Cryotechniques for light microscopy
Mark Donovan and Henry Preston

Introduction
Freezing of unfixed or fixed tissue may be required for a number of reasons:

The major application of cryotechnology is in rapid diagnostic processing as part of an intraoperative consultation. The production of sections for lipid demonstration, enzyme histochemistry and immunocytochemistry differ little from the methods employed in rapid diagnostic applications.

Theory of freezing
The reasons for freezing a tissue sample are to provide a hardened matrix for sectioning or to preserve the morphological, biochemical or immunological properties of a cell or tissue. The use of a low temperature can eliminate many of the problems associated with standard practices of chemical fixation and paraffin or resin embedding. In practice however, the process of transforming water into ice can dramatically alter the physical and chemical structure of cells and tissue.

When pure water is cooled, large hexagonal ice crystals form as a result of homogeneous and heterogeneous nucleation and subsequent growth of ice crystal nuclei. The transformation of water into ice crystals occurs at a temperature of OC and a pressure of 1 atmosphere. Where a very high rate of cooling exists, cubic rather than hexagonal ice crystals will form. These are far smaller and produce less distortion on formation. If the rate of cooling is increased further to within the order of 104K/sec small volumes of water can be solidified without the formation of ice crystals at all.1 This solid form of ice, known as vitreous ice, exists in a temperature dependent, irreversible phase transition with cubic ice, hexagonal ice and water.2 The critical temperatures at which these phase transitions occur have only been accurately determined for pure water3 but the values for intracellular water are considered to be significantly higher (fig 1).4-5

The formation of large, hexagonal ice crystals as the result of slow freezing occurs firstly in the less concentrated extracellular fluid. This produces an osmotic difference between the extracellular and intracellular fluid which results in a loss of intracellular water and the subsequent shrinkage of the cell as the osmotic balance returns. The increased ionic concentration within the cell ruptures membranes and denatures the protoplasm. The eventual formation of ice crystals within the cell, when residual intracellular fluid freezes, may then mechanically fracture the cell. These effects are collectively known as ice crystal artefacts.

The damage produced by ice crystals depends upon the size and type of crystal formed during the freezing process. Large hexagonal ice crystals will produce major structural damage to cells and tissue whilst smaller cubic ice crystals cause less cellular damage. Ideally an extremely rapid cooling rate should be used as this will produce vitreous ice without crystal damage. The aim when freezing the sample, therefore, is to limit ice crystal formation as much as possible through control of the cooling rate.

Factors which effect the cooling rate of the sample include:

Freezing of the specimen will require selection of an appropriate cryogen and procedures which maximise the heat exchange between cryogen and specimen. The cooling rate attainable is particularly influenced by specimen size. Above a critical specimen size optimal freezing will only occur to a certain depth and the cooling rate in the deeper parts of the sample will be slow enough to allow the formation of hexagonal ice crystals with subsequent tissue damage. It is possible to reduce the level of ice crystal formation through the use of cryoprotectants which reduce the rate of ice crystal nucleation through freezing point depression.

Cryoprotectants
Cryoprotectants are thought to increase viscosity at sub zero temperatures thereby decreasing the mobility of water molecules. The constrained water molecules are prevented from forming ice crystal nuclei and ice crystal formation is inhibited.

Cryoprotectants in common use include non-penetrating agents such as polyvinyl pyrrolidone (PVP), dextran, hydroxyethyl starch and sucrose or penetrating agents such as dimethyl sulphoxide (DMSO), glycerol, ethylene glycol and dimethyl formamide (DMF). Of these a solution of 0.5 mol/l sucrose in 3.5 mol/l DMSO allowed to infiltrate the sample at room temperature prior to freezing is highly recommended.6

Cryogen selection
A number of cryogens with differing absolute temperatures and levels of heat transfer efficiencies at the sample-cryogen interface are available. Selection requires consideration of the degree of acceptable ice crystal damage in addition to convenience, availability and cost.

Liquid nitrogen (-196C)
Produces rapid freezing although overcooling can cause cracking and brittleness. A layer of gaseous nitrogen can form at the sample surface reducing the heat transfer considerably. The use of talc to coat the sample may overcome this effect.

Liquid nitrogen - isopentane (-150C)
The use of a second liquid such as isopentane or hexane which is supercooled by immersion in liquid nitrogen is very effective in reducing freezing artifact. This combination has been recommended for histochemistry although the preparation is somewhat lengthy for rapid diagnostic procedures.

Electric cryobath - isopentane (-60C)
Commercial thermoelectric cryobaths which cool isopentane or hexane have the advantage of easier preparation and although the cooling rate is not as rapid, freezing artifacts are minimal.

Carbon dioxide gas (-70C)
Freezing attachments which connect to a cylinder of carbon dioxide rapidly freeze a sample by heat exchange from the expanding and vaporizing gas. These attachments are simple and efficient for routine work but freezing artifacts are produced.

Solid carbon dioxide (-70oC)
The use of solid chunks of carbon dioxide (dry ice) held against the specimen can be effective and is acceptable for routine freezing.

