section 3
After removal from the animal, tissues must be 'fixed' to preserve the living structure as closely as possible 5,6. The ideal fixative prevents tissue autodigestion (autolysis); inhibits bacterial or fungal growth (preserves); makes the tissue resistant to damage during subsequent processing, embedding and sectioning stages; is isotonic with the tissue and therefore does not alter tissue volume; does not distort any part of the tissue structure; does not dissolve tissue components and is not detrimental to the tissue component being studied.
Fixatives are divided into two main groups on the basis of their reaction with soluble proteins, namely coagulant and non-coagulant fixatives.
Coagulant fixatives, such as ethanol and mercuric chloride, are of very limited use in ISH 7,8,9,10. Ethanol dehydrates, coagulates and precipitates cellular proteins, nucleic acids and carbohydrates. It is a non-additive fixative : no covalent bonding occurs between the ethanol fixative and tissue components. Consequently, macromolecules such as mRNA are not firmly anchored within the tissue and are likely to be lost during post-fixation processing procedures.
The most commonly used fixatives for ISH are the non-coagulant, cross-linking aldehydes formaldehyde, paraformaldehyde and glutaraldehyde 11,12,13,14,15. These compounds fix tissue by introducing cross-links between different tissue components, particularly proteins, nucleic acids and lipids. Cross-linked mRNAs are stable and securely retained during subsequent tissue processing. Aldehydes are also progressive fixatives : the degree of cross-linking produced in a tissue is proportional to the fixation time.
Cross-links are generated between numerous reactive groups, but especially -NH2 groups, eg. the formaldehyde fixation of lysine residues 5 : -
-(CH2)4NH2 + H2CO ® -(CH2)4NH-H2COH
lysine side group formaldehyde ........................
-(CH2)4NH-H2CO + H2N(CH2)4- ® -(CH2)4NH-CH2-HN(CH2)4-
lysine + formaldehyde lysine side group methylene bridge
Formaldehyde is the most common fixative in routine use. It is easy to prepare and stock solutions made up in phosphate buffered saline keep for months at room temperature. Commercial formaldehyde solutions contain ~40% w/v (~12M) formaldehyde in water and 10 - 15% methanol. Formaldehyde is very water soluble and at low concentrations (4%) exists mainly as the monomer HO-(H2CO)-H. At high concentrations (40%) it gradually polymerises and precipitates as paraformaldehyde, HO-(H2CO)n-H where n is >100.
Formaldehyde causes small changes in tissue volume during fixation, especially when dissolved in phosphate buffer, and the resulting morphology is very good for light microscopy. It penetrates tissue rapidly initially, but large tissue pieces fixed by immersion in formaldehyde solutions may not be fully fixed for days.
Bouin's fixative, formaldehyde with acetic and picric acids, gives excellent morphology at the light microscope level but it has been suggested to cause some degradation of nucleic acids.
Paraformaldehyde is the most common fixative used for ISH. Tissue fixation is fast enough to prevent degradation of mRNA, but it is also easy to control the degree of fixation. Unlike formaldehyde, paraformaldehyde-containing fixative solutions have a short shelf life and need to be made up fresh each time.
Paraformaldehyde is sold as a powder, which polymerises further with age. Old paraformaldehyde powder is difficult to dissolve and will not adequately retain mRNA within the sections during ISH.
Glutaraldehyde is a very strong cross-linking di-aldehyde fixative. It is used for electron microscopy and for ISH with oligonucleotide probes14 (4% w/v glutaraldehyde and 20% w/v ethylene glycol in phosphate buffered saline). The small oligonucleotide probes can easily penetrate the densely cross-linked cytoplasmic matrix generated by glutaraldehyde fixation. Glutaraldehyde has been reported to generate artefacts with cRNA : mRNA ISH 7.
Tissues can be fixed either before or after sectioning.
