Embedding tissues in resin meets the requirement of tissue support for semi-thin sections with improved morphology. These agents are also readily available, relatively inexpensive, easy to prepare and handle, of sufficiently low viscosity to allow for short infiltration times and consistently thin sections to be cut and enable a range of staining methods to be used. Ideally resins should also demonstrate stability, uniform polymerisation (preferably at room temperature) and water solubility; they should be degradable, non toxic and produce minimal artefacts. Unfortunately none of the resin media currently available meet all of these demands.⁵ Excessive shrinkage in particular, is a common artefact that is more pronounced with methacrylates than with epoxy resins.

Processing
Tissue processing is often performed manually. Suitable automatic processors are commercially available but the expense of purchase is not warranted for the low number of specimens normally embedded in resin.

Glass or polypropylene vessels are suitable for manual processing with a wide range of resins. Infiltration is improved with agitation (mechanical stirrer or haematological rotator).

A paper label, written with indelible ink or pencil, should be included with the specimen during all stages of processing and incorporated into the final block as a permanent record. Computer generated labels produced on printers with carbon ribbons are also suitable.

Fixation
Standard fixation schedules with a wide range of fixatives are compatible with resin techniques. The commonly used formalin and mercury based solutions are particularly effective for providing sharp cytological and nuclear detail.

Dehydration
Dehydration is not necessary with all resin embedding protocols. When required ethanol and acetone are the dehydrants normally used. Unlike acetone, ethanol is not miscible with most resins and requires a linking agent (clearing agent) after dehydration. The linking agent normally used is propylene oxide (2-epoxypropane).

Impregnation
This is accomplished by progressively replacing the fixative or dehydrant with resin. With the more viscous resins impregnation is best performed under vacuum.

Processing schedules
Processing will vary depending upon the type and dimensions of the specimen. Tables 1 to 6 should provide a manual processing schedule suitable for most eventualities using epon-araldite resin whilst Table 7 gives a methacrylate protocol. Infiltration is quicker if vacuum is applied.

Resins
Epoxy resins (araldite,⁶ epon, Spurr⁷)
The term epoxy (or 'epoxide') refers to a chemical group in which an oxygen atom is bonded to two carbon atoms to form a three membered ring.⁸ An epoxy resin is a substance which contains the above groups and is also capable of polymerisation to form a rigid three dimensional structure with cross linking between molecular chains. Once formed the structure is not reversible. Epoxy resins have high mechanical strength, are easily polymerised and produce little shrinkage.⁸ These resins are relatively viscous which can lead to problems with infiltration and irregular polymerisation. Slow stirring can overcome these difficulties however vigorous agitation should be avoided as it generates air bubbles which interfere with the process and can inactivate the accelerator.⁵

Epoxy compounds are 'hard' resins from which 0.5 to 1 µm thick sections can be cut using either a glass or diamond knife on a motor driven microtome. Retraction of the block on the upstroke so that the cut face does not contact the back of the knife is a particular advantage. Hard resins cannot be cut on basic rotary microtomes using steel knives.⁹ Low viscosity epoxy resins such as vincyclohexane dioxide, diglycidyl ether and diepoxyoctane are capable of infiltrating large specimens and, when set, are more easily cut⁵ (Table 8).

Occupational exposure to epoxy resins is a common cause of allergic contact dermatitis.^10-11 These agents are also probable carcinogens, primary irritants, and systemically toxic.¹² They are fire hazards and environmental pollutants.

Resin mixtures
Epon araldite¹³
Araldite, Epon, hardener and accelerator are best obtained from the same source for compatibility. Table 8 lists various epoxy resins, hardeners and accelerators which are suitable.

Araldite embedding solution
REAGENT REQUIRED
1 Araldite stock solution
Araldite 10 g
Epon 10 g
Hardener 15 g
Mix thoroughly using a mechanical shaker. The stock is stable and will keep for up to 3 weeks if stored in a sealed, disposable polythene container at -20°C. Allow to reach room temperature before use.