Solid carbon dioxide - hydrocarbon slurry (-79C)
Pellets of solid carbon dioxide are mixed with isopentane, hexane, acetone or ethanol to form a slurry. The specimen may be either plunged directly into this mixture or into a container of pure hydrocarbon cooled in the hydrocarbon-carbon dioxide slurry.

Aerosol sprays (-50C)
Commercial aerosol freezing sprays containing gases liquefied under pressure are a simple and convenient method for freezing specimens for general routine work. However the heat transfer is slow and uneven making it prone to freezing artifact.

Thermoelectric cooling (-40C)
This type of cooling employs the Peltier effect which relies upon the use of two metal conductors. When a current is passed between the two conductors, heat is absorbed at the junction so a sample in contact with this surface will undergo heat transfer and freeze. This type of cooling device is employed as part of the stage of some freezing microtomes.

Refrigerated contact (-30C)
The use of compressor driven refrigeration coils to cool a solid metal block of aluminium or insulated cabinet forms the basis of the freezing techniques employed in most cryostats. The cooling rate is slow but the method is simple.

Frozen sections
The production of a frozen section would appear to be straightforward. Intra- and inter-cellular water is frozen to produce a hard matrix to enable thin slicing of the tissue. The reality is more complex with an interplay of specimen type, microtome knife, microtome and environmental temperatures. As with any microtomy, the condition and set-up of the microtome will greatly influence the outcome.

When sectioning frozen tissue the sample must be firmly attached to the microtome stage or chuck. Saline or water will hold the sample when frozen however, embedding media are preferred as these also provide a supportive and protective aid to sectioning. The most widely used embedding medium is OCT, an aqueous solution of glycols and resins which provides an inert matrix for sectioning. Alternatives to OCT compound include 30% bovine albumin and von Apathy's gum syrup.

Microtomes in frozen sectioning
Any of the various forms of microtome (rotary, rocking, moving knife, sledge or ultra microtome) can be used to produce sections of frozen tissue. The only modification necessary is a means of maintaining the tissue in a frozen state while sectioning. Piped carbon dioxide, refrigerated coils, and thermoelectric devices have all been applied to microtome design to satisfy this requirement.

The term 'cold knife' is sometimes used to describe techniques or devices in which only the specimen and microtome knife are cooled but sectioning is performed at room temperature. When the entire microtome and specimen are completely enclosed in a cooled chamber the term 'cold cabinet' or 'cryostat' is applied. Of the cryosectioning equipment available7 the freezing microtome and cryostat are the instruments most frequently encountered.

Freezing microtome
The standard freezing microtome consists of a radial arm attached to a central pivot. On this arm two clamps hold a wedge profile microtome knife with its cutting edge inclined to the horizontal plane. The knife is moved in a horizontal arc over a specimen chuck mounted on a central threaded spindle. When the knife is moved back and forward, the chuck is advanced upward by a micrometer mechanism in proportion to the angle of the arc through which the knife is moved. The tissue is mounted onto the specimen chuck or freezing stage which is traditionally made of hollow metal with holes in its sides. When compressed carbon dioxide is piped into the chuck expanding and vaporizing gas absorbs heat and the tissue on the stage is rapidly frozen. A separate nozzle may also direct carbon dioxide onto the knife in order to lower its temperature.

Alternatively, freezing stages based on a refrigerated coil or thermoelectric module are available. A particular advantage of thermoelectric cooling is the capability to adjust accurately and maintain the temperature of the tissue and knife.

The freezing microtome is used mainly in the sectioning of hard or dense tissues such as cartilage and bone, or for cutting large sections of neurological tissue. Sections less than 5 m in thickness can be difficult to obtain.

FIXATION
Tissue to be sectioned using the freezing microtome may be either fresh or fixed. The production of good quality sections from unfixed tissue however is extremely difficult due to the problem of maintaining a low optimum cutting temperature of the frozen tissue in a room temperature environment. Cut sections tend to thaw quickly and are sticky to handle because of the unfixed proteins present. It is preferable therefore to fix the tissue before sectioning unless this is precluded by the proposed staining procedure. The traditional approach has been to fix tissue for 1 to 2 minutes in 10% formalin or normal saline heated to boiling. Formol calcium is also suitable but alcoholic fixatives must be avoided as alcohol inhibits the freezing point of the tissue.

INFILTRATING MEDIA
Sections from certain fixed tissues such as liver, spleen and brain have a tendency to shatter on cutting. This is particularly so for the 20-40 m sections often required for neuropathology. In order to reduce the brittleness of these tissues a specimen can be infiltrated with a protective medium. Various media have been utilised including 30% sucrose in 10% formalin,8 1% gum acacia in 30% sucrose,9 10% glycerol in 4% formaldehyde with 2% dimethyl sulphoxide10 and an aqueous glycerine-gelatine mixture.11 The use of OCT compound has also been proposed12 with the method of Ishii et al13 being particularly simple and effective. All of the infiltrative methods however, are time consuming taking from several hours to several weeks.

