carbol-Safranin, and Neutral red.

FORMULA AND TECHNIQUES HAEMATOXYLIN AND HAEMALUM 1. Heidenhain’s Iron Hematoxylin:

Sol. A 5% (approx) aqueous iron ammonium sulphate (use lilac crystals.

Sol. B 0.5% (approx) aqueous hematoxylin (dilute a stock 10% alcoholic solution with water)

• Bring paraffin sections down to water. • Mordant in Sol. A for 1 hour at 37oC or up to 12

hours at room temperature. • Rinse (do not wash for long). • Stain in Sol. B until quite black (about the same

time as for mordant). • Rinse • Decolorize in saturated alcoholic picric acid until

staining is correct.


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• Wash well in running water. (All picric acid must be removed).

• Counterstain if desired, dehydrate, clear and mount. 2. Weigert’s Iron Hematoxylin: Sol. A 1% alcoholic haematoxylin Sol. B Liq. Ferri. Perchlor.(B.P.) 4ml Distilled water 100ml Conc. HCl 1ml

• Mix equal volumes of Sol. A and B in test tube and pour the mixture on to the slide. (The mixture will only keep for a few hours).

• Stain for 15-30 minutes • Rinse • Decolorize in acid alcohol • Wash, counterstain, etc.

3. Erlich’s Haematoxylin: 2% alcoholic haematoxylin 100ml Water 100ml Glycerine 100ml Glacial acetic acid 10ml Potassium alum To excess

• Plug bottle with wool (not cork) and leave in the light to ripen for a few weeks. The stain is usually fit for use when it has darkened to a rich Burgundy color.

4. Mayer’s Acid Haemalum: Haematoxylin 1g }

Sodium iodate 0.2g }Dissolve overnight Potassium alum 50g }then add: Distilled water 1000ml} Chloral hydrate 50g Citric acid 1g


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• Boil 5 minutes. Ready for use when cool.

5. Harris’ Haematoxylin: 10% alcoholic haematoxylin 10ml 10% aqueous ammonium alum 200ml

• Mix and bring to boil. When boiling, add 0.5g yellow mercuric oxide. When the solution turns purple, plunge flask into water and cool rapidly. When cold, filter and add 4ml glacial acetic acid. This stain is artificially oxidized and can be used immediately.

• Good as nuclear stain in frozen section for fat as well

as in cytology smears. 6. Delafield’s Haematoxylin: Saturated aqueous ammonium alum 40ml Haematoxylin 4g Absolute alcohol 25ml

• Mix and ripen for a few days and then add 100ml of glycerine and add 100ml of methyl alcohol. Ripen for a further six weeks. Note: Liq. Ferri. Perchlor is an aqueous

solution of ferric chloride. Iron ammonium alum is iron ammonium

sulphate. Potassium alum is potassium aluminum

sulphate. Ammonium alum is ammonium

aluminum sulphate.


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• It is very convenient to make up a 10% alcoholic solution of haematoxylin as a stock solution from which all the above can be made by diluting with alcohol or water.

• Such a stock hematoxylin slowly

ripens on its own.

7.Cole’s haematoxylin:

Haematoxylin 3g in 25ml of 75 OP spirit. Add this to 500ml of warmed D.W., dissolve and 100ml of 1% alcoholic iodine. Add 1,400ml of saturated aq. am. alum (mordant).

• Bring to boil – must boil – cool and filter • Use: No need for ripening.

COUNTERSTAINS Most pathologist use Eosin as a general counterstain, but some prefer more complex mixtures such as phloxine-tartazine. Van Gieson or even Masson’s trichrome. In this department, Eosin is used as a routine and other counterstains are only employed to demonstrate particular tissue elements. Eosin is used as a 1% aqueous solution of the water soluble yellowish variety. If staining is found to be weak, the dye can be improved by adding 2% calcium chloride or a trace of acetic acid. Too much acid causes precipitation. The dye will usually grow molds but these are harmless and can be filtered off.


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To get good results, tisuues should be heavily overstained with eosin and then differentiated in running water. This also improves the keeping properties of the section. In a good preparation, red cells, muscle and connective tissue should be in three distinct shades. ROUTINE HAEMALUM AND EOSIN STAINING: Any fixative. Paraffin Sections. 1. Warm slide and treat with xylene to remove wax. 2. Treat with alcohol to remove xylene. 3. Rinse in water. 4. Treat with Lugol’s iodine to remove mercury if necessary 5

minutes. 5. Treat with 5% sodium thiosulphate till white. 6. Rinse in water. 7. Stain in any haemalum till overstained. 8. Rinse in water. 9. Differentiate in acid alcohol till nuclei only are stained. 10. Rinse. 11. Scott’s tap water substitute 5minutes. 12. Wash in water for a few minutes. 13. Stain in Eosin till section is bright red (5-15 minutes). 14. Wash in running water till eosin is differentiated. 15. Dehydrate in alcohol. 16. Clear in xylene. 17. Mount in D.P.X. NOTES: If desired, sections may be stained for 2-5 minutes in

celestine blue and rinsed in water between stages 6and 7. If this is done, 5 minutes in haemalum usually suffices.

• Lugol’s iodine: Potassium iodide 2, water 100. • Acid alcohol: Conc. Hydrochloric acid 9ml.

Absolute methylated spirit 99ml OTHER NUCLEAR STAINS: 1. Celestine Blue:


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• Dissolve 2.5g of iron ammonium alum in 50ml of distilled water.

• Add 0.25g of celectine blue B. • Boil, cool and add 7ml of glycerol

2. Alum Carmine: (Carmalum) Carminic acid (BDH) 1g 5% aqueous ammonium alum 200ml Salicylic acid 0.2g • It is impossible to overstain with this. If the result is

too weak, rinse and stain for 1 minute in neutral red.

3. Carbol-Safranin:

• Melt 0.5g of phenol in a dry flask under the hot tap.

• Mix in 0.1g of safranin to make dark red sludge. • Grind together 0.25g of starch and 0.25g of

destrine. • Add 50ml of water, with further grinding. • Heat to 80oC. Cool, filter and dissolve the

carbolsafranin sludge in this filtrate. • Stain for 10 minutes and differentiate in the

dehydrating alcohol. 4. Neutral red- Carbol fuchsin: 1% aqueous neutral red 15 parts Carbol fuchsin (Ziehl Neelsen) 1 part

• Stain for ½ to 2 mins (alcohol tends to remove the stain).

Notes: Scott’s tap water substitute:

Potassium bicarbonate 2g Magnesium sulphate 20g Distilled water 1 liter


25

This neutralizes the acid and makes the section blue without loosening the section from the slide. It is not necessary if tap water is alkaline but does save time.

Alcohol: For all dehydrating of tissues and sections we use absolute methylated spirit which is 74 overproof and consists of methyl alcohol with 90% a small impurity of methyl alcohol. It is about 99% alcohol and is as good as and cheaper than absolute alcohol. For making up solutions however, pure absolute ethyl alcohol should be used.

Xylene: Some batches acid. If so, they will cause fading of slides. This can be cured by standing the stock xylene over marble chips.

FROZEN SECTIONS Indications: 1. To stain fats. 2. Rapid diagnosis during an operation. 3. Some special stains (e.g. neurologia or histochemical tests). 4. To minimize artifacts. 5. To cut very thick sections. Fixation:

• Formalin: All other fixatives make tissues too brittle. For speed, small blocks may be fixed for 2 minutes in formol-saline at 80oC, but sections obtained in this way are of poor quality.

• Embedding in gelatin is useful for friable tissues.

Cutting:

• Set the microtome at 15μ and start with the knife about 1mm (not less) from the surface of the chuck.


26

• Face the fixed block carefully, lay the flat surface (moist but not wet) downwards on the chuck, and press firmly so that it makes contact all over, especially at the corners.

• Freeze, with short sharp burst of CO2 until a thin white line of frozen tissue appears along the lower margin of the block. The block is now fixed to the chuck and the knife should be brought across to slice off the upper part of the block before the plane of section freezes. The flat surface for sectioning is now prepared. Continue freezing until the new upper surface of the block just balances, then let it thaw slightly.

• Start cutting. The ideal is to cut through tissue half thawed, supported by frozen tissue a few μ below.

• Wipe each section off the blade with the little finger of the left hand wet in 50% alcohol: the knife is thereby kept wet with alcohol and sections run up on its surface much more easily. Float each section off the finger in a dish of water. The first few sections are usually useless. Once one good section is obtained a series out at intervals of about 5 seconds will usually stay in the right zone of semi-thawing. Do not forget an occasional blast of CO2 to keep the bottom of the block frozen to the chuck (the next section after this is usually a failure). Do not cut right down into the chuck.

Liver and kidneys are the easiest tissues to cut. Necrotic material and normal lung the hardest. With practice favourable tissues may be cut at 10μ. The nearer the chuck the knife is set the less time it takes to freeze the tissues and the better one’s control of the temperature at the cutting surface, but do not set at less than 1mm without a good practice.


27

Fat Stains: See the section on Sudan III

Gelatin embedding for friable tissues:

• Wash out formalin with running water for 6-8 hours.

• Place washed block in a covered pot containing the following mixture for 4-6 hours at 37oC. Gelatin 15g Glycerine 15ml Distilled water 70ml Thymol A crystal

• Transfer block to a fresh lot of gelatin mixture in a small mould (e.g. paper) and allow to cool.

• Harden the block of gelatin in 10%-20% formol saline overnight. Trim off all excess gelatin and re-immerse block in formol saline for 8 hours (or till convenient).

Gum for freezing:

Gum arabic (ground to powder) 50g Cane sugar 20g Distilled water 50ml Thymol 0.05g

Mix and filter through muslin is necessary. RAPID FROZEN SECTIONS DURING AN OPERATION: • Select and trim a suitable thin block to tissue about

3mm thick. • Press on to a wetted microtome chuck. • Freeze rapidly with CO2 • Place on the microtome of the cryostat and cut

sections. • Pick up sections while still frozen on the side of the

knife with a coverslip. (They thaw and adhere). • Place on a warm plate (e.g. microscope lamp) till just

drying.


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• Stain the unfixed section for 1 minute in 1% polychrome methylene blue.

• Rinse in water till the depth of staining is right. • Rapidly dehydrate, clear and mount. • Place on a slide face down avoiding bubbles and

examine. (This unfixed section cannot be kept). Result:

Nuclei - Blue Cytoplasm -Very pale blue Fibrous tissue - Pink Elastic tissue - Unstained but refractile and greyish

Mast cells - Purple

Further sections can be stained and kept permanently as follows: • Dry section in air (NOT on warm plate) 15 seconds. • Fix in the following solution for 1 minute.