2 Working solution
Add 2 g accelerator to every 100 ml stock solution and mix thoroughly. If the embedding solution is to be diluted with acetone or propylene oxide the dilution is made after the addition of accelerator.

ACRYLIC RESINS (METHACRYLATES)
Methacrylic acid [CH2=C.(CH3)COOH] and acrylic acid [CH2=CH.COOH] are the basic acrylic compounds commonly used in Histology. Methacrylates rapidly infiltrate fixed, dehydrated tissues at room temperature, however, damage to tissue is common during polymerisation because of marked, variable shrinkage of tissue components.⁸ These resins are more suited to embedment of hard materials such as undecalcified bone.

Methacrylates are 'soft' resins from which 0.5 to 2 µm thick sections can be readily obtained using tungsten carbide, glass or diamond knives. Sectioning is optimal if a motor driven microtome with retraction of the block on the upstroke is used. However, a non-motorised microtome can be effective provided that it is capable of retracting the block.

BUTYL/METHYL METHACRYLATE
Butyl and methyl methacrylate are similar esters which can spontaneously change from the monomer (liquid state) to polymer (solid state); the process is hastened if an accelerator (benzoyl peroxide) is added.⁸ An inhibitor is normally incorporated into the commercial product to prevent spontaneous polymerisation during storage. This agent can be removed by washing the monomer in a separating funnel with 5% aqueous sodium hydroxide.^3,14

Partial polymerisation before infiltration partly overcomes the problem of variable shrinkage of tissue elements (see earlier). Partially polymerised methacrylate remains fluid enough to infiltrate tissue whilst there is far less shrinkage after full polymerisation.⁸

Partial Polymerisation of Methyl Methacrylate
REAGENT REQUIRED
1 Methyl methacrylate polymer
Benzoyl peroxide (dry) 1 g
Methyl methacrylate (washed) 100 ml
The resin is washed in 5% aqueous sodium hydroxide before use.

2 5% aqueous sodium hydroxide
Sodium hydroxide pellets 5 g
Distilled water 100 ml

METHOD
1 Add benzoyl peroxide to the washed methyl methacrylate in a pyrex beaker.
2 Place in a water bath and raise the temperature to betwen 80 and 82°C (measure from a thermometer in the methacrylate). The temperature should not be allowed to exceed 82°C (violent frothing followed by full polymerisation will result).
3 The solution will begin to thicken. When it is a consistency similar to glycerol cool rapidly in cold water to room temperature.
4 The solution is stable and can be stored in a tightly stoppered container at 4°C for up to 12 months.

GLYCOL METHACRYLATE
This is the ethylene glycol monoester of methacrylic acid.⁸ It forms addition polymers, at low temperatures, after initiation of polymerisation with benzoyl peroxide and is the water miscible form of butyl methacrylate. These two properties make it ideal for enzyme studies. Tissue preservation tends to be better than that obtained with methyl methacrylate processing.¹⁵

REAGENTS REQUIRED
Monomer (Infiltration mixture)
Glycol methacrylate 80 ml
2-butoxyethanol (plasticiser) 16 ml
Benzoyl peroxide (initiator) 1.2 g
Stir until the initiator has dissolved. This solution can be stored for up to 6 months and may be filtered and reused.

Polymer (Embedding mixture)
To 40 parts of monomer add 1 part of following solution:
Polyethylene glycol 400 (plasticiser) 15 ml
N:N dimethyl aniline (accelerator) 1 ml
Mix thoroughly and use immediately.

Glycol methacrylate will polymerise more rapidly than methyl methacrylate (1 to 2 hours at room temperature or in 4 to 6 hours at 4°C ).

Special procedures
Enzyme demonstration
METHOD
1 Glycol methacrylate polymer is mixed with distilled water (Table 9) and used as the infiltrating medium at 4°C.
2 The solution should be changed at 6 hourly intervals.