Gelatin Infiltration Method11
REAGENTS REQUIRED
1 Gelatin 16 g
2 Glycerine 15 ml
3 Distilled water 70 ml
4 Thymol one crystal
Dissolve the gelatin in distilled water and add glycerin. Store at 4C (thymol acts as a preservative) and melt in a 37C water bath for use.

METHOD
1 Fix tissue in 10% formalin overnight.
2 Wash thoroughly in running water for 6 - 8 hours.
3 Impregnate in gelatin-glycerine mixture at 37C for 6 hours.
4 Transfer tissue to fresh gelatin-glycerine mixture, embed in a suitable mould and allow to set at 4C.
5 Trim block and harden the gelatin-glycerine by fixing in 10% formalin overnight or until ready for sectioning.

OCT Infiltration Method13
METHOD
1 Wash fixed tissue in running water.
2 Infiltrate in neat OCT for 5 hours at room temperature.
3 Remove from OCT and blot off excess.
4 Infiltrate in fresh OCT for 4 hours at room temperature.
5 Remove from OCT and blot off excess.
6 Infiltrate in fresh OCT for 15 hours at room temperature.
7 Embed in fresh OCT and freeze for sectioning.

TECHNICAL NOTES
1 The volume of OCT during infiltration should be in excess of 30 times the volume of tissue.
2 Times given are for a specimen of 5 mm thickness and should be reduced or increased for smaller or larger specimens.
3 Sections are best cut on a cryostat with a chamber temperature of -5C.

SETTING UP THE FREEZING MICROTOME (TRADITIONAL CO2 TYPE)
The microtome should be firmly clamped to the edge of a bench or table. The accompanying carbon dioxide cylinder, unless a rising-pipe type, must be inverted or have its valve lower than the base by 20 cm. Connection to the microtome is by means of a flexible, metal-reinforced hose which must not be twisted or allowed to kink.

Tissue Freezing And Sectioning14-15
METHOD
1 Open the valve on the carbon dioxide cylinder slightly (one half to one turn).
2 Place a small amount of saline, water or OCT on the freezing stage and on this place the tissue.
3 Apply gentle downward pressure on the tissue to hold it in place and freeze using short bursts of carbon dioxide. This produces quick freezing while conserving carbon dioxide and preventing blockage of the freezing stage outlets by carbon dioxide snow. The tissue is adequately frozen when the whole piece becomes white and firm.
4. Raise or lower the centre spindle until the surface of the tissue is just in contact with the knife. Set the section thickness control to the required setting. Cutting is best done using a firm, steady motion and gentle pressure. Cool the knife with short bursts of carbon dioxide.
5 Allow the most superficial portion of the tissue to thaw. Cut a number of sections until the required consistency and thickness are obtained.
6 Remove the debris from the knife and place a drop of water on the upper surface of the knife where the sections will accumulate. Cut the desired number of sections.
7. Remove sections from the knife with a saline moistened brush. Alternatively, with unfixed tissue and using a cooled dry knife, the section may be picked up directly onto a warmed slide. The section will thaw on contact and adhere to the slide surface.
8. Transfer the free sections to a dish of normal saline using light movement to separate each.
9. When cutting is complete, turn off the carbon dioxide cylinder valve and release the pressure in the connecting hose. The microtome and knife should be well cleaned, dried and suitably oiled after use to prevent rusting.

MANIPULATION OF FREE FLOATING SECTIONS
The frozen sections produced can be handled in several ways.

Haematoxylin And Eosin Staining Of Free Floating Sections
METHOD
1 Using a bent glass rod place sections in a small dish of distilled water.
2 Transfer to alum haematoxylin (Harris', Gill's, Mayer's or Carazzi's) for 1-5 minutes. The actual time required will vary according to section thickness and haematoxylin used. The section will need to be retained on the glass rod during the staining steps otherwise it will be lost in the staining solution.
3 Transfer through two changes of distilled water.
4 Transfer to alkaline water for 1 minute (to blue the nuclei).
5 Wash in three changes of distilled water.
6 Transfer to 1% aqueous eosin for 30 seconds to 1 minute.
7 Transfer to 95% ethanol for 1 minute.
8 Transfer through three changes of absolute alcohol.
9 Transfer through two changes of xylene or xylene substitute.
10 Transfer to a final change of xylene or xylene substitute and using a glass rod, mount the sections on a slide and allow to drain.
11 Mount in synthetic mounting medium.

RESULTS
Nuclei - blue
Cytoplasm, collagen - shades of pink

The cryostat
To obtain frozen sections consistenly below 5 m in thickness it is necessary to use a cryostat. The cryostat overcomes the problem of temperature maintenance inherent in the freezing microtome by housing the entire sectioning apparatus (a rotary, rocking or sledge microtome) in an insulated, thermostatically controlled refrigerated cabinet with some if not all the controls being operated on the outside. Temperature control of the cabinet is generally within the range of ambient to -40C. Most currently available cryostats have an area within the cabinet for rapidly freezing a specimen onto a chuck whilst some also have the facility to maintain specimen temperature independently of the cabinet temperature during sectioning. This is of particular advantage for rapid diagnostic work where the optimum cutting temperature for different tissue types can be adjusted quickly without altering the cabinet temperature.