Formalin (commercial) 10ml Acetic acid 3ml Alcohol (Abs. Meth. Spirit) 87ml

• Rinse in tap water. • Celestine blue 1 minute. • Rinse. • Haemalum 1 minute. • Rinse. • Acid alcohol rapidly. • Scott’s tap water substitute until blue. • Rinse. • Rosin 5 seconds. • Rinse. • Dehydrate, clear and mount in D.P.X.


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SPECIAL STAINING METHODS

METHODS FOR CONNECTIVE TISSUE FIBERS: The proper recognition of connective fibers and their relationship to the other structures is often of vital importance in the diagnosis of lesions. Numerous methods are available and an understanding of their properties is necessary before the right one can be chosen. Collagen fibers: • Van –Gieson’s stain: is traditional and easy to apply but it

will only stain relatively coarse fibers and it readily fades. It is still useful for the study of coarse fibers such as leiomyoma, breasts and arteries (after elastic stains).

• Mallory’s Trichrome: (Picro-Mallory). This stains finer fibers than does Van Gieson and will pick out in blue the basement membranes and interstitial connective tissue in liver, kidney, heart, etc. Its greatest value in this respect is the staining of interstitial tissue of the glomerular tuft. In addition to its use for collagen it stains firbrin red. It can also be used to differentiate the cells of the pituitary.

• Mason’s Trichrome: This has similar properties to Mallory’s stain but stains connective tissue green. It is less useful for kidney but it is easier to use than Mallory’s. It will stain the striations of muscle.

• Silver Impregnation: This is undoubtedly the best method for demonstrating the finest fibrils and will distinguish them from coarser collagen by color (black and brown respectively). It is especially valuable for demonstrating the structure of tumors and lymph nodes and will also show the finest fibrils in early organization. This method is now reliable, rapid and applicable to nearly all fixatives and is rapidly replacing the earlier dyes. It cannot be used for glomerular basement membranes.

• Periodic acid Schiff: Carbohydrate in fine connective tissue is colored red. Used for glomerular basement membranes.


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• Period acid methenamine silver (Jones): This method demonstrates the connective tissue carbohydrate in black. The color is intense so sections must be very thin. Only used for glomerular basement membranes. (See under PAS methods).

Elastic fibers: Four methods are available:

• Verhoeff: This method is rapid easy and reliable. It fails to demonstrate the finest fibers. It can be easily combined with Van Gieson’s to give a very useful stain for vessels.

• Weigert: The stain is difficult to prepare and ready made commercial samples are unreliable. When good, the stain picks out the finest fibrils and is superior to the rest.

• Orecin: The natural dye is poor. The synthetic dye (G.T. Gurr) is a good stain. It is better than Verhoeff but not quite good as Weigert.

• Aldehyde fuchsin: A recent, reliable and easy method. Comparable to Weigert in sensitivity.

VAN GIESON’S STAIN: Use any fixative and paraffin sections.

1. Bring sections down to water and remove mercury if necessary.

2. Stain nuclei with an iron hematoxylin (e.g. Weigert) 15 minutes or with celestine blue followed by haemalum 10 minutes. Rinse and differentiate 10 min. Tendency should be to under-differentiate.

3. Wash in water. 4. Rinse in alcohol. 5. Stain for 1-3 minutes in Van Gieson’s

mixture. 6. Rinse quickly in distilled water or pour the

stain off. Blot off. 7. Dehydrate rapidly in absolute alcohol.


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8. Clear in xylol. 9. Mount in DPX.

Formula : 1% aqueous acid fuchsin 10ml Saturated aqueous solution of picric acid 90ml

• Dilute this with an equal volume of water and boil for 3 min. to ripen.

Results:

Nuclei - Black to dark brown Collagen - Red Other tissues - Yellow

Note: • The fuchsin is removed by water and the

picric acid by alcohol. It is often better to omit step 5 and dehydrate as quickly as possible. Alkaline water removes fuchsin very quickly.

• Picric acid decolorizes alum haematoxylin, hence a more resistant one must be used.

• In place of acid fuchsin in Van Gieson, one may use poncean or sirdus red. These are good for amyloid.

PICRO-MALLORY STAIN Any fixative. Fix paraffin sections on albuminized slides.

1. Stain nuclei with iron haematoxylin or celestine blue and haemalum.

2. Rinse in tap water. 3. Differentiate in picro-orange (1) until only nuclei are

stained (3-5 minutes). 4. Wash in water till only R.B.C. are yellow. 5. Stain in acid fuchsin mixture (2) till tissue is red (5-10

min.)


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6. Rinse in 2% acetic and differentiate in red differentiator (3) till only required elements are red (up to 5 min.).

7. Wash in tap water (10 seconds). 8. Stain in aniline blue (4) till connective tissue is well

stained (10 minutes). 9. Rinse in 2% acetic. 10. Differentiate in blue differentiator (5) till only

connective tissue is stained blue (not less than 1-2 minutes).

11. Dehydrate rapidly, clear and mount in D.P.X.

SOLUTIONS:

1. Picro-orange: 80% alcohol saturated with picric acid 100ml Orange G 0.25g

2. Acid- Fuchsin Mixture: Acid fuchsin 0.5g Ponceu Red 0.5g 1% acetic 100ml

3. Red differentiator: Stock differentiator* 40ml 95% alcohol 40ml Water 20ml

4. Aniline Blue solution: • Aniline blue dissolve in 100 ml

boiling distilled water. • Add 2.5 ml glacial acetic acid.

Cool and filter. 5. Blue differentiator: Stock differentiator* 20ml Water 80ml *Stock differentiator: Absolute alcohol 100ml


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MASSON’S TRICHROME STAIN: Any fixative. Paraffin sections.

1. Bring sections down to water. 2. Stain nuclei by either iron hematoxylin or celestine

blue and haemalum, rinse and decolorize. 3. Wash in tap water. 4. Stain for 1-5 minutes in Masson’s Ponceu-fuchsin. 5. Rinse in acid water*(tap water washes the dye out). 6. Treat with 4% aqueous phosphomolybdic acid till

collagen is not darker than pale pink. 7. Rinse in acid water. 8. Stain 2-5 minutes in Masson’s light green **(till

collagen is green). 9. Rinse in acid water.

10. Dehydrate, clear and mount. Results: Nuclei - blue-black Cytoplasm - light red Muscle - dark red Red cells - bright red Hyaline and fibrin - bright red Collagen and mucin - green *Ponceu-fuchsin: Ponceu 2 R 0.7g Acid fuchsin 0.35g Glacial acetic acid 1.0ml Distilled water 100ml **Light green: Light green 2g Glacial acetic acid 2 ml Distilled water 100ml

HEIDENHAINS MODIFICATION OF MALLORY’S ANILINE BLUE METHOD Any fixative. Paraffin sections.


34

(The following technique is taken directly from Mallory’s book) Staining solutions:

• Azocarmine Azocarmine B 0.25-1cm Distilled water 100ml Acetic acid, glacial 1cc

If azocarmine g is used instead of B add 0.1g to the 100ml water, bring it to a boil, cool to room temperature and filter through coarse filter paper in the paraffin oven at 51oC. After cooling add the 1cc of glacial acetic acid. • Aniline blue: Aniline blue 0.5g Orange G 2g Distilled water 100ml Acetic acid, glacial 8ml Boil and filter after cooling. For staining, dilute the stock solution 1:3 with distilled water. Method of staining: 1. Stain in the azocarmine solution in a glass-covered

dish in paraffin oven at 51oC to 55oC for 45-60 minutes, then cool at room temperature 5-10 minutes.

2. Wash in distilled water. 3. Differentiate in an alcoholic solution of aniline made

up as follows: Aniline 1cc Alcohol, 90% 1000cc.

4. Rinse in acetic acid alcohol made up as follows for 30 seconds to 1 minute:

Acetic acid, glacial 1cc Alcohol, 95 percent 100cc.

5. Mordant in 5 percent aqueous solution of phosphotungstic acid 1-3 hours.

6. Wash quickly in distilled water. 7. Stain in the aniline blue solution 1-3 hours. 8. Wash very quickly in water. 9. Differentiatee in 95% alcohol followed by absolute.

10. Clear in xylol and mount in balsam.


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Results: Collagen - deep blue Chromatin - red Muscle tissue - reddish to orange Erythrocytes - red Neuroglia - reddish Mucin - blue Pituitary alpha granules - red Pituitary beta granules - blue

GORDON AND SWEET’S SILVER IMPREGNATION OF RETICULUM This method can be used after all ordinary fixatives including Helly’s. 1. Cut paraffin sections and attach firmly to slides. 2. Take sections down to water (remove mercury if necessary). 3. Oxidize for 1-7 minutes in acid permanganete solution.

0.5% aqueous potassium permanganete 50ml 3.0% sulphuric acid 2.5ml

4. Wash in water. 5. Bleach until white in 1% oxalic acid or 10% hydrobromic

acid (about 1 minute). 6. Wash in two changes of glass distilled water. 7. Mordant for 2-15 minutes in 2% aqueous iron alum. 8. Wash in 2 to 3 changes of distilled water. 9. Impregnate for 5-7 second in Wilder’s silver bath.

10% silver nitrate 5cc - Add ammonia drop by drop until precipitate is not quite

dissolved. - Add 3% sodium hydroxide 5cc


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- Titrate with ammonia until not quite clear. - Make up to 50cc with double distilled water. - Filter into a bottle.

10. Wash briefly in distilled water. 11. Reduce in 10% aqueous formalin* in tap water (about ½

min.) 12. Wash in water. (If the sections are over-impregnated,

repeat the process from step 7). 13. Tone in 0.2% yellow gold chloride 1-3 minute (optional) 14. Wash in tap water. 15. Fix in 5^% sodium thiosulphate 5 minutes. 16. Wash well in tap water. 17. Dehydrate, clear and mount. Notes:

• All silver solutions should be made up in chemically clean glassware. The Wilder’s silver bath will keep for 3-6 months.

• If desired a light counterstain can be used between steps 16-17. Carmalum or neutral red-fuchsin are recommended.

• Steps 3-5 constitute “Mallory’s bleach”. It can be repeated; this is said to increase the sensitivity of the method.

*This formalin may be neutral or slightly alkaline but not acid.