Small pieces of tissue ( < 1 cm3 eg mouse embryos) can be immersed in ~ 10 x the tissue volume of fixative solution, a process called immersion-fixation. Initially, aldehyde fixatives penetrate tissue rapidly, but the rate falls with time. The centre of a large piece of tissue can take days to fix fully, with consequent degradation of tissue components such as mRNAs. Whole organs or animals can be fixed by perfusion of the vasculature with fixative solution. Such perfusion-fixation ensures rapid, even fixation of large tissue volumes.
Fresh tissue can be frozen, sectioned, recovered onto slides and then fixed.
Fixatives for ISH should ideally retain mRNA within the tissue whilst preserving tissue morphology, but should not significantly raise background. At the same time they should allow probes to penetrate the tissue and hybridise with the maximum number of target molecules. For ISH using radio-labelled cRNA probes, these apparently conflicting requirements are best met by paraformaldehyde. Using fresh-frozen sections, the S/N ratio with paraformaldehyde fixation is better than that with formaldehyde. Both are much better than glutaraldehyde.
Mouse tissues fixed with paraformaldehyde tend to become brittle. This is exacerbated by subsequent paraffin embedding and is more noticeable for some tissues than others, eg pancreas and whole eyes may be very difficult to section. Damage sustained during tissue sectioning will also tend to raise non-specific background. Formaldehyde-fixed, paraffin-embedded mouse tissues are usually much softer and consequently easier to cut. In this case, the S/N ratio obtained on formaldehyde-fixed sections may be better than that with paraformaldehyde fixation.
Both background and specific hybridisation signal are generally higher on paraffin sections of perfusion-fixed tissues than on frozen sections of the same tissue. However, both S/N ratio and tissue morphology are better with perfusion-fixation. Cross-linking of the target mRNAs before sectioning prevents mRNA digestion during sectioning and also ensures retention during tissue processing.
Fresh or fixed animal tissues need to be supported by embedding in a solid medium before thin (2 - 6 mm) sections can be cut from them.
Fresh tissue or sucrose-infiltrated, perfusion-fixed tissues can be supported by freezing in optimal cutting compound - OCT - embedding medium during sectioning. OCT is a mixture of polyvinyl alcohol and polyethylene glycol which surrounds but doesn't infiltrate the tissue. It freezes at ~ -8oC.
Quick and easy to embed tissue
Resulting frozen block is relatively easy to cut
OCT is fully water soluble
It is harder to cut serial sections from frozen blocks than from paraffin wax blocks
Once embedded, the tissue blocks need to be kept frozen
Perfusion or immersion-fixed tissues can be supported during sectioning by embedding in semi-synthetic paraffin wax. Tissues are first dehydrated through graded alcohols and then 'cleared' with an antemedium such as toluene, xylene or Histosol. Paraffin wax and ethanol are immiscible, whereas antemedia are miscible with both compounds. After clearing, the tissues are infiltrated with paraffin wax.
A range of waxes are available, whose melting temperatures are proportional to their chain length and in the range 42 - 60oC. Thin sections, ie ~ 2 mm and thicker, can be cut from tissues infiltrated wax with a melting point of 56 - 58oC.
Easy to cut thin serial sections
Once embedded, the tissue blocks can be kept indefinitely at room temperature
Once cut, tissue sections can be kept indefinitely at room temperature
Morphology often better than the equivalent frozen section.
It takes much longer to embed tissue in paraffin than in OCT and requires specialist
equipment
Tissues shrink during paraffin infiltration and may become very difficult to cut,
especially animal tissues fixed with paraformaldehyde and cleared with xylene
Tissue must be fixed before paraffin-infiltration
A variety of resins are available for embedding. They are routinely used for electron microscopy and high resolution light microscopy. Resins are rarely used for ISH although some workers have obtained good results with methacrylate embedding 16, eg of drosophila embryos
This method ensures rapid, even fixation of the entire animal. The tissues / organs of interest are dissected out for further processing after fixation. This particular protocol was designed for paraformaldehyde fixation, but can be used for any of the aldehyde fixatives.