NOTE the tissue is not dehydrated.¹⁶

Lipids
Tissue treated with osmium tetroxide and embedded in resin is suitable for accurate quantitation of lipid volume.^17-18

METHOD
1 Fix cores of tissue (commonly liver) in 10% neutral buffered formalin for 24 hours.
2 Wash the samples briefly in distilled water.
3 Place into a mixture of equal parts 5% potassium dichromate and 1% osmium tetroxide (this is a very toxic solution and great care is required in its preparation) for 6 hours at room temperature.
4 Wash in running water for 2 hours.
5 Process as indicated in Table 2.

RESULT
Fat droplets dense black
Background relatively clear
Other stains can be performed on sections from the treated biopsy.

AUTORADIOGRAPHY
High resolution autoradiographs have been obtained from tissue sections cut at 2 µm after embedding in glycol methacrylate. A range of tissues were examined including spleen, testis, small intestine and colon. Plastic sections were stained with the Feulgen reaction then processed for autoradiography.¹⁹

LOW TEMPERATURE EMBEDDING MEDIA
To overcome problems associated with high temperature denaturation of proteins and loss of enzymes, special low temperature acrylic resins, based on acrylate and methacrylate, have been formulated. They are used at temperatures ranging between -35°C and -70°C and are polymerised by UV irradiation.^20-21

HYBRIDISATION HISTOCHEMISTRY
Hybridisation histochemistry has been successfully performed on sections from epoxy resin and lowicryl embedded material.^22-23

Microtomy
Glass knives are better suited to sectioning resin embedded material than steel knives as the molecular glass edge is much sharper. Long edged glass knives (Ralph knives) are the most effective,²⁴ however, suitable semi-thin sections can be obtained using steel knives. If glass knives are used the block size is limited by the width of the knife (usually 6.4 mm or 9 mm) and the length of the microtome cutting stroke. Water troughs can be attached directly to glass knives.

Impregnated tissue tends to expand more than clear resin therefore, to avoid folds in cut sections, resin should be trimmed as close as possible to the embedded tissue before sectioning (approximately 1 mm).²⁵ Face trimming the block should be carried out in steps no greater than 5 µm to avoid damage to the face and is best accomplished immediately after polymerisation, while the block is still warm. Sections are generally cut longitudinally to minimise the mechanical stress placed upon the block and knife.

Dry sectioning is preferred for epoxy resin whilst wet is better for methacrylate. Sections, cut using a glass knife fitted with a trough, will float directly on to the water from which they can be transferred to a pool of water upon a clean, glass slide. This in turn is placed on a hotplate set at 60°C to expand and dry the section. A moist cotton bud can be used to collect resin sections which are cut dry.²⁶ The cotton bud is rolled with the section as it travels down the knife. Adhesives are generally not required but are recommended for decalcified or undecalcified bone. A guide to section thickness for a range of tissues and staining procedures can be found in Table 10.

A method has been described for preparing thin resin sections without using a microtome.²⁷ Thick sections (0.5 mm) are cut with a diamond saw after embedding in Spurr resin. The section obtained is glued to a glass slide, ground to approximately 75 µm then polished. The polished sections can be stained, after removal of the resin, by H&E.

Resin removal
Hard resins provide a barrier to dyes and markedly reduced staining clarity. Removal of this type of resin is therefore recommended before staining (resin cannot be removed from ‘soft’ methacrylate sections). Epoxy resins are easily degraded by alcoholic sodium hydroxide,²⁸ potassium hydroxide,²⁹ sodium methoxide³⁰ or bromine vapour.³¹

Saturated Alcoholic Sodium Hydroxide or Potassium Hydroxide
REAGENT PREPARATION
1 Sodium hydroxide saturated in absolute ethanol
Place sodium hydroxide pellets in a reagent bottle, fill with absolute ethanol and shake well. The supernatant can be used after 24 hours.
2 Potassium hydroxide saturated in absolute ethanol
Place potassium hydroxide pellets in a reagent bottle, fill with absolute ethanol and shake well. Seal the reagent bottle and store until the supernatant turns brown (2-3 days). Filter before use.