WEIGERT’S ELASTIC STAIN The whole success of this method depends on preparing a good batch of stain. Endless formula have been published but much depends on the making of the batch and factors necessary for a good batch are not all known. A good batch keeps for about a year. We have found the following method works well:

Crystal-violet 2.5g Basic fuchsin 2.5g Dextrin 1.0g Resorcin 10.0g Distilled water 500ml 30% sol. Ferric chloride 62ml (B.I. anhydrous)


37

• Heat the distilled water to nearly boiling in a large evaporating basin. Mix the dyes and dextrin and dissolve in the hot water. Add the resorcin and bring to the boil. When boiling add slowly the freshly prepared ferric chloride solution, stirring continuously with a glass rod. It is important to keep the mixture boiling, though not too vigorously. Continue boiling and stirring for a further 2 minutes or so to coarsen the precipitate. Cool and filter through a Buchner funnel and filter flask attached to the section pump. Wash the deposit with distilled water until the drips are colorless and the bulk of the filtrate a clear azure blue. Usually 8-10 liters is sufficient. The filter paper is now removed and dried overnight in the incubator, when the deposit is scraped off and dissolved in 550ml of absolute ethyl alcohol, to which has been added 1 ml of conc. hydrochloric acid, by simmering on an electric hot plate or water bath for 30 minutes or so. Cool and filter, then add 19ml conc. hydrochloric acid and allow to stand 24-28 hour before use, when the color should be a dark greenish-blue.

Staining Solution:

Stock solution 35ml 70% alcohol 30ml (These amounts may have to be varied slightly with each fresh batch of stain).

Technique of Use: Any fixative. Paraffin sections. 1. Xylol 2. 2 changes of absolute alcohol. 3. Blot and dip in 0.5% colloidin (as this is a long

staining method) 4. Harden colloidin in 70% alcohol for 5 minutes. 5. Bring sections down to water. 6. Remove mercury (as described under routine

haemalum and eosin staining.


38

7. Treat with permanganete and oxalic acid as in silver reticulin method).

8. Wash in alcohol. 9. Rinse in alcohol. 10. Stain in Weigert’s stain for 8-24 hours (time depends

on the strength of the batch. 11. Wash in alcohol (74 Op or ordinary acid alcohol

until only elastic tissue is stained.) 12. Remove colloidin with ether-alcohol mixture or

acetone. 13. Wash in water. 14. Follow with full procedure for Van Gieson’s stain.

Result: Elastic fibers - dark blue-black VERHOEFF’S ELASTIC STAIN:

1. Bring sections down to water. 2. Stain in Verhoeff’s mixture* till black (about15 mins.) 3. Rinse in water. 4. Differentiate in 2% ferric chloride until only elastic

fibers and nuclei are stained. 5. Wash in water. 6. Rinse in distilled water. 7. Counterstain with Van Gieson ½ to 1 minute. 8. Rinse in distilled water, dehydrate, clear and mount.

*Verhoeff’s stain: 5% unripened alcoholic haematoxylin 10cc 10% aqueous ferric chloride 4cc Lugol’s iodine 4cc


39

(Neither the haematoxylin nor the ferric chloride should be old; mix in the order stated).

Result: Elastic tissue - black Nuclei - grey-black Collagen - red Other structures - yellow - This is an excellent, easy and reliable combination of stains.

Its only fault is the failure of very fine fibrils to stain but for most practical purposes it is perfectly adequate.

- Prolonged staining in Van Gieson removes the elastic stain. SYNTHETIC ORCEIN STAIN FOR ELASTIC TISSUE:

Synthetic orcein 1g Dissolve in 80% alcohol 100ml Add conc. HCl 1ml (The stain keeps well.)

1. Bring sections down to water and remove mercury if necessary.

2. Rinse in alcohol. 3. Stain in orcein for ½ to 1 hour (longer does no harm). 4. Rinse out excess stain with acid alcohol. 5. Wash well in water (this improves the contrast). 6. Counterstain with safranin, neutral red, or Van Gieson

(½ minute only) as desired. Result:

Coarse elastic fibers - reddish-brown Fine elastic fibers - dark-brown Other structures - counterstain only

ALDEHYDE FUCHSIN STAIN FOR ELASTIC TISSUE: Preparation of the stain:

• Add 1ml of conc. HCl and 1 ml of paraldehyde to 100ml of a 0.5% basic fuchsin in 60-70% alcohol. Keep at room temperature until the mixture darkens to a deep violet (about 24hours). The stain gradually alters its properties with age, staining more rapidly and strongly when fresh.


40

METHOD:

1. Bring sections to water. 2. Oxidize sections in Lugol’s iodine for 10 minutes to 1

hour. 3. Remove iodine with 5% thiosulphate for 1 minute. 4. Immerse in aldehyde-fuchsin for 5 minutes to 2 hours,

depending on the tissue component to be stained. 5. Rinse in 60-70% alcohol. 6. Counterstain if desired (orange G, light green, fast

green). 7. Dehydrate in alcohol. 8. Clear in xylene. 9. Mount in D.P.X.

Times of staining: Elastic tissue 5 minutes Pancreatic β-cells 15-30 mins. Pituitary β- granules 30 mins to 2 hours Mast cells granules 5-10 mins. Color: Deep purple

STAINING METHODS FOR DEMONSTRATING PARTICULAR CELLS:

There are certain cells in structure that are difficult to recognize or find and need special stains for their demonstration. The following methods are the ones that we have found most useful: Striped muscle: Lendrum’s “Phostox” or P.T.A.H. Smooth muscle: P.T.A.H. Lissamine fast red.

Mallory’s or Masson’s trichrome


41

Bone marrow cells: Giemsa Eosinophils: Phloxine Tartrazine Plasma cells: Pappenheim Mast cells: Thionine, toluidine, or

Polychrome methylene blue. Aldehyde fuchsin.

Pituitary cells: P.A.S. or Heidenhain’s Mallory Method

Enterochromaffin cells: Diazo method Bile canaliculi: P.T.A.H. Islets of Langerhans: Aldehyde fuchsin (β cells)

MALLORY’S PHOSPHOTUNGSTIC ACID HAEMATOXYLIN (P.T.A.H.) Any fixative. Paraffin sections. 1. Bring sections down to water. If necessary remove mercury

with iodine and remove iodine with alcohol (hypo. ruins the stain).

2. Postchromate* for ½ hour. Wash in water. 3. Differentiate the chromate for 1 minute in acid

permanganete. 4. Wash in water. 5. Treat with 1% oxalic acid till white. 6. Rinse in water and transfer to Mallory’s stain* for 12-24

hours. 7. Shake off excess stain (do not wash, water removes the red

component). 8. Dehydrate in alcohol (this differentiates the blue

component). 9. Clear and mount.

*SOL. A. 10% HCl in methylated spirit. SOL. B. 3% aqueous potassium dichromate. Mix 1 part of SOL. A. and 3 parts of SOL.B. and pour on slide.

*Mallory’s stain: Haematoxylin (or haematin) 0.1g Phosphotungstic acid 2.0g


42

Distilled water 100ml - Dissolve separately and mix. Leave to ripen for several

months. Artificial ripening has not proved satisfactory in our lands. When ripe, the stain keeps for several years.

Result: Fibrin and neuroglial fibrils and red cells- dark blue Nuclei - light blue Collagen - Rose red

Note: The balance of red and blue colour depends on the postchromating and the removal of the chromate by permanganete; the more chromate left in, the darker the blue. The times given above are average ones.

LENDRUM’S “PHOSTOX” Any fixative. Paraffin sections. 1. Bring down to water. 2. Mordant in Lugol’s iodine for 1 hour. 3. Pour off and without washing treat with permanganete and

oxalic acid, as above. 4. Wash in distilled water. 5. Stain in Mallory’s stain till muscle is stained blue (a few

hours). 6. Dehydrate, clear, and mount as above. PHLOXINE TARTRAZINE Fix in formalin or F.M., avoid chromate or Bouin. 1. Bring down to water and remove mercury if necessary. 2. Stain nuclei with haemalum. 3. Differentiate in acid alcohol. 4. Wash in tap water. 5. Stain for ½ hour in phloxine.


43

6. Rinse in water. 7. Treat with tartrazine** in cellosolve till the desired

components are red and the residue yellow. 8. Dehydrate in alcohol, clear and mount. - Water removes the tartrazine at once but it can be restained in a

few seconds so a rinse at stage 7 may be used to ascertain if staining is correct. A few seconds more in tartrazine restores the yellow background.

- Red phloxine is replaced by yellow tartrazine in the tissue

components in the following order and the process can be stopped at any desired point. • Collagen • Cell cytoplasm • Muscle • Erythrocytes • Fibrin • Mast cells and eosinophils • Nucleoli • Inclusion bodies.

- This is generally useful and easy to counterstain. *Phloxine solution: Phloxine 0.5g Calcium chloride 0.5g Distilled water 100ml **Tartrazine solution: Saturated solution of tartrazine N.S., in cellosolve (ethylene glycol monoethyl ether). LENDRUM’S LISSAMINE FAST RED FOR UNSTRIPED MUSCLE: Any fixative. Paraffin sections. 1. Bring section down to water and remove mercury if

necessary. 2. Stain 5 minutes in celestine blue mixture. 3. Rinse in water.


44

4. Stain in haemalum for 5-10 minutes, rinse in tap water and differentiate if necessary.

5. Blue in Scott’s tap water substitute for 3 minutes then wash. 6. Lissamine fast red for 5 minutes (1% in 1% acetic acid). 7. Immerse in 1% ohosphomolybdic acid, 5 minutes in 56oC

oven or in a bowl of hot tap water (the solution takes some time to warm up).

8. Rinse. 9. Tartrazine 5 minutes (1.5% in 1.5% acetic acid). 10. Brief rinse in 65% alcohol (optional). 11. Complete dehydration with absolute alcohol using

dropping bottle. 12. Clear xylol, mount in D.P.X. Result:

Nuclei - blue Muscle, red cells and some cell granules - red Background - yellow

Note: The Lissamine fast red solution goes off in a matter of weeks.

Prolonged staining does not seem to help. The times given are arbitrary and it may be desirable to use the phosphomolybdic acid cold if the red comes out too quickly. A little further red comes out in the tartrazine so do not push differentiation too far and if necessary shorten the time in tartrazine.

This method utilizes the basic technique of Mallory’s trichrome.