The animal is sub-lethally anaesthetised and then perfused via the left ventricle or aorta. First, blood is washed out of the vasculature with a solution of sodium nitrite (vasodilator) and heparin (anticoagulant) in phosphate-buffered saline. Then the animal is fixed with paraformaldehyde
Anaesthetic Methoxyfluorane, Abbot Laboratories, Sydney, - or similar inhalation anaesthetic. Alternatively, an injectable anaesthetic such as Avertin can be used Sodium nitrite as powder, Ajax Univar, Catalogue No 492, (500g or less) Preservative-free sodium heparin Fisons, Thornleigh, NSW Phosphate buffered saline as Phosphate Buffered Saline Tablets Dulbecco 'A' (Magnesium and Calcium-free), Oxoid, Unipath, Catalogue No BR14a, boxes of 100 tablets Paraformaldehyde Extra Pure Paraformaldehyde powder, Merck, Catalogue No 4005.1000 (500g or 1 kg) 70% alcohol Reagent grade, BDH
Perfusion apparatus 'Home made' version is 2 x 20 ml syringes, one full of nitrite and one full of fixative, attached via a 3-way tap to a soft plastic catheter the same external diameter as the internal diameter of the aorta of a mouse. If using an inhalation anaesthetic, use a nose jar containing a pad of anaesthetic-saturated cotton wool during the perfusion
1 Make up fresh NaNO2 - heparin - PBS solution. Mix 1 part PBS in MilliQ water with 1 part 1% (w/v) NaNO2 in MilliQ water. Add heparin to a final concentration of 10 units / ml. Leave at room temperature prior to use 2 Make up fresh 4% (w/v) paraformaldehyde in PBS. For 100 ml of fixative, dissolve 1 PBS tablet in 100 ml of MilliQ water and then heat to 65oC in a microwave oven. In a fume hood, add 4 g of paraformaldehyde powder and shake. The paraformaldehyde powder should go into solution fully in ~ 1 - 2 minutes. Cool to room temperature before use
1 Sublethally anaesthetise the mouse with Methoxyfluorane or equivalent inhalation anaesthetic in a killing jar 2 When almost dead, remove mouse from the jar and pin out on a board with an anaesthetic-filled nose jar in place 3 Open the thorax by completely removing the ribs and sternum to fully expose the heart and lungs 4 Make a small incision in the lateral wall of the left ventricle halfway between the atrio-ventricular groove and the apex of the heart. Insert the catheter up into the aortic outflow tract taking care not to damage the mitral valve 5 Clamp the catheter in place with a small pair of forceps and then remove the right atrium 6 Flush the animal with NaNO2 - heparin - PBS solution until the effluent from the right atrium is clear 7 Fix with 4% paraformaldehyde in PBS until the animal is stiff 8 Remove the organs of interest and post-fix by immersion in fresh fixative solution for 2 - 12 hours 9 Process through to paraffin wax the next day OR store in 70% ethanol
1 Make ~ 5 ml NaNO2 - heparin - PBS solution per mouse to be perfused. The final solution is 1/2 strength PBS and 0.5% sodium nitrite, which is roughly isotonic 2 Make ~ 25 ml 4% paraformaldehyde in PBS per mouse to be perfused. The pH of the final solution (at 22oC) should be that of the PBS ie 7.2. If the paraformaldehyde powder does not dissolve initially, heat the solution briefly or add a few drops of sodium hydroxide. If it still will not dissolve, get a fresh stock of paraformaldehyde. Old paraformaldehyde does not fully dissolve nor does it fix well enough for ISH 3 An injectable anaesthetic such as Avertin works just as well as Methoxyfluorane 4 If the mitral valve is damaged by the incision or during insertion of the catheter, perfusion of the pulmonary rather than the systemic circulation occurs. Unless this is an important animal, or you are interested in the lungs, start again 5 Inject the nitrite and fixative very slowly to prevent oedema / damage to the organs 6 Cells left in the blood vessels become 'cross-linked' in situ on contact with fixative. They occlude the vessel lumen and prevent further perfusion with fixative solution 7 With a good perfusion, the tail and hind legs will go very stiff within a few minutes. If only the fore legs go stiff, some head and