METHOD
1 Transfer resin sections directly from a hot plate into a coplin jar containing alcoholic sodium or potassium hydroxide and leave for 2 to 5 minutes.
2 Wash in three changes of absolute ethanol.
3 Rinse in 70% ethanol.
4 Wash in running tap water for 5 minutes.

Staining methods
Staining techniques for paraffin sections are generally applicable to resin sections with minimal or no modification once the resin has been removed. These include periodic acid Schiff (PAS), periodic acid Schiff/diastase (PAS/D), Perls’ prussian blue, toluidine blue/nile blue sulphate for mast cells, Von Kossa and reactions (Table 10). However, resin cannot be removed from methacrylate embedded tissue and some aqueous dyes, particularly trichrome stains, penetrate the resin poorly. In these instances staining times may need to be increased to produce satisfactory results.

Enzyme histochemistry and immunohistochemistry can also be performed on deplasticised resin sections using the same techniques as those used for paraffin sections³² and with a range of polyclonal antisera.³³

GENERAL STAIN
Toluidine blue is commonly used as a general stain for epoxy resin sections.³⁴ Removal of resin is not required and the technique is useful for determining the general morphology of a specimen before sectioning further.

Toluidine Blue Rapid Stain
REAGENT REQUIRED
Toluidine blue (CI 52040) 1 g
Borax 5 g
Distilled water 100 ml
Mix and stand overnight then filter before use.

METHOD
1 Flood a section, on a hot plate, with toluidine blue solution for up to 30 seconds (do not allow the section to dry).
2 Wash in running tap water for a few seconds.
3 Remove excess water from around the section then dry on a hot plate.
4 The section can be examined uncovered or mounted if a permanent preparation is required.

RESULTS
Tissue components - blue

OTHER METHODS
The following modified methods have proven successful when applied to sections after removal of resin.

Haematoxylin and Eosin
Routine H&E procedures will generally give satisfactory results although staining times may need to be increased for optimum results. Buffered alcoholic eosin or picro-eosin are a more suitable counterstains than aqueous or alcoholic eosin.

REAGENTS REQUIRED
Buffered alcoholic eosin
1% eosin Y (CI 45380) in 95% ethanol 250 ml
0.2 mmol/l sodium acetate in 95% ethanol 80 ml
1 mmol/l acetic acid in 95% ethanol 170 ml
95% ethanol to 2 l
Counterstain for 5 minutes followed by dehydration in absolute ethanol.

Picro-eosin
Eosin Y (CI 45380) 10 g
Potassium dichromate 5 g
Picric acid (saturated aqueous) 100 ml
Absolute ethanol 100 ml
Distilled water 800 ml
Mix in the proportions given then dilute 50:50 with distilled water for use. Stain for 4 to 6 minutes followed by a brief wash in water before dehydrating and mounting.

Jenner-Giemsa Stain (for bone marrow trephines)³³
REAGENTS REQUIRED
1 Jenner solution
Jenner dye 10 g
Methanol 2.5 l
2 Giemsa stock solution
Giemsa powder 38 g
Methanol 250 ml
Glycerin 250 ml
Place in a flask and plug the neck loosely with cotton wool. Warm in a water bath at 56°C for 1 hour stirring frequently. Cool and filter before use.
3 Giemsa working solution
Giemsa stock 1 ml
0.2 mmol/l Walpole's acetate buffer pH 4.7 2 ml
Distilled water 37 ml

METHOD
1 Immerse section in Jenner solution for 15 minutes.
2 Transfer directly to Giemsa working solution for 20 minutes.
3 Differentiate in acetone (4 to 5 quick dips), repeat with fresh acetone.
4 Treat with 1:1 acetone:xylene (6 quick dips).
5 Treat with xylene then mount in neutral mounting medium.