MAY GRUNWALD-GIEMSA STAIN ON PARAFFIN SECTIONS: Fix in Helly. Other fixative are not as good. Embed in paraffin and cut sections as soon as possible. 1. Bring section down to water and remove mercury.


45

2. Wash well in tap water for 5 minutes, when wash in two changes of buffered distilled water at pH 6.8.

3. Stain in a mixture of equal parts of May Grunwald solution and buffered (pH 6.8) distilled water for 1 hour.

4. Without washing, transfer to 1 in 20 dilution of Giemsa in buffered water for 2 hours.

5. Rinse in buffered distilled water. 6. Differntiate in glycerine-ether-alcohol mixture, controlling

under the microscope. This takes a few seconds only. Gurr’s glycerine-ether mixture diluted 1 in 4 with pure absolute ethyl alcohol is used and should be prepared fresh each time.

7. Dehydrate rapidly in absolute alcohol. 8. Clear in xylene. 9. Mount in Gurr’s neutral mountant. Result: • With fixation in Helly, the stain should be like ordinary

Romanowsky’s stain on films. Material fixed in formalin alone will not take up enough red stain. This may be partly corrected by adding a trace of eosin (alc. sol.) to the alcohol used for dehydrating.

Note: Buffer tablets at various pH can be bought G.T. Gurr. GIEMSA ALTERNATIVE (BARRET’S MODIFICATION) 1. Bring to water and stain nuclei lightly Mayer’s Haemalum

(3-5 minutes). 2. Differentiate and wash. 3. Rinse in distilled water.


46

4. Stain for 15-20 minutes in Orange mixture 0.5ml Buffer 0.5ml Acetone 1.0ml

5. Rinse in water and then in distilled water. 6. Stain for 18-24 hours in Orange mixture 0.15ml

Blue mixture 0.15ml Buffer 3.0 ml Distilled water 57.0ml

7. Blot, dehydrate quickly in alcohol. Clear and mount in D.P.X. Orange mixture: 1% Erythrocin 1 part 1% Orange G 2 parts Distilled water 2 parts Blue mixture: 1% methylene blue 2 parts 1% Toluidine blue 1 part Distilled water 17 parts Buffers: PH 5.8 pH 6.4 M/15 KH2O4 45ml 34ml M15 Na2HPO4 5ml 16ml

Note: The staining mixtures should be made up freshly from the stock solutions of 1% dyes. The two buffers can be tried because so tissues vary in their reactions. A satisfactory result can usually be obtained with one. METHYL GREEN-PYRONIN METHOD (PAPPENHEIM) This method offers a distinction between deoxyribonucleic acid (DNA) and ribonucleic acid (RNA) by making use of the high


47

affinity of the latter for pyronin and of the former for methyl green. It is important to avoid undue heating during processing. If a high temperature wax is used for embedding, or if sections are fixed to the slides on a warm tray at 85oC, alterations in the structure of both nucleic acids occur. These are most important and obvious in the case of DNA. RNA is readily lost by even early autolysis. Fresh tissue and good fixation are essential. Method: 1. Bring paraffin sections to water. Cryostat sections,

briefly post-fixed in 5% acetic-ethanol should be rinsed in distilled water.

2. Stain for 6 minutes in methyl green-pyronin*. 3. Blot gently with filter paper. 4. Dehydrate rapidly in absolute acetone. 5. Rinse in equal parts of acetone and xylene. 6. Rinse in 10% acetone in xylene. 7. Clear in two changes of clean xylene. 8. Mount in D.P.X. Result : DNA - green to bluish green (or purple) RNA - red *Preparation of the stain: • Make up a 2% aqueous solution of Pyronin Y

(not all samples will perform adequately). • Make up a 2% aqueous solution of methyl green and

extract it with chloroform in a separating funnel until no more violet enters the chloroform layer (this may take very many washes).

• For use, mix 12.5ml Pyronin Y and 7.5ml methyl green with 30ml distilled water. (Alternatively 30ml of M/5 acetate buffer of pH 4.8 may be employed).

HISTOCHEMICAL METHODS


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In histological diagnosis it is often necessary to find out the nature of various substances in the tissues. Their presence may be suggested by the appearance of the routine section or they may be known to occur in the type of lesion diagnosed. In any case, their presence should be tested and verified and not guessed. In the following pages methods are described for recognizing the substances most often encountered in diagnostic routine. FATTY SUBSTANCES: Ordinary paraffin sections are useless for the recognition of fats. Sometimes, as in liver, they suggest that fat is present but in most tissues they give no hint. It is therefore, essential to cut frozen sections if there is any question of fat being present. Many different fatty substances occur in disease and it is not possible to distinguish them individually in tissue sections. Nevertheless certain broad groupings are possible by utilizing several different tests. The tests used and the types of fat recognizable are as follows: Types of fat recognizable: 1. Neutral fats: (glycerides of fatty acids).

Occur in adipose tissue, in fatty infiltration and in degenerative lesions.

2. Free fatty acids: Occur in old inflammatory and degenerative lesions and in fat necrosis.

3. Cholesterol and its esters: Occur in atheroma, old hemorrhages, old inflammatory lesions, in sebaceous glands, adrenal, corpus luteum, testis.

4. “Myelin" fats A group of complex fatty substances which includes the phospholipids. Examples are normal myelin and Gaucher’s and Niemann Pick disease.

Tests:


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1. Stain frozen sections with Oil Red O or Sudan II & IV.

2. Reaction to polarized light: • Examine unstained frozen sections

mounted in water or jelly by a polarizing microscope or by an ordinary microscope with one “Polaroid” glass below the condenser and the other above the objective lens. Anosotropic substances show bright when the field is blacked out by rotating the glasses at right angles. Isotropic substances do not.

Note: There must not be any prism between the polaroids, i.e. no inclined unit or binocular unit.

3. Solubility: Most of the group of “myelin” fats are rendered insoluble by fixatives containing formalin. They then persist in paraffin sections after chloroform, xylene, etc. and can be stained by Sudan black (but not by Sudan II or IV or by Oil Red O).

Reactions of fats: 1. Neutral fats:

• Strongly positive staining with Sudan or Oil Red O.

• Isotropic • Totally soluble in fat solvents.

2. Fatty acid: • Weak staining with Sudan II &IV or Oil

red O variable, weaker than neutral fats. • Normally isotropic but may be anisotropic. • Totally soluble in fat solvents.

3. Cholesterols and its esters: • Cholesterol gives a weak Sudan or Oil red

O stain but the esters stain more strongly. • Anisotropic in cold but become isotropic

on heating and revert on cooling. • Totally soluble in fat solvents.

4. Myelin fats:


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• Stain with Sudan III or IV or with Oil red O.

• Anisotropic. • After formol fixation, they are insoluble in

fat solvents and give Sudan black staining in paraffin sections.

SUDAN STAIN FOR FATS These dyes, Sudan II & IV (Scharlach R) and also Oil red O, depend for their action upon their greater solubility in body fat than in the solvent used. A saturated solution of one dye will still dissolve the other and this is made use of in preparing the Sudan staining solution. The method is applicable only to frozen sections. It will work after most watery fixatives but formalin is preferable because the sections are less brittle to handle. 1. Cut frozen sections and receive into distilled water or diluted

formalin (cut sections can be stored in the latter). 2. Rinse sections in 70% alcohol. 3. Stain for not more than 1 minute in Sudan

solution* taking care to avoid letting the section fold over. (Folded sections stain unevenly- prolonged staining will dissolve out fine droplets of fat).

4. Rinse in 70% or 50% alcohol to remove excess stain.

5. Rinse in water holding section under till alcohol has diffused.

6. Counterstain for a few minutes in alum haematoxylin diluted 1 in 2 or 1 in 4 with distilled water (control depth of staining with microscope).

7. Differentiate if necessary in 0.5% HCl in 50% alcohol.

8. Wash well in distilled water to which a few drops of strong ammonia have been added.

9. Mount in glycerine jelly (see mountants). *Sudan stain:


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Mix equal quantities of dry Sudan III and Sudan IV powder and place them in a clean dry bottle. Fill the bottle with Herzheimer’s mixture (equal parts of acetone and 70% alcohol) and shake well. Leave solution for a few days to obtain a saturated solution. For use, pipette off some of the clear supernatant fluid; this is cleaner and more economical than filtering. When using keep vessels covered to avoid evaporation and precipitation of stain. Result: Fat - Orange-red Nuclei - Blue OIL RED STAIN FOR FAT Several dyes of the oil red series can be used as fat stains. The best in our hands has been oil red. It is used in a partly aqueous solvent which helps to prevent solution of small fat droplets during staining. Stock Oil red O:

• Make up a saturated (0.5%) solution of Oil red O in isopropyl alcohol.

• Stock staining solution: • For use, dilute 6ml with 4ml distilled water and

allow to stand for 24 hours. Decant the supernatant. This may be kept in a tightly stoppered bottle for up to 6 months, filtered as necessary through a No.42 Whatman paper, directly on to the sections.

Method: 1. Cut formalin-fixed sections or use unfixed cryostat

sections mounted on coverslips. 2. Rinse briefly in water. 3. Rinse in60% isopropyl alcohol. 4. Stain in Oil red solution for 10 minutes. 5. Differentiate briefly in 62% isopropyl alcohol*. 6. Wash in water. 7. Counterstain and mount as for Sudan stain avoiding

ammonia at stage 8.


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*Keep tightly stoppered. Absorbed moisture may dilute to below 60%. This will cause precipitation of Oil red.

The following alternative method is better for small quantities of fat:

Triethyl phosphate 60ml} “T.E.P.” Distilled water 40ml}

• Saturate some of this “T.E.P.” with oil red O at 56oC overnight. Cool and filter.

Method: 1. Rinse frozen sections in T.E.P. 2. stain in Oil red O in T.E.P. 10 minutes. 3. Rinse out excess stain in T.E.P. 4. Wash in distilled water. 5. Stain nuclei in haemalum. 6. Rinse in water. 7. Differentiate in T.E.P. containing 1% HCl. 8. Wash, blue, and mount in glycerine jelly.

(Fine fat droplets will not dissolve out in T.E.P. this being an aqueous solvent).

SUDAN BLACK B Sudan black B is a fat soluble dye like Sudan III and Sudan IV (Scharlach R) and will stain all the fats that can be stainedby them. A number of other lipid substances are rendered insoluble by formalin fixation and can be stained in ordinary paraffin sections by Sudan black (but not by Sudan III or IV). These lipids include red cell envelops, lipofuscin, kerasin (Gaucher’s disease) and myelin sheaths. For the latter, Sudan black is a rapid alternative to the slower Weigert-Pal. Sudan black is soluble in xylene and sections must be mounted in a watery medium (glycerine jelly or Aparhy’s medium).