The cryostat is generally left operating at all times. It is particularly important to maintain the ventilation ducts of the refrigeration unit free of obstruction as poor air flow will result in reduced compressor life and efficiency. The cryostat microtome should be kept clean of tissue debris and lubricated regularly with a low temperature oil.

THE MICROTOME KNIFE
A wedge shaped (C profile) knife that is faultlessly sharp and of high quality stainless steel to minimize rusting is required for optimal sectioning. The knife should be cleaned with xylene followed by alcohol after each use. Applying a coating solution, teflon spray or rubbing the knife with a metal polish is recommended to reduce friction during sectioning. A spare microtome knife should always be stored in the cryostat so that it is maintained at cabinet temperature.

Setting the correct knife angle,ideally between 30 to 50, will prevent damage to the block face and reduce buckling of the section. An alternative to the traditional wedge knife is to use a disposable blade. These offer a consistently sharp edge, easy changing and no corrosion problems.

Knife Coating Solution
REAGENTS REQUIRED
1 Solution A
Copper Sulphate 10 mg
Stannous chloride 15 mg
Distilled water 1 l
2 Solution B
Sodium thiosulphate 1 kg
Sodium chromate 0.1 gm
Distilled water 1 l

METHOD
1 Rinse the clean, sharpened microtome knife in distilled water.
2 Place in solution A for 10 seconds.
3 Rinse in distilled water.
4 Place in Solution B for 20 seconds.
5 Rinse in distilled water.
6 Allow to air dry.

THE ANTI-ROLL PLATE
To prevent sections from curling as they are cut most cryostat microtomes are fitted with an anti-roll plate. The plate is usually manufactured from perspex and is intended to sit about 50 m above the knife surface so a cut section passes between it and the knife. Small ridges on the plate or cellulose tape applied to each corner provide the elevation. The plate must be positioned parallel with the cutting bevel of the knife and protrude slightly beyond it and also be kept free of tissue debris during sectioning. The application of teflon spray will reduce friction and section adhesion to the anti-roll plate.

OPTIMAL CUTTING TEMPERATURE
In most cases the freezing of tissue will occur at a temperature below that at which the cryostat will produce the best sections and a short period of time will be required before the tissue warms to operating temperature. Sectioning hard, overcooled tissue should be avoided as it dulls the knife edge and can damage the tissue surface. As the optimum cutting temperature is measured at the knife edge, the capability of some cryostats to independently control the temperature of the cryostat cabinet, microtome knife and specimen chuck is particularly advantageous. Maintaining the microtome knife at a temperature a few degrees lower than the tissue greatly assists sectioning. In other cryostats the same effect can be obtained by applying an aerosol freezing spray or fragments of dry ice to the microtome knife.

The optimum cutting temperature for a range of tissue types is given in Table 1. Due to differences in composition, cellularity, connective tissue and fat content the optimum temperature for sectioning different tissues varies. Since a cryostat will require time to reach the optimum temperature for any given tissue it is usual practice to maintain a setting of -20C as a compromise. Most tissue will cut effectively within ± 5C of this temperature. The exceptions are liver, brain, spleen and uterine curettings which shatter in this temperature range and are sectioned -10C.

ORIENTATION OF THE SPECIMEN
The production of a frozen section is greatly assisted by having a well-orientated specimen with a flat cutting surface within a small rim of embedding medium. The use of plastic cryomoulds or circling the rim of a round stub style of chuck with tape to produce a well are of great value in orientation of the specimen within the embedding medium.16-17 The use of a mounted, cooled chuck for freezing and creating a flat cutting surface has also been suggested18. Very small specimens are best supported on a mound of embedding agent pre-frozen on a chuck. This provides extra elevation above the surface of the chuck to enable safe sectioning.

FREEZING METHODS FOR CRYOSTAT SECTIONING
The selection of a particular freezing procedure will be determined largely by individual requirements. The following methods are suitable in most situations but are listed in decreasing order of preference.

Liquid Nitrogen - Isopentane Method (-150C)
REAGENTS REQUIRED
1 Isopentane
2 Liquid nitrogen
3 Suitable embedding compound (such as OCT Compound)

METHOD
1 Place 50 ml of isopentane in a pyrex or polypropylene beaker.
2 Immerse the beaker in a dewar or styrofoam container of liquid nitrogen.
3 Stir the isopentane until opalescent (about 2-3 minutes).
4 Remove the isopentane beaker from the liquid nitrogen.
5 Place OCT compound (or similar) on a specimen stub or in a cryomould and orientate the specimen within it.
6 Immerse the specimen into the cooled isopentane until frozen.
7 Place the frozen block into the cryostat for sectioning or store at -70C until required. The specimen will require warming to its optimal cutting temperature before sectioning.