neck organs may be adequately fixed for ISH 8 Use the equivalent of ~ 10 x the tissue volume of fixative for post-fixation 9 The post-fixation time is determined largely by the tissues harvested. A minimum time of ~2 hours ensures that the tissue survives paraffin-infiltration. Longer post-fixation times improve morphology but also make the tissue more brittle and much more difficult to cut. 2 - 4 hours post-fixation gives good morphology and it is possible to section tissues such as whole eye and pancreas. Other tissues, such as spleen may require up to 8 hours post-fixation but are consequently become very brittle 10 If the tissue cannot be paraffin-infiltrated immediately, it can be stored for long periods of time in 70% ethanol. Do not leave it in the fixative or store in PBS
Instead of perfusing the animal with fixative solution, small pieces of tissue (< 1 cm3) can be fixed by simple immersion in 4% paraformaldehyde in PBS, made up as in 3.2.1 above. This method is often reserved for small mouse embryos which cannot be perfusion-fixed
Dissect out the tissue and immerse in the equivalent of ~ 10 x the tissue volume of fixative. Fix overnight - either at room temperature or at 4oC - and process through to paraffin wax the next day OR store in 70% ethanol
The fixed tissue is dehydrated through graded alcohols, cleared with an antemedium and then infiltrated with paraffin wax.
Alcohols Reagent grade 70% and absolute alcohols, BDH. Make up 80% and 90% stocks with distilled water Clearing agent Reagent grade Histosol, xylene (sulphur-free) or toluene, BDH Paraffin wax Paraplast 58oC melting point paraffin wax
Processing machine or Histokinette Plastic cassettes for tissue processing
1 Place the fixed tissue in plastic processing cassettes 2 Process using the Histokinette : 70% alcohol 21/2 hours 22oC 70% alcohol 21/2 hours 22oC 80% alcohol 1 hour 22oC 90% alcohol 1 hour 22oC Absolute alcohol 1 hour 22oC Absolute alcohol 11/2 hours 22oC 50% (v/v) alcohol in toluene 1 hour 22oC 100% toluene 1 hour 22oC 100% toluene 1 hour 22oC Paraffin wax 1 hour 60oC Paraffin wax 1 hour 60oC S = 14 1/2 hours processing time 3 Either embed the tissue immediately, or wrap the tissue in its plastic processing cassette in aluminium foil and store at room temperature
1 This protocol is only a guide, but it has worked very well for ISH using paraformaldehyde-fixed mouse and rat tissues, including pancreas, spleen and whole eyes 2 For small tissue pieces, shorter processing cycles can be used 3 All processing except the paraffin wax infiltration is best performed at room temperature 4 Histosol and xylene clearing agents make mouse tissues very brittle. Toluene is much more gentle on the tissue although it is probably more toxic to the operator - carcinogenic - handle with care 5 Mouse tissues do not usually have to be defatted : if this is necessary, use chloroform
After the tissue is fully infiltrated with paraffin wax, it is embedded in excess wax in a square block which acts as a support during sectioning. Using an embedding centre, a metal mold on a hot plate is filled with molten paraffin wax. The tissue is orientated appropriately, making sure that there are no air bubbles trapped between the tissue and the molten wax. The wax-filled mold containing the tissue is then cooled on a cold plate. The block is now ready to section but can be stored indefinitely at room temperature
5 - 6 mm sections are cut with a microtome, floated onto a water bath and then recovered onto silane-coated slides, ready for ISH
Microtome for paraffin blocks Forceps Cold water bath Non-coated slides Water bath set at 42oC AES-coated slides
1 Cool the block to be cut on ice 2 Pre-heat the water bath to 42oC
1 Cut a ribbon of sections at 5 - 6 mm 2 If the sections are very crumpled, pick up the ribbon with forceps and float it first onto the surface of a cold water bath. Tease out the wrinkles with forceps 3 Detach individual paraffin sections from the ribbon, collect the sections onto non-coated slides and transfer them to the 42oC water bath 4 Recover individual sections on AES-coated slides. Multiple sections can be placed on a single slide 5 Dry the sections onto the slides at 37oC. If desired, then melt the paraffin wax at 60oC