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Method: 1. SOL. A. Sudan black B 1% in isopropyl

alocohol SOL. B. Sodium borate 1% aqueous - Mix equal parts of SOL. A. and SOL. B., stand for 10 minutes, filter.

2. Bring sections down to water. 3. Stain for 10 minutes. 4. Differentiate in 60% isopropyl alcohol in water. 5. Wash in distilled water. 6. Counterstain with a red nuclear dye (Carmalum is the best). 7. Wash in water. 8. Mount in glycerine jelly or Apathy’s medium. Result: Lipids stain black *Sudan black can also be used as saturated solution in 70%

alcohol and differentiated in 70% alcohol but staining takes much longer.

COPPER PATHALOCYANIN IN METHOD FOR PHOSPHOLIPIDS: Fix tissues in formalin or preferably, in formol-calcium. Embed in paraffin the usual way. Method: 1. Bring sections to absolute alcohol. 2. Stain in 0.1% Methanol Fast Blue 2G* in 90-100% alcohol

at 60o for 8-16 hours. 3. Rinse in 70% alcohol and bring to water. 4. Differentiate in 0.05% aqueous lithium carbonate for ½ to 2

hours. 5. Rinse in water. 6. Counterstain in 1% aqueous neutral red, up to 30 minutes

(the neutral red forms a deep blue complex with the phophalocyanin-phospholipid compound. This is very insoluble and, if preferred, staining with neutral red may be


54

done before stage 4 which subsequently needs only a very short period.

7. Rinse in water. 8. Blot dry and transfer rapidly through 70 and 100% alcohol

to xylene. 9. Mount in D.P.X. Result: Phospholipids - purplish blue (e.g. myelin sheaths and dark blue to red cell envelops) Nuclei and nucleoli - red Protein - pale sky blue or reddish shades N.B.:The presence of water in the alcoholic solution of

phthalocyanin increases the staining of non-lipid components in the tissues, but also the speed of staining of lipids.

MUCOID SUBSTANCES Mucin is not a chemical entity. The group of slimy fluids ordinarily spoken of as “mucus” comprises a large number of compounds which apart from their slimy physical character, all contain carbohydrate of some form in chemical combination with protein or aminoacid complexes and sometimes with sulphuric acid. This group contains substances of great diversity and no single stain will demonstrate all of them. The following list gives the most useful methods and their behavior with different mucins. Mucin-carmine P.A.S. Alcian green Toluidine blue Stomach ± or - + ± - Duodenal mucosa + + + or ± - Brunner’s glands - + - - Colon + ± + - Salivary gland + + ± - Pancreatic duct ± + + - Bile passages + + + or ± - Bronchus + + + ± Cervix + + + ± Endometrium - + ± - Bartholin’s gland + + + - Prostate ± + + - Seminal vesicles - + + - Ovarian cyst + + + -


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Cartilage + or ± + or ± + or ± + Umbilical cord ± ± ± + Aorta - - ± + Myxoma ± + or ± + or ± + Chordoma ± ± ± + SOUTHGATE’S NUCLARMINE STAIN Any fixative. Paraffin section. 1. Take down to distilled water. 2. Stain in Weigert’s iron haematoxylin 5 minutes of celestine

blue and haemalum. 3. Differentiate in 1% acid alcohol if necessary and blue again

in tap water. 4. Stain for 15-30 minutes in mucicarmine solution* diluted

1:5 with distilled water. 5. Wash in tap water. 6. Dehydrate, clear and mount. Result: Mucin - reddish Nuclei - blue Mucicarmine solution: Carmine 1g Aluminum hydroxide 1g • Add 100ml of 50% alcohol and then add 0.5g of

anhydrous aluminum chloride (beware of frothing and use a 500ml flask).

• Boil for exactly 2 ½ minutes. • Cool and filter. Note:Cheap carmines seem to work better than purified ones. The

mixture frothes when being prepared. PERIODIC ACID-SCHIFF TECHNIQUE (P.A.S.) This method depends on the production of aldehyde from 1:2 glycol groups present in carbohydrates by oxidation with periodic acid. The aldehydes are combined with Schiff’s reagent to form a red compound dye in situ.


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There are two different ways of using the technique, the first (McManus variant) uses a 1% aqueous periodic acid, the second (Hotchkiess variant) a 0.8% solution in buffered 70% alcohol, following by a reducing rinse. The McManus or watery P.A.S. gives the stronger reacting and should be used to demonstrate fungi amoebae, basement membranes etc. The Hotchkeiss or alcoholic P.A.S. gives a weaker reaction and should be used to demonstrate mucin and pituitary basophils. Both methods stain glycogen and if this is likely to be present it should be removed by 20 minutes incubation with saliva (or diastase) before staining. Strong P.A.S. (McManus) 1. Bring section to water and remove mercury. 2. Treat with 1% aqueous periodic acid 5 minutes. 3. Wash briefly in water (diluted). 4. Schiff’s solution 20 minutes. 5. Wash in water. 6. Stain nuclei with haemalum. 7. Differentiate and blue in the usual way. 8. Dehydrate, clear and mount. Result: Basement membranes of kidney and other organs - deep red Most fungi - deep red Connective tissues - reddish pink Hotchkiss Method: 1. Bring section down to water and remove mercury. 2. Rinse in 70% alcohol or meth. Spirit. 3. Immerse in Periodic acid solution at room temperature. 4. Rinse in 70% alcohol or meth. Spirit. 5. Immerse in reducing solution* for 1 minute.


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6. Rinse in 70% alcohol or meth. Spirit. 7. Immerse in Schiff’s solution** for 4 minutes or more. 8. Wash in running water for 10 minutes. 9. Stain nuclei lightly with celestine blue and haemalu (about

2-3 minutes in each). Differentiate and blue as usual. 10. Counterstain in Orange G*** for 10 seconds (optional). 11. Wash in water till pale yellow (about 80 seconds). 12. Dehydrate, clear and mount in D.P.X. Result: Mucin - red Nuclei - blue Background - yellow R.B.C.’s and aci - yellow Period Acid Solution: Periodic acid 0.4g M/5 sodium acetate 5.0ml

(M/S=27.2g hydrated salt in 1000ml) Abs. Ethyl alcohol 35.0 ml Distilled water 10.0ml (Keep between 17-22oC in dark. Keeps about 14 days). *Reducing bath:

Potassium iodine 1.0g Sodium thiosulphate pentahydrate 1.0g Abs. Ethyl alcohol 30.0ml Distilled water 20.0ml 2N Hydrochloric acid 2.5ml (use 20% of conc. HCl)

(Ignore any sulphur deposit. Keep between 17-22oC. Lasts up to 14 days.)

**Schiff’s reagent:


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• Basic fuchsin 0.5ml • Dissolve in boiling distilled water 100ml • And cool to 55oC and add N/HCl 10ml • Cool at temperature • and add sodium meth. bisulphate 0.75g • Leave 48 hrs. in dark at 4oC

(in tightly stoppered flask) • Shake with granulated charcoal

for 1 minute to remove pink color. 2g • Filter • Store in tightly stoppered flask at 4oC

(Lasts about 6 weeks at 4oC) *** Orange G Orange G (G.I. 27) 2.0g Phosphotungstic acid 5% Aqueous 100ml Stand for 24 hours and use supernatant.

PERIODIC ACID METHENAMINE SILVER METHOD (Jones)

This method is based on the periodic acid-Schiff but uses methenamine silver to detect the aldehyde. The staining is extremely intense- too intense for ordinary 5-6μ sections. It s practical use is the demonstration of glomerular basement membranes but it is the only really satisfactory on sections 2 μ thick. Method: Any fixative. Paraffin sections (2 μ).

1. Sections to water and remove mercury if necessary. 2. Rinse well in distilled water. 3. Oxidize in 1% aqueous periodic acid 15-20 minutes. 4. Rinse in distilled water. 5. Wash well in tap water for 15 minutes. 6. Rinse in distilled water. 7. Place in hexamine silver solution* for 1-2 hours.

Examine the section at ½ hourly intervals after the first hour. When the glomerular basement membranes is sharply defined proceed as follows:


59

8. Rinse well in distilled water. 9. Tone in 0.2% gold chloride for 5 mins. 10. Rinse well in distilled water. 11. Fix in 5% hypo for 2-3 mins. 12. Wash well in tap water. 13. Counterstain section by haematoxylin and eosin.

Result: Nuclei - black Basement membrane - black Cytoplasm and muscle - pink-red *Working Hexamine silver solution:

• Add 2ml of 5% borax to 25 ml distilled water. Mix well.

• Then add 25ml of stock hexamine silver solution.

Stock hexamine silver solution: 5% aqueous silver nitrate 5ml 3% aqueous hexamine 100ml (Hexamethylene-tetramine)

ALCIAN GREEN (2 G%) Method:

1. Bring paraffin sections to water. 2. Treat with 1% acetic acid for 2 minutes. 3. Stain in 1% alcian green in 1% acetic acid for 30

mins.. 4. Wash in running water. 5. Counterstain nuclei as required, with carmalum or

haematoxylin. 6. Dehydrate in alcohols. 7. Clear in xylene. 8. Mount in D.P.X.

Result: Acid mucopolysaccharide stain green. *If any procedure involving treatment with acids is to follow this stage, it is advisable to immerse the sections first in 0.3% sodium carbonate for 30 minutes. This converts the bound alcian green into an insoluble pigment.


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METACHROMASIA This is defined as the staining of a tissue component so that the absorption spectrums of the resulting tissue-dye complex differed sufficiently from that of the original dye, add from its ordinary tissue complexes, to give a marked contrast in colour. For most purposes it is limited to the effects seen with the thiazine dyes (Toluidine blue, Azure A, Methyl blue, Thionin). With the first of those a bright red change is called gamma metachromasia, and a change to purple- beta chromasia. Gamma metachromasia signifies the presence of free electronegative surface changes of a certain minimum density. These are most commonly attributable to the SO3H groups of sulphated mucopolysaccharides. Method: Fixation has a pronounced effect on the intensity of metachromasia. Paraffin sections can be used, cryostat sections postfixed if necessary, are preferable.

1. Bring section to distilled water. 2. Stain in 0.1% Toluidine blue in 30% ethanol for 5-20

mins. 3. Rinse in 95% alcohol. 4. Dehydrate in absolute alcohol. 5. Clear in xylene. 6. Mount in D.P.X.