Dry Ice-Acetone-Isopentane Method (-79C)
REAGENTS REQUIRED
1 Acetone
2 Isopentane
3 Dry ice (solidified carbon dioxide)
4 Suitable embedding compound (such as OCT Compound)

METHOD
1 Place 30 ml of isopentane in a pyrex or polypropylene beaker.
2 Prepare a carbon dioxide-acetone slurry by adding solid carbon dioxide pieces to a dewar of acetone.
3 Immerse the beaker in the dewar of carbon dioxide-acetone slurry.
4 Allow the isopentane to cool.
5 Place OCT compound (or similar) on a specimen stub or in a cryomould and orientate the specimen within it.
6 Immerse the specimen into the cooled isopentane until frozen.
7 Place the frozen block into the cryostat for sectioning or store at -70C until required. The specimen will require warming to its optimal cutting temperature prior to sectioning.

Cryostat Quick Freeze Attachments (-40C to -30C)
REAGENTS REQUIRED
Suitable embedding compound (such as OCT Compound)

METHOD
1 Place OCT compound on a microtome specimen stub or cryomould and orientate the specimen within it.
2 Quickly transfer the stub to the freezing bar or spot in the cryostat chamber.
3 Apply the heat exchanger to the tissue. This will accelerate cooling and create a flat, even surface on the specimen.
4 When the OCT and tissue are frozen remove from the freezing area and place in microtome for sectioning.
5 Depending on the tissue type the specimen may require slight warming to its optimal cutting temperature.

Cryostat Sectioning Procedure14,19
The various difficulties which might arise during section preparation and their possible causes are listed in Table 2.

METHOD
1 Select a representative piece of tissue trimmed to no larger than 2 cm x 2 cm x 4 mm. The 4 mm thickness is particularly important to minimize freezing artifact.
2 Place the tissue on a chuck with a base of OCT compound (or similar).
3 Quickly freeze the tissue with either
(a) Liquid nitrogen with or without isopentane bath,
(b) Solid carbon dioxide - isopentane slurry or
(c) Cryostat quick - freeze facility.
If these methods are not available the following will suffice:
(d) Solid carbon dioxide contact or
(e) Aerosol cryospray.
4 Secure the specimen chuck in the microtome (microtome should be in a locked position) and ensure the microtome advance mechanism is in the start position.
5 Release the microtome lock and advance or retract the chuck position until the knife edge just touches the block. Rough trim the superficial surface of the block in small steps (15-25 m) until an even, full face is achieved. Remove tissue debris from the knife with a soft brush or tissue.
6 Position the anti-roll plate, check the thickness setting and automatic advance selector. Allow the cryostat and/or specimen to reach the optimum cutting temperature.
7 Cut sections using a slow even motion, except for hard tissue which requires a firmer stroke. The section should glide smoothly under the anti-roll plate. Alternatively, a soft brush can be used to keep the section flat as it glides out on the knife surface.
8 With the section sitting flat on the knife surface, lower a clean labelled slide onto the section. It is best to rest one edge of the slide on the knife surface about 2 cm beyond the section and gently lower the other end towards the section. When the slide is about 1 mm from the knife the section will lift onto the slide from the knife surface.
9 Fix the slide rapidly in a fixative of choice, unless the staining procedure precludes fixation.
10 Wash the slide briefly in running water and proceed with the staining procedure desired.

TECHNICAL NOTE
Cryostat sections of fresh tissue adhere well to clean glass slides due to the naturally sticky nature of unfixed proteins. With fixed tissue or long staining protocols sections may become detached unless the slides are coated with an adhesive. Adhesives recommended include glycerin-albumin,15 albumin,20 chrome-glycerine jelly,14 chrome-gelatin11 and chrome-gelatin-formaldehyde.2 1 Poly-l-lysine and 5% polyvinyl alcohol can also be used.

FIXATION OF SECTIONS
With the frozen section firmly attached to the glass slide the decision to fix sections of fresh tissue will depend upon the diagnostic urgency, potential infectiveness and the staining procedure to follow. A number of fixatives are recommended for routine use with frozen sections. These are summarised below (except for Zamboni's Fixative, details of formulae can be found in other chapters).

Formol Acetic Alcohol (FAA) is a good fixative for rapid diagnostic applications. Excellent nuclear preservation and enhanced section adhesion.

Formol Alcohol is similar to FAA, a good fixative for rapid diagnostic work.

Carnoy's Fluid provides excellent nuclear preservation and is suitable for rapid diagnostic work. The defatting action of the chloroform makes it useful for breast sections where the fat can cause section detachment during staining.

Neutral Buffered Formalin (4%) is Suitable for routine use.

Acetone (100%) gives excellent antigen and histochemical preservation at 4C. It is used for immunocytochemical procedures where it may be coupled with periodate-lysine-paraformaldehyde post fixation for improved morphology22.

Periodate-Lysine-Paraformaldehyde (PLP) must be prepared fresh each day. Used at 4C for 8 minutes it is highly recommended for immunocytochemistry as it provides good morphological and antigen preservation.