Reticulin (Naoumenko)³⁵
SECTION PREPARATION
Resin sections for reticulin staining should be cut at 4 µm.

REAGENTS REQUIRED
1 Acidified potassium permanganate
0.1% aqueous potassium permanganate 95 ml
3% aqueous sulphuric acid 5 ml
2 Silver solution
8% aqueous ammonium nitrate 7 ml
Distilled water 35 ml
4% aqueous sodium hydroxide 8 ml
10% aqueous silver nitrate 3.8 ml
Make up the solution immediately before use. Add the ingredients in the order give with gentle shaking.
3 1% aqueous oxalic acid
4 2% aqueous iron alum
5 0.1% aqueous gold chloride
6 5% aqueous sodium thiosulphate
7 0.2% formalin

METHOD
1 Deplasticise section and rehydrate.
2 Oxidise in acidified potassium permanganate for 1 minute.
3 Wash in distilled water.
4 Bleach in 1% oxalic acid for 1 minute.
5 Wash in distilled water.
6 Treat with 2% iron alum for 1 to 2 minutes.
7 Wash in distilled water.
8 Immerse in ammoniacal silver solution for 1 to 2 minutes.
9 Rinse briefly in 70% ethanol.
10 Immerse in 0.2% formalin in a coplin jar with agitation for 30 seconds.
11 Immerse in fresh 0.2% formalin in a coplin jar for 1½ to 2 minutes.
12 Rinse in distilled water.
13 Tone in 0.1% gold chloride (optional) for 1 minute.
14 Rinse in distilled water.
15 Fix in 5% sodium thiosulphate for 1 minute.
16 Rinse in distilled water.
17 Dehydrate, clear and mount in neutral mounting medium.

RESULTS
Reticulin - Black

Masson trichrome
REAGENTS REQUIRED
1 Slidders iron haematoxylin³⁶
Haematoxylin (CI 75290) 0.5 g
Aluminium chloride 5 g
Ferrous sulphate 5 g
95% ethanol 50 ml
Distilled water 50 ml
Hydrochloric acid (conc) 1 ml
9% aqueous sodium iodate 1 ml
Dissolve the haematoxylin in 95% ethanol and aluminium chloride and ferrous sulphate in distilled water. Combine the two solutions then add concentrated hydrochloric acid and sodium iodate. Allow the solution to ripen for 48 hours before use. The solution remains stable for 4 to 6 weeks.
2 Biebrich scarlet/acid fuchsin
1% aqueous biebrich scarlet (CI 26905) 90 ml
1% aqueous acid fuchsin 10 ml
Glacial acetic acid 1 ml
3 Aniline blue staining solution
Aniline blue (CI 42755) 1 g
Distilled water 99 ml
Glacial acetic acid 1 ml
4 Differentiating solution
Phosphomolybdic acid 5 g
Phosphotungstic acid 5 g
Distilled water 200 ml
5 Acid alcohol solution
Absolute ethanol 700 ml
Distilled water 295 ml
Hydrochloric acid (conc) 5 ml

METHOD
1 Deplasticise and rehydrate sections.
2 Mordant in Bouin's fluid at 56°C for 1 hour.
3 Wash in water.
4 Stain with Slidder's haematoxylin for 5 minutes.
5 Wash in water.
6 Differentiate briefly in 0.5% hydrochloric acid in 70% ethanol.
7 Wash in water.
8 Stain with biebrich scarlet solution.
9 Rinse briefly in water.
10 Differentiate in differentiating solution for 10 minutes.
11 Wash in water.
12 Stain in aniline blue solution for 4 minutes. 13 Wash in water, dehydrate, clear and mount in a neutral mounting medium.

RESULTS
Nuclei - Blue/black
Cytoplasm - Red
Collagen - Blue

References
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