Result: • Sulphate containing mucopolysaccharides or lipids

will show gamma (red) metachromasia. • Phosphate esters (nucleic acids) may show beta

(purple) metachromasia. • An orange red filter may be used to change the red

blue sequence to green red. This is more easily distinguished.

CARBOHYDRATE SUBSTANCES

GLYCOGEN (BEST’S METHOD) Glycogen is water soluble but is largely retained in tissues by being trapped in the proteins by ordinary fixation. To detect small


61

quantities, special fixatives should be used- Alcohol carnoy or best ice cold Bouin. Glycogen is readily stained in P.A.S. but a second slide must be stained after 20 minutes incubation with saliva to show that the stainable substance is diastase labile. Best’s glycogen carmine method is specific. Method:

1. Fix in ice cold Bouin, dehydrate in alcohol. 2. Transfer to alcohol-ether (equal parts) 1-2 hours. 3. ½ or 1% celloidin in ether overnight (use “Nucol”

356A/9). 4. Shake off excess celloidin and harden block in

chloroform 1 minute. 5. Clear in benzene till translucent (½ to 2 hours). 6. Impregnate in paraffin wax – 3 changes in hours. 7. Cut sections, mount and bring down to water. 8. Stain in haemalin 5 minutes (do not prolong time). 9. Differentiate and rinse (do not wash). 10. Stain in Best’s stain* 15-30 minutes. 11. Without rinsing in water, differentiate** for 5-30

seconds and inspect under microscope. 12. Wash in 80% alcohol. 13. Dehydrate, clear and mount.

Result: Glycogen - red Best’s Carmine Stock solution: (Lasts 3 months in ice chest)

Carmine 2g} Boil gently for 5 mins. Cool Pot. Carbonate 1g} and filter. Add to filtrate Pot. Chloride 5g} 20ml of ammonia.

Aq. dest 60ml} *For staining: Stock solution carmine 15.0ml} Ammonia(.880) 12.5ml}Lasts 2-3 weeks Methyl alcohol 12.5ml} **Best’s differentiation: Abs. alcohol 8ml Methyl alcohol 4ml Aq. dest 10ml


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• If boiling period is insufficient the stain may be less fast (more easily removed by the differentiator at stage 11).

PIGMENTS

The following colored materials may be encountered in sections: • Haemosiderin

Golden to rusty brown granules. Gives positive iron reaction with Berl’s stain.

• Haematodin (Bilirubin) Reddish-brown crystalline deposits. Negative iron reaction. See bilirubin below.

• Formalin pigment Jet black granules present in stale tissues which in blood and fixed in formalin. Can be removed by alcoholic picric acid (Immerse slides for 2 hours).

• Melanin Variable color from very pale brown to nearly black negative iron reaction. Bleached out by acid permanganete. Gives positive Masson’s silver bestand positive Schmori’s stain.

• Carbon Totally insoluble in anything. Jet black. Gives no reaction with any test.

• Bilirubin

Greenish-brown granular masses. Negative iron reaction Gmelin’s and Stein’s tests.

• Lipofuscins Brown granules in parenchymous cells. Iron test may be positive (due to associated iron). Positive satin for fat in frozen sections. Stains by Schmorl’s method.

• Malaria pigment Resembles formalin pigment and carbon. Soluble in conc. sulphuric acid.

• Haemoglobin Peroxydase stains with leuco patent blue.


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PERL’S REACTION FOR IRON Any fixative but chrome salts better avoided.

1. Bring paraffin sections down to distilled water. 2. Place in slightly warmed mixture of equal parts of

20% hydrochloric acid and 20% potassium ferrocyanide 5minutes.

3. Rinse in distilled water. 4. Counterstain in 1% aqueous neutral red, or other red

dye. 5. Wash rapidly in tap water. 6. Dehydrate, clear and mount.

Result: Iron (ferric salts) - dark blue Tissue - red

• To intensify the staining treat the slide after step 3 with hydrogen peroxide (10vols.) for 5 seconds, then wash for 5-10 minutes in running water.

Note: To avoid false reactions use iron-free analytical reagents.

Make up the ferrocyanide reagent freshly each tome (the strength need not be very accurate).

GMELIN REACTION FOR BILIRUBIN AND HAEMATODIN Method:

1. Bring section to water. 2. Apply a 50/50 mixture of conc. nitric and absolute

alcohol. 3. Apply coverslip and wipe off excess reagent. 4. Ring edge with hot paraffin wax (optional). 5. Examine under the microscope.


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• A spectrum of colors from red to green spreads out from masses of bilirubin or from haematodin crystals.

STEIN’S METHOD FOR BILE PIGMENTS Method:

1. Bring sections to water. 2. Treat with Lugol’s iodine 2 parts, tinc. Iodine 1 part

mixed for 12-18 hours. 3. Wash in running water 5 minutes. 4. Treat with 5% aqueous sodium thiosulphate

30seconds. 5. Counterstain with Mayer’s carmalum 3-18 hours. 6. Wash in water, dehydrate, clear and mount.

Result: Bile pigment - dark greenish-black SCHMORL’S METHOD FOR MELANIN,LIPOFUSCINS: etc. Method:

1. Bring sections to water. 2. Immerse in ferric-ferricyanide solution* for 5-10

minutes, wash in 1% acetic acid and examine under microscope.

3. Repeat stage 2 if necessary, until melanin is dark blue but background is clear.

4. Wash thoroughly in running water. 5. Counterstain nuclei with 1% aqueous neutral red.. 6. Dehydrate rapidly, clear and mount in D.P.X.

Result: - Reducing substance - dark blue. (This include melanin enterochromaffin granules,

lipofuscin, and SH groups). - Nuclei - red

*Ferric-Ferricyanide solution 1% Ferric chloride 30ml}bothfreshly prepared 1% potassium ferricyanide 4ml } Distilled water 6ml Mix, fliter and use within 30 minutes.


65

MASSON’S SILVER METHOD FOR MELANIN The argentaffin cells of the intestine also gives this reaction. Note that this reaction differs fundamentally from silver impregnation method in that no reducer is used. Reticulin stains do not show up melanin because they are too rapid and because the oxidizers used in most of them bleach the melanin. MASSON’S METHOD: Fix in any ordinary fixative, avoiding chromates.

1. Routine paraffin sections. 2. Bring to distilled water. 3. Leave overnight in Fontana’s ammoniacal silver* in

the dark in a covered jar. 4. Rinse in distilled water. 5. Fix in 5% “hypo”,1-2 minutes. 6. Counterstain with carbol-safranin. 7. Dehydrate, clear and mount.

*Fontana’s silver solution: • To 20cc of 10% AgNO3, add strong ammonia

drop by drop till only a few granules of the first formed precipitate remain; if the mark is overshop, add more AgNO3 to faint palescence.

• Add 20cc of distilled water. Allow to settle for a day and decant into a dark bottle. Filter each jar full before use and do not use for more than a few sections. (Keeps in the bottle for a month or two).

• If it is intended to do any large number of sections, commercial buffered hexamine silver may be used. It gives less deposit and being nearly neutral does not loosen sections from the slide but we have found it less reliable.

1. Take 100ml of 3% hexamine. 2. Add 5ml of 5% silver nitrate. The

precipitate redissolves.


66

3. Add 5ml of borate buffer of approx. pH (to 3% boric acid add a little phenolphthalein, then NaOH till a pink color is just precipitable).

4. Make up to 200 with distilled water. LISON – DUNN TECHNIQUE FOR HEMOGLOBIN

1. Fix in neutral (preferably buffered) formalin for 24-48 hours but preferably not longer.

2. Cut paraffin sections and bring to water. 3. Stain in leuco patent blue* 5 minutes. 4. Rinse in water. 5. Counterstain with neutral red.** 6. Dehydrate, clear and ount. Result:

Haemoglobin - dark blue Oxidase granules - dark blue Nuclei - red *Leuco patent blue: 1% aqueous leuco patent blue 100ml Add powdered zinc 10g Glacial acetic acid 2ml

• Boil the mixture until it is pale straw color. • Cool and filter. This is the stock solution.

For use, make up: Stock solution 10ml Glacial acetic acid 2ml 10 vol. Hydrogen peroxide 1ml Filter before use. **1% neutral red in 1% aqueous acetic acid.

DIAZO METHOD FOR


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ENTEROCHROMAFFIN CELL GRANULES Fix tissues in any formalin containing fixative. Oxidizing agents such as dichromate and chromic acid, should be avoided if possible.

Method: 1. Bring sections to water. 2. Treat for 30 seconds with a dilute (1mg/ml) solution of

the stable diazotate of 5-nitro-anisidine (Fast red salt B, I.C.I. 1bd) also known as Echtrotsalz B, in 0.1 H voronal acetate buffer at pH 9.2.

3. Wash thoroughly in running water. 4. Satin nuclei with Mayer’s haemalum, 6-10 minutes. 5. Wash in running water for 30 minutes. 6. Dehydrate in alcohol, clear in xylene and mount in

D.P.X. Result:

Argentaffin coll granules - fiery orange red Nuclei - dark blue Cytoplasmic structures - yellow

Note: Any one of a large variety of stable diazotates can be

used most satisfactory results are obtained with those giving reddish azo dyes. The colors given with granules in carcinoid tumors are more usually brownish-red.

MISCELLANEOUS SUBSTANCES

FEULGEN REACTION FOR D.N.A. This reaction depends on the fact that aldehyde (potential aldehyde) groups are produced by hydrolysis or deoxyribonucleic acid in fixed tissues by N HCl at 60oC. After hydrolysis the sections are washed and transferred to Schiff’s solution which gives a red compound with aldehydes. The time of hydrolysis varies with the fixative (5-15 minutes for most fixatives).


68

Method: 1. Bring sections to water (remove mercury if necessary). 2. Rinse briefly in cold N HCl. 3. Incubate in N HCl (pre-heated) at 60oC for the

optimum hydrolysis time (find by experiment if necessary).

4. Rinse in col N HCl and then in distilled water. 5. Transfer to Schiff’s solution ½ - 1 hour. 6. Drain and rinse in three changes of freshly prepared

bisulphate solution (5ml 10% K2S2O5, 5 ml N HCl, water to 100ml)

7. Rinse in water. 8. Counterstain (1% aqueous light green or fast green). 9. Dehydrate, clear and mount in D.P.X.

STAINS FOR FIBRIN Fibrin stains red with picro-Mallory or Masson’s trichrome but these are not specific. It stains dark blue with Mallory’s phosphotungstic acid and this is the most reliable method. It can be stained blue black by a modified Gram stain(below) but this method depends on proper differentiation and is, therefore, not completely reliable. Method: Any fixative, paraffin sections.