Zamboni's fixative is used at 4C for 30 minutes and is recommended for immunocytochemistry.

Zamboni's Fixative
REAGENTS REQUIRED
1 Solution A
Saturated picric acid (store at 4C)
2 Solution B
Paraformaldehyde 100 g
Distilled water 400 ml
Heat to 60oC. Slowly add 1-3 drops of 1 mol/l sodium hydroxide, stirring until solution is clear.
3 Solution C
Sodium dihydrogen orthophosphate 3.31 g
Di-sodium hydrogen orthophosphate 33.7 g
Distilled water 1 l

METHOD
The working solution is prepared by mixing the following:
Solution A 150 ml
Solution B 100 ml
Solution C 750 ml

Staining of frozen sections
The procedures outlined are designed for rapid staining of frozen sections. Although haematoxylin and eosin staining offers a familiar picture, polychrome staining has the advantage of speed and simplicity.

Rapid Haematoxylin and Eosin (H&E) Procedure
REAGENTS REQUIRED
(See Hematoxylin)

METHOD
1 Slide mounted frozen sections (fresh or appropriately fixed) are washed briefly in water.
2 Stain in haematoxylin solution (Harris', Gill's, Mayer's or Carazzi's) for 30 seconds to 2 minutes.
3 Rinse in water.
4 If necessary, differentiate in 1% acid alcohol.
5 Wash in water.
6 Wash and blue the section in alkaline tapwater (or equivalent) for 30 seconds.
7 Counterstain in 1% eosin for 5 - 30 seconds.
8 Rinse in water.
9 Dehydrate and mount in synthetic mounting medium.

RESULTS
Nuclei - blue to blue/black
Cytoplasm, collagen - shades of pink

Toluidine Blue and Thionine
These closely related dyes offer a good and extremely rapid staining reaction for diagnostic work. The staining pattern is particularly useful for lymph node and brain evaluation due to the metachromatic qualities of the staining solutions.

REAGENTS REQUIRED
1 Toluidine Blue or Thionine 0.5 g
2 Ethanol 20 ml
3 Distilled water 80 ml
4 Phenol 0.5 g

METHOD
1 Use fixed or air dried sections.
2 Apply stain to the section for 20 seconds.
3 Rinse in water.
4 Mount in water or an aqueous mounting medium. (For permanent mount, dehydrate in acetone, clear in acetone-xylene, then in xylene and mount).

RESULTS
Nuclei - Blue
Cytoplasm, muscle and connective tissue - pink to purple

Polychrome Methylene Blue
This is a rapid metachromatic stain which is also of value in the diagnosis of frozen sections. The stain, however, requires considerable time to mature before it is ready for use.

REAGENTS REQUIRED
1 12% Aqueous potassium carbonate 8.0 ml
2 1% Aqueous methylene blue 100 ml
Mix together and boil gently for 60 minutes.
Cool to room temperature then add:
3 10% Aqueous citric acid 4.0 ml
Store in loosely stoppered bottle and allow to oxidize for 12 months before use.

METHOD
1 Use a fixed or air dried section.
2 Apply staining solution for 30 seconds.
3 Rinse in water.
4 Mount in water or aqueous mounting medium.

RESULTS
Nuclei - Blue
Cytoplasm, muscle, connective tissue - pink to purple

Alcoholic Pinacyanide23
This metachromatic stain is particularly recommended for staining thyroid sections.

REAGENTS REQUIRED
1 Pinacyanole 0.5 gm
2 70% ethyl or methyl alcohol 100 ml

METHOD
1 Use fixed or air dried section.
2 Apply stain to the section for 5 - 15 seconds.
3 Rinse in water.
4 Mount in water or aqueous mounting medium.

RESULTS
Nuclei - blue
Cytoplasm, collagen - pink
Muscle and elastic tissue - violet
Plasma cells - red
Haemosiderin - orange
Thyroid and pituitary colloid and amyloid - bright red

Phloxine - Methylene Blue - Azure B
This method gives a permanent stain with a similar appearance to H&E. It is recommended particularly for unfixed tissue where it gives good nuclear definition and clear staining.

REAGENTS REQUIRED
1 Solution A
Phloxine 0.5 g
Acetic acid 0.2 ml
Distilled water 100 ml
2 Solution B
Methylene Blue 0.25 g
Azure B 0.25 g
Borax 0.25 g
Distilled water 100 ml
3 0.2% acetic acid

METHOD
1 Rinse section briefly in water and drain.
2 Apply staining solution A for 1 minute.
3 Wash in water for 10 seconds and drain.
4 Apply staining solution B for 30 seconds.
5 Remove excess stain in 0.2% acetic acid in distilled water. Agitate the slide gently until stain ceases to flow from the section (about 20-30 seconds).
6 Differentiate in three washes of 95% ethanol.
7 Dehydrate in two changes of absolute ethanol, clear in two changes of xylene and mount in synthetic mounting medium.