1. Bring to water. 2. Stain lightly with carmalum, eosin and carbol safranin. 3. Wash. 4. 1% aq. sol. Aniline methyl violet * 3-4 minutes. 5. Wash in tap water. 6. Lugol’s iodine 3-4 minutes. 7. Wash, Blot well. 8. Decolorize and dehydrate in aniline xylol** until

cytoplasm is practically colorless and fibrin dark-blue. 9. Blot very thoroughly. 10. Wash well in xylol and mount. Result: Fibrin - dark-blue

Nuclei - reddish-brown


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(Gram Positive organisms are also stained by this method) *Aniline methyl violet Solution A: Methyl violet 2.5g } Mix and filter. 95% alcohol 12cc } Solution B: Aniline water 100cc Aniline water: Aniline 2cc }Shake well and filter. Aq. distilled 98cc } **Aniline xylol: Aniline 1 part Xylol 2 parts AMYLOID Amyloid stains metachromatically with various violet dyes. It stains red with congo red and the stained material is tropic. It stains with Thioflavian T and the stained product fluoresces in ultraviolet light. It stains non-specifically with PAS (McManus) METACHROMASIA WITH METHYL VIOLET

1. Paraffin sections take down to water. 2. Stain in 1% aqueous methyl violet (or crystal violet) 5

minutes, or better, stain for 1 hour or more in a very dilute solution (a few drops of 1% solution in 50ml of distilled water).

3. Differentiate in dilute acetic acid (4 drops of glacial in 100ml of water) or 0.5-1.0% oxalic acid until amyloid is pink and other tissues violet.

4. Wash well in distilled water to remove acid. 5. Stand section for as long as feasible (more than 1

hour). 6. Without rinsing, in Apathy’s gum acacia mountant. Result:

Amyloid - pink Background - blue to violet


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A more sensitive stain can be obtained with cryostat sections of fresh tissue: 1. Fix cryostat section in 10% formol saline 10 mins. 2. Wash in water. 3. Stain in 1% methyl violet 5 minutes. 4. Wash in water. 5. Part differentiate in 1% acetic acid. 6. Wash in water. 7. Complete differentiation and counterstain in2%

aqueous methyl green (chloroform washed). 8. Wash well in water. 9. Mount in glycerine jelly or blot dry, rinse in xylol and

mount in D.P.X. Result: Amyloid - red

Nuclei - green Mast cells - blue Background - clear

HIGHMAN’S MODIFICATION OF BENNHOLD’S CONGO RED METHOD

1. Stain in Congo red (0.5% in 50% alcohol) for 1-5 minutes.

2. Differentiate in 0.2% potassium hydroxide in 80% alcohol for 1-3 minutes.

3. Rinse in water. 4. Counterstain with alum haematoxylin. 5. Wash inwater. 6. Dehydrate, clear and mount in D.P.X. Result: Amyloid - red

Nuclei - blue


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THIOFLAVIN T STAIN This is probably the most sensitive method.

1. Any fixative. Paraffin sections. 2. Bring to water. 3. Stain in haemalum if desired. 4. Rinse in distilled water. 5. Stain in Thioflavin T* 3-5 minutes. 6. Wash off with 1% aqueous acetic acid. 7. Differentiate in 1% acetic acid 10-20 minutes. 8. Rinse in water. 9. Mount in Apathy’s medium. 10. Examine under ultraviolet light using exciter filter

BG 12 and barrier filter O.G.4 or O.G. 5 or alternatively with exciter filter U.G.I. or U.G.2 with a colorless U.V. barrier filter.

Result: Amyloid fluoresces yellow *Thioflavin T 1% aqueous solution. Store in cold room (4oC)

and filter before use. CALCIUM Calcium deposits stain a dark blue black with haematoxylin but the staining is due to trace of iron in the deposit and is not specific. Von Kossa’s method stains calcium black but here the reaction is due to the acid radical of the calcium deposits and is also not strictly specific. It is, however, a very useful method. Alizarin is a specific stain. VON KOSSA’S STAIN

1. Any fixative that does not contain free acid. 2. Bring paraffin sections down to distilled water. 3. Place in 1.5% aqueous silver nitrate (freshly prepared

from a 10% stock solution) and leave in the light for 1 hour.

4. Wash in distilled water.


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5. Remove excess silver the “hypo”. 6. Counterstain in 1% neutral red, or carbol safranin. 7. Rinse in tap water. 8. Dehydrate, clear, and mount.

ALIZARIN RED’S 1. Fixation as above. Paraffin sections. 2. Bring to water. 3. Stain in Alizarin red S solution* 1-5 mins. 4. Rinse rapidly in distilled water. 5. Blot section dry. 6. Counterstain in 0.5% toluidine blue 5-10 seconds. 7. Rinse rapidly in distilled water. 8. Blot. 9. Dehydrate in two changes of acetone. 10. Mount in D.P.X. Result: Calcium - orange red Nuclei - blue

*We use Revector Alizarin red S (Sodium Alizarin sulphonate 5800S) 2% aqueous Alizarin red

• Add 10% ammonia (880 diluted 1 in 10) drop by drop until the solution turns deep iodine color.

• Allow to stand for 5 minutes. • Filter.


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METHODS FOR ENZYMES THE CALCIUM COBALT METHOD FOR ALKALINE PHOSPHATASE (after Gomori) PARAFFIN SECTIONS: Thin blocks of tissue should be fixed, according to Gomori’s method (1946) in two or three changes of cold absolute acetone at 4oC for 24 hours. There are many subsequent ways in which the tissues may be embedded in paraffin wax, most of them being minor variations designed to minimize destruction of the enzyme. As a routine procedure the following method gives good results.

1. Transfer the blocks progressively at ½ hourly intervals to absolute ethanol, absolute ethanol-ether with one or two changes, and then to 1% celloidin.

2. Drain off excess celloidin and harden in chloroform. 3. Clear in benzene. 4. Embed in paraffin wax, avoiding prolonged exposure

to the high temperature of the wax bath. 5. Cut sections at 5μ and mount on albuminized slides. 6. Dry the slides for an hour at 37oC. 7. Store at 4oC until required for incubation.


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METHOD PROPER AND CRYOSTAT SECTIONS: (After fixation from step 3)

1. Remove wax from the slides by brief immersion in right petroleum.

2. Pass tap water via absolute acetone. 3. Incubate for ½ hr. to 16 hrs. at 37oC in the following

medium: 10ml 2% sodium B-glucorophosphate 10ml 2% sodium diethyl barbiturate 20ml distilled water 2ml 2% calcium chloride 1ml 2% magnesium sulphate

4. Rinse in running water. 5. Treat with 2% cobalt nitrate or acetate 3-5 minutes. 6. Rinse well in distilled water. 7. Treat with a dilute solution of yellow ammonium

sulphide 1-2 minutes. 8. Wash in water, counterstain in 1% eosin, 5 minutes if

desired. 9. Dehydrate in alcohol, clear in xylene and mount in

canada balsam. Result: Various structures are stained black or brownish-black

in tissue processing alkaline phosphatase activity. FROZEN SECTIONS:

1. Cut sections 10-15μ thick and mount on clean glass slides without any adhesive.

2. Dry in air at room temperature for 1-2 hours. 3. Incubate in the substrate solution for ½-4 hrs. 4. Wash in water treat with 2% cobalt solutions, wash

treat with dilute yellow ammonium sulphide. 5. Counterstain in 1% aqueous eosin 5 minutes 6. Wash in running water 5 mins.


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7. Mount in glycocino. THE LEAD-NITRATE METHOD FOR ACID PHOSPHATASE Suitable material: Cryostat, post-fixed and paraffin sections.

1. Paraffin and post fixed sections to water. 2. Incubate at 37oC for ½-16 hours, 1-2 hours for cryostat

sections; longer for others) in 0.01M sodium b-glycerophosphate in 0.05M acetate buffer (pH 5.0) containing 0.004M lead nitrate.

3. Wash briefly in water. 4. Immerse in 1-5% yellow ammonium sulphide 1-2

minutes. 5. Wash in water. 6. Counterstain if required in eosin. 7. Wash well and mount in glycerine jelly.

Result: Acid phosphatase activity is shown by a black precipitate of lead sulphide.

COUPLING AZO DYE METHOD FOR ALKALINE PHOSPHATASE Method:

1. Fix with thin slices of tissue in 10% neutral formalin at 4oC for 10-16 hours.

2. Cut frozen sections 10-15μ thick and mount on clean slides without adhesive.

3. Allow to dry in air for 1-3 hrs. to ensure adherence. 4. Dissolve 10-20mg sodium a-naphthyl phosphate in

20ml 0.1M veronal acetate buffer (pH 9.2). Add 20mg of the stable diazotate of 4-benzoyl amino-2: 5-dimethoxyaniline and stir well. Filter on to the slides sufficient to cover each section adequately and incubate at room temperature. (17-22o) for 15-60 minutes. (Alternatively use the same quantity of the stable diazotate of 4-chloro-o-anisidine; or of 5-chloro-o-toluidine, and proceed in the same manner.


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5. Wash in running water for 1-3 minutes. 6. Counterstain in Mayer’s haemalum, 4-6 minutes. 7. Wash in running water for 30-60 minutes. 8. Mount in glycerine jelly.

Result: The sites of alkaline phosphatase activity are colored black with salt 2 or brick red with salts 7 and 9;Nuclei- dark blue.

PARAFFIN SECTIONS: Cold acetone-fixed, paraffin-embedded, or cold formalin-fixed paraffin-embedded as given in method for esterase. Method:

1. Bring sections to water via absolute acetone after removing the paraffin wax with light petroleum.

2. Cover with freshly made and filtered substrate-diazonium salt mixture as above.

3. Incubate for 30 minutes to 4 hours (Salt 2), or for up to 2 hours (salt 7), or up to 12 hours (salt 9).

4. Wash inwater, counterstain as above and blue in running water.

5. Mount in glycerine jelly. • (The use of salt 9 is particularly

recommended). Result:

With salt 9, the sites of alkaline phosphatase activity appear dark reddish-brown localization is excellent.

Nuclei - blue


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STANDARD COUPLING AZO DYE METHOD FOR ACID PHOSPHATASE Suitable material: cryostat sections postfixed, formol-fixed frozen sections or freeze dried sections. Method:

1. All sections to water. 2. Incubate at 37oC in the following solution.

• 10-20mg sodium a-naphthyl phosphate in 20ml 0.1 veronal acetate buffer (pH 5.0). Add 20mg fast garnet GBC (salt 18). The solution is mixed and filtered on the sections. Incubation times vary from 1-45 minutes.

3. Wash well in running water. 4. Counterstain if required in haematoxylin. 5. Wash in water. 6. Mount in glycerine jelly.

NAPHTHOL AS – PHOSPHATE METHOD FOR ALKALINE PHOSPHATASE Material: Post fixed cryostat sections, pre-fixed frozen sections,

freeze dried sections, paraffin sections. Stock solution:

Dissolve 25mg naphthol AS-MX (or B1) in 10ml N,N-dimethyl formamide, adding 10ml water and sufficient molar sodium carbonate (2-3 drops) to bring the pH to 8.0. After the addition of 300ml water the volume is brought up to 500ml by


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adding 0.2M tris buffer (pH 8.3). The slightly opalescent solution is stable at room temperature for several months.

Method: 1. Sections to water. 2. Incubate in stock solution containing 1mg/ml fast red

T.R. (salt 9) for 10-30 minutes at 22oC. 3. Wash briefly. 4. Counterstain if required. 5. Wash well. 6. Mount in glycerine jelly.

Result: Alkaline phosphatase activity - red

NARHTHOL AS – PHOSPHATE METHOD FOR ACID PHOSPHATASE

Material: Post fixed cryostat sections, pre-fixed frozen sections, freeze dried sections.

Solutions:

A. Pararosanilin HCl. Stock solution. B. 4% sodium nitrite in distilled water. C. Veronal-acetate stock solution

buffer pH 9.2 D. Dissolve 100mg Naphthol AS-TR

(or B.I.), phosphate in 10ml N,N-dimethyl formamide.

Incubating medium:

Dilute 5ml of solution C with 12ml distilled water and add 1ml of solution D. Mix 0.8ml each of solutions A and B and add to the solutions. Adjust the pH of the whole solution to pH 5.0 with N NaOH.

Method:

1. Sections to water. 2. Allow to dry. 3. Place in incubating medium for 30-90minutes at room

temperature.


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4. Wash in water. 5. Counterstain in methyl green if required. 6. Wash in water. 7. Mount in glycerine jelly or D.C.M.

Result: Acid phosphatase activity - red THE a-NAPHTHOL ACETATE METHOD FOR ESTERASE Method:

1. Use either cold-formalin fixed frozen sections, without washing out the formalin, or paraffin sections fixed in cold acetone or in cold-formalin, and dehydrated with cold acetone.

2. Incubate for 1-15 minutes at room temperature in the following medium: Dissolve 20mg a-naphthol acetate in 0.25ml acetone and add 20ml 0.1M phosphate buffer (pH 7.4)*.

3. Wash in running water 2 minutes. 4. Counterstain in Mayer’s haemalum 4-6 minutes. 5. Wash in running water for at least 30 minutes. 6. Mount in glycerine jelly.

*Alternatively, add 0.2ml 1% a-naphthol acetate in acetone to 10

ml of the buffer. Result: Esterase - black Nuclei - dark-blue


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METHODS FOR MICRO-ORGANISMS IN SECTIONS The routine method is Gram’s stain. This is admirable for Gram positive organisms but not wholly satisfactory for Gram negative ones. GRAM’S STAIN:

1. Any fixative, paraffin sections. 2. Bring down to water. 3. Stain for 1 minute in Gram’s crystal violet*. 4. Rinse in water. 5. Mordant in Lugol’s iodine ½ minute. 6. Rinse. 7. Differentiate in acetone till no more clouds of stain

come out (about 3 seconds). 8. Rinse in water 9. Counterstain in neutral red-fuchsin. 10. Dehydrate quickly in alcohol (this takes some of the

counterstain out). 11. Clear and mount in D.P.X.

*Crystal violet:


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• Dissolve 2mg of crystal violet in 20ml of 96% alcohol. Add 80ml of 1% aqueous ammonium oxalate.

As an alternative method to this use the modified Gram’s method given as fibrin stain.

If Gram negative organisms fail to stain well, stain an additional section with simple methylene blue. This shows all organisms well but does not distinguish between them. (Giemsa’s stain also stains organisms blue.

GROCOTT HEXAMINE SILVER METHOD FOR FUNGI Stock hexamine silver solution: 5% aqueous silver nitrate 5ml 3% aqueous hexamine 100ml (hemamethylenetetramine)

• Mix; a white precipitate forms but dissolves on shaking. (Store at 4oC, keeps for months).

Working silver solution: 5% aqueous Borax 2ml }Mix and add Distilled water 25ml} Stock hexamine silver solution 25ml Light green counterstain: Light green 0.2g } Distilled water 100ml}Mix and dilute Glacial acetic acid 0.2ml }1 in 5 for use Method:


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1. Hydrate section. 2. Oxidize in 5% aqueous chromic acid 1

hour. 3. Rinse in tap water. 4. Treat with 1% aqueous sodium

metabisulfite 1 minute. 5. Wash in tap water 5-10 minutes. 6. Rinse in 3-4 changes of distilled water. 7. Place in working silver bath in paraffin

oven (56oC) until section is yellow to golden brown (about 30-45 min).

8. Remove with wax coated forceps, rinse in distilled water and examine. Fungi- brown to black.

9. Rinse in 3 changes of distilled water. 10. Tone in0.1% gold chloride 2-5 minutes. 11. Rinse in distilled water. 12. Fix in 2% sodium thiosulphate. 13. Wash well in tap water. 14. Counterstain in light green solution ½ to 1

minute. 15. Rinse, dehydrate, clear and mount.

Result: Fungi, mucin, melanin, glycogen - black Background - green ZIEHL-NEELSEN CARBOL FUCHSIN STAIN (For tubercle bacilli in tissue sections). This stain is merely a slight modification of that ordinarily employed on bacteriological smears. Any fixative may be used, but mercurial fixatives are best. 1. Bring paraffin sections down to water. 2. Flood slide with carbol fuchsin* and heat to

steaming (not boiling). Keep hot for 15 minutes.

3. Rinse in distilled water.


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4. Differentiate in acid alcohol until the section is a faint pink when washed in water and the red cells are still pink.

5. Wash in running water for 5 minutes. 6. Stain lightly in haemalum or in methylene

blue. 7. Dehydrate, clear, and mount.

*Carbol fuchsin: Basic fuchsin 10g Phenol crystals 50g Alcohol 100ml Distilled water 1000ml • It is important that the counterstain should not

be too dark, otherwise bacilli are difficult to find.

WADE FITE METHOD FOR MYCO. LEPRAE IN SECTIONS 1. Warm section to melt wax and blot. 2. Remove rest of wax in

Peanut oil 1 part Xylene 2 parts

3. Blot dry. 4. When dry, place in water (avoid alcohol).


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5. Remove mercury if necessary. 6. Stain 30 minutes in carbol-fuchsin at room

temperature. 7. Wash in tap water. 8. Decolorize for 2 minutes only in 0.5% HCl in

70% alcohol. 9. Wash well. 10. Counterstain nuclei in Mayer’s haemalum, 5

minutes. 11. Blue in tap water. 12. Blot and dry (avoid alcohol). 13. Clear in xylene. Mount. Result: Myco. leprae - red Nuclei - blue

SILVER STAIN FOR SPIROCHAETES The method depends on the impregnation of the spirochaetes with silver nitrate and its subsequent reduction to an insoluble black oxide. There are several modifications of Levaditi’s original


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technique. The present one has given the best results in our hands. The tissue is stained on block.

1. Fix in formal saline (other fixatives will not work).

(Avoid any impurity of mercury). 2. Cut thin slices of tissues, 2-3mm thick. 3. Absolute alcohol, 24 hours. 4. Wash in aq. dist. Until block sinks (several changes). 5. Place in 100ml of silver bath* 24 hours at 37oC. 6. Add 0.25ml of glacial acetic acid to the 100ml of silver bath

and leave for four more days at 37oC. 7. Wash well (2-3 hours) in aq. dist. (several changes). 8. Place in reducing fluid** at room temperature for 2-3 days. 9. Wash 2-3 hours in aq. dist. (several changes). 10. Dehydrate in alcohols, clear, embed, cut sections and

mount on slides. 11. Remove wax with xylol and mount in balsam carefully; do

not press the coverslip as the section is brittle and will disintegrate.

Result: Spirochaetes - black Nuclei - Greyish Background - yellow

Silver bath: Silver nitrate 1.5g Double distilled 50ml Water Abs. alcohol 50ml

**Reducing fluid: Tanin 3g } Gellic acid 5g }Solution should be at Fused sodium acetate 10g }least 3 months old. Aq. dist. 350ml} FLUORESCENT MICROSCOPY (For tubercle bacilli in tissue sections) Use a monocular microscope with normal objectives and eyepieces. The condenser should be of the simplest (Abbe) type


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unless a quartz one is available. The high dry obhective (1/6th) is usually sufficient – if an immersion lens is required, a nonfluorescent medium such as glycerol can be used, ordinary immersion oil causes no trouble, however, ultraviolet light, filtered through wood glass and a layer of aqueous copper sulphate* passes through the specimen and into the microscope. In the ocular, a UV absorbing filter (Ilford minus blue, Wratten 2B) is placed to prevent ultraviolet light from reaching the eye. Method:

1. Bring paraffin sections down to water. 2. Stain at 60oC for 10 minutes (on rack over water bath)

in Carbol –Auramine-Rhodamine.** 3. Wash in tap water. 4. Decolorize in 0.5% HCl in 70% ethanol for 1-2

minutes. 5. Wash in tap water. 6. Mount in 80% glycerol for viewing. If permanent mounting is required, dry the slide at 56oC for 2 hours and mount in D.P.X. via dilute D.P.X. in xylene.

Result: Acid fast bacilli - bright reddish-gold rods which fluoresce green

*Aqueous copper sulphate: Hydrated CuSO4 8g Conc. Ammonia (S.G. 0.88) 100ml Aq. dest. 160ml

**Carbol-Auramine-Rhodamine Auramine O 1.5g Glycerol 75ml RhodamineB 0.75g Phenol liquefactum 10ml Aq. dest. 50ml

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His to Pathology Department