RESULTS
Nuclei and bacteria - blue
Collagen and muscle - bright rose to red
Erythrocytes - scarlet

Methyl Violet for Amyloid24
This stain demonstrates amyloid deposits in frozen sections.

SECTION PREPARATION
Use unfixed sections or sections fixed in 70% methanol

REAGENT REQUIRED
1 Methyl violet 0.5 g
Distilled water 100 ml
2 1% acetic acid

METHOD
1 Apply stain to section for 1 minute.
2 Wash in two changes of distilled water for 30 seconds each.
3 Apply 1% acetic acid for 10-30 seconds.
4 Wash in tap water for 1-5 minutes.
5 Blot off excess water but do not allow to dry.
6 Mount in glycerin.

RESULTS
Amyloid - red purple

Storage of frozen sections
The suitability of cryostat sections for short or long term storage will be determined by the stability of the component under investigation which is in turn regulated by the initial fixation procedure. Fixatives such as Zamboni's or PLP are highly recommended.

Fixed, slide mounted cryostat sections can be stored temporarily in an air tight plastic container at 4C with limited loss of enzyme or antigenic activity. Unfixed sections for fluorescence microscopy may also be stored temporarily in this manner with little deterioration. Longer term storage, however, requires lower temperatures of -80C or less. Sections should be wrapped in aluminium foil and placed in an airtight container. One of the problems associated with long term storage is desiccation and denaturation of various components. Pre-treating sections with 6% polyethylene glycol, dimethyl formamide, sucrose or other cryoprotectants can prevent this change. Alternatively, wet freezing at -80C in a glycerol-sucrose mixture is recommended.

Storage of frozen tissue
A tissue sample selected for storage must be representative of the specimen, contain no necrotic areas, pose no infectious hazard and be no more than 3 mm in thickness. Once selected the sample should be frozen immediately and not be allowed to thaw before storage.

Frozen tissue must be stored below -70C using a low temperature freezer or liquid nitrogen storage unit. The lower temperature (-196C) provided by liquid nitrogen is preferred as very little degradation occurs below -130C. Short term storage at -20C is possible but not advisable. Avoid staoring tissue in a cryostat chamber as the defrosting cycle will produce thawing and refreezing of the sample. All samples must be enclosed in a protective layer of foil or plastic wrap and sealed in an airtight container to limit desiccation.

The need for high-quality unfixed tissue samples stored in an appropriate manner is particularly important in immunocytochemistry and molecular biology. Procedures such as Polymerase Chain Reaction (PCR) require preservation of intact DNA, mRNA or proteins with cellular preservation being of little importance. In contrast, immunochemistry and quantitative cytometry require retention of tissue morphology and cellular integrity. In view of these considerations, three possible freezing protocols have been proposed:22

Snap Freezing
Applicable to samples for nucleic acid and protein extraction (PCR, Southern blotting, Northern blotting, Immunoblotting). Immunocytochemical staining is possible in specimens after short term storage but long term storage should be avoided as it produces desiccation and denaturation of antigenic sites.

METHOD
1 Ensure representative tissue sample is no larger than 1.5 cm x 1.5 cm x 0.3 cm.
2 Enclose the tissue in a single layer of aluminium foil or plastic wrap to limit dessication.
3 Snap freeze the sample using liquid nitrogen, liquid nitrogen-hydrocarbon mixture, carbon dioxide-hydrocarbon slurry or carbon dioxide contact.
4 Place the frozen, foil wrapped tissue into an appropriately labelled plastic cassette.
5 Place the cassette in a small labelled plastic bag, seal and store at -70C to -196C.

Freeze in OCT
Applicable to samples for immunocytochemistry, hybridization histochemistry and flow cytometry. OCT compound (or similar) is used to protect cell integrity and limit desiccation of the sample.

METHOD
1 Ensure representative tissue sample is no larger than 0.8 cm x 0.8 cm x 0.3 cm.
2 Place the tissue in a cryomould filled with OCT.
3 Freeze by immersion in liquid nitrogen, liquid nitrogen-hydrocarbon, solid carbon dioxide-hydrocarbon slurry, carbon dioxide contact or cryostat quick freeze device.
4 Wrap the frozen OCT block in a single layer of aluminium foil to limit dessication.
5 Place the frozen, foil wrapped block into an appropriately labelled plastic cassette.
6 Place the cassette into a small labelled plastic bag, seal and store at -70C to -196C.

Fixed And Frozen
Applicable to samples for hybridization histochemistry (particularly localization of specific mRNA's) and flow cytometry.

METHOD
1 Ensure representative tissue sample is no larger than 0.8 cm x 0.8 cm x 0.2 cm.
2 Fix the specimen in 25 ml of 4% paraformaldehyde at 4C for 2 hours.
3 Discard the paraformaldehyde and replace with cold 30% sucrose. The specimen will float in the sucrose.
4 Leave overnight at 4C. The infiltrated specimen should sink to the bottom of the container.
5 Transfer the sample to a cryomould filled with OCT (or similar).
6 Freeze by immersion in liquid nitrogen, liquid nitrogen-hydrocarbon, solid carbon dioxide-hydrocarbon slurry, carbon dioxide contact or cryostat quick freeze device.
7 Wrap the frozen OCT block in a single layer of aluminium foil to limit dessication.
8 Place the frozen, foil wrapped block into an appropriately labelled plastic cassette.
9 Place the cassette into a small labelled plastic bag, seal and store at -70C to -196C.

The freezing and storage protocols outlined are not the only methods available. The value of any storage procedure must be gauged by the quality of results obtained with the stored sample once it is thawed and processed. However, attention paid to sample selection, adequate freezing procedures and a well maintained cold storage facility should provide high quality material for current and future testing.

Miscellaneous cryotechniques
Freeze substitution
The process of freeze substitution involves immersing a frozen specimen in a dehydrating agent held at low temperature. The dehydrant, which may be an organic solvent or fixative-organic solvent mixture, penetrates the tissue very slowly as the specimen thaws and gradually replaces ice within the sample. If the dehydrating agent is a fixative the specimen will also fix in the process although the degree of preservation will depend upon the temperature and nature of the fixative solution. Once dehydrated the sample may be infiltrated with paraffin wax or resin to provide a matrix for sectioning.

The main advantage of freeze substitution is the retention of water soluble substances which are held in place by freezing and then fixed in place during substitution. The quality of preservation is dependent upon many factors. Of particular importance is rapidly freezing the sample at the first stage of processing. Ideally, substitution should progress by replacement of vitrified ice by the solvent solution. The substitution should occur at a low phase change temperature to avoid recrystallization into larger ice crystals which produce intracellular damage. (The recrystallization temperature will vary with the type of specimen but is within the range of -70C to -130C for biological tissues). However, the substitution temperature will also influence the ability of the substitution agent to take up water.

At the lower limit of the melting point of the substitution fluid, viscosity is increased and penetration rate is decreased. For example, at -70C the penetration rate of ethanol is only 0.5 mm/day. Harris25 recommends the use of molecular sieves to capture the water released during substitution and improve the efficiency of the exchange. The limiting features of freeze substitution include the loss of constituents by diffusion through the gradient formed by the substituting fluid, the denaturation of protein by the action of the solvent or fixative and the swelling of the cells by the ice-fluid exchange.

A number of substitution (dehydration) agents are available,7 some of which are listed in Table 3.

Freeze-Substitution25
METHOD
1 Rapidly freeze a sample no larger than 3 mm3 in iso-pentane cooled to -170C with liquid nitrogen.
2 Transfer the frozen sample onto liquid nitrogen-cooled acetone (196C).
3 Substitute in acetone at -90C with constant agitation for 8-12 hours. Add molecular sieve to aid dehydration.
4 Warm to -60C for 8 hours, then to -30C for 8 hours.
5 Wash twice in acetone at -30C.
6 Warm to 4C in acetone.
7 Embed in:
(a) Resin: use conventional epoxy or methacrylate infiltrating procedures.
(b) Paraffin wax: chloroform at 4C for 12 hours, followed by chloroform at room temperature, wax at 54C (two changes 15 minutes each) then embed in fresh wax.

Freeze drying
The process of freeze drying is used to remove water from frozen tissue by sublimation of the solid phase at a low temperature and under vacuum. Sublimation can only occur when the partial pressure of the water vapour of the ice exceeds that of the atmosphere. In practice, the process of drying a frozen sample takes place under a vacuum of 10-3 Torr or greater and at a temperature difference sufficient to heat the ice crystals in the sample and provide energy for sublimation to water vapour. The transfer of water molecules from ice to vapour, however, removes heat from the environment causing a drop in temperature and a reduction in the rate of sublimation. A constant temperature must therefore be maintained in the sample so that sublimation proceeds at a rate equivalent to the heat input. Pearse7 suggests an optimal drying temperature of -30C to -40C.

Sublimation provides a layer of dried tissue around the periphery which inhibits further sublimation. More heat applied to overcome this barrier will cause damage whilst too little will fail to dry the centre of the sample. To overcome this inhibition sublimated water molecules must be removed from the surface of the sample. In so doing a concentration gradient between the inner and outer regions of the sample is generated and this encourages sublimation of ice from the centre of the sample. Removal of water vapour is accomplished through the use of a vapour trap (such as phosphorous pentoxide). The combined use of heating to promote sublimation and a vacuum system with appropriate vapour trap provides the conditions necessary to establish a vapour concentration gradient with continuous vaporization from the sample. The frozen-dried sample is extremely hygroscopic and rehydration occurs very rapidly unless the specimen is maintained at a temperature slightly higher than ambient. The freeze-dried tissue can be infiltrated with a resin using a low temperature.

References
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4 Barnard T. Rapid freezing techniques and cryoprotection of biomedical specimens. Scanning Microscopy 1987; 1:1227-1224

5 Lyon H. Theory and strategy in histochemistry. Berlin: Springer-Verlag. 1991

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25 Harris JR. Electron microscopy in biology - a practical approach. Oxford: Oxford University Press. 1991