1 Make sure that the water bath is clean and free from fragments of tissue and paraffin, or bacteria. These will all stick to the slides and can generate spurious hybridisation 'signals'. Fill the water bath with distilled water and heat it to 42oC. Hotter water will stretch the section or even melt the paraffin 2 A section of 5 - 6 mm should contain enough mRNA to generate a hybridisation signal, but will also be thin enough to give reasonable resolution 3 The ribbon of sections can be floated directly onto the warm water bath if there are few wrinkles 4 When using the cold water bath, a few drops of 70% alcohol placed directly under a badly wrinkled section will help stretch it out 5 If AES-coated slides are used to recover sections from the cold water bath, the sections will stick to the slide and can't float off onto the warm water 6 It is easy to use two different probes eg sense and antisense, on two different sections on the same slide. The only proviso is that there is enough 'land' between the sections for separate coverslips 7 Melting the wax at 60oC after drying the sections doesn't seem to affect the subsequent hybridisation signal, even if the slides have been stored for long periods of time after cutting
After air drying or melting the paraffin wax, the sections can be stored indefinitely at room temperature in a dust-free container eg Kartel Slide Containers for 100 slides
The animal is perfusion-fixed and the organs dissected free as per 3.2.1 pg 13, or small tissue pieces are immersion-fixed as per 3.2.2 pg 15. The paraformaldehyde fixed-tissue can be embedded in OCT, frozen and then sectioned using a conventional cryomicrotome (cryostat). However, before it is embedded and frozen, the tissue must be infiltrated with sucrose which acts as a cryoprotectant
Sucrose Analar grade, BDH, Product number 10274 (500 g) Sodium dihydrogen orthophosphate NaH2PO4.H2O - BDH, Product number 10245 (500 g) Disodium hydrogen orthophosphate Na2HPO4 - Analar Grade, BDH, Product number 10249 (500 g) Sodium azide Analar Grade, BDH, Product number 10369 (100 g)
1 Make up fresh 100 mM sodium phosphate buffer (PB) For 100 ml of PB pH 7.2, add 6.84 ml of 1M Na2HPO4 and 3.16 ml of 1M NaH2PO4 to 90 ml of MilliQ water. Autoclave the solution and check the pH (at 22oC) before use 2 Make up the sucrose infiltration solution : 20% (w/v) sucrose in 10 mM PB pH 7.2 For 100 ml, add 20 g of sucrose to 10 ml of 100 mM PB pH 7.2 and make up to a final volume of 100 ml with sterile MilliQ water. Add sodium azide to a final concentration of 0.1% (w/v)
1 Immerse the tissue in the equivalent of ~ 10 x the tissue volume of sucrose solution : it will float initially and then sink when it is fully infiltrated 2 The tissue is now ready to freeze-embed in OCT, although it can be stored at 4oC in sucrose for short periods of time
1 Lots of things grow in sucrose - use sterile solutions where ever possible 2 There is a wealth of superstition surrounding sucrose infiltration. The concentration of sucrose used by different workers varies from 5% - 30%. It is dissolved in water, phosphate buffer or phosphate-buffered saline. Some of these solutions eg 5% sucrose in water, will be hypotonic and cause the tissue to swell. Others eg 30% sucrose in PBS, are hypertonic and cause the tissues to shrink. 20% sucrose in phosphate buffer should be roughly isotonic Various protocols using graded (5, 10, 15 and 20%) sucrose solutions are also used. Anecdotally, they may be beneficial for fragile tissues such as retina 3 Azide is very toxic 4 Prolonged storage of tissue in sucrose increases the chance of it becoming infected with bacteria or fungi. Also, if the fixation was inadequate, the tissue will slowly disintegrate to form a sludge on the bottom of the container
The sucrose-infiltrated tissue is first orientated and then frozen in OCT embedding medium which acts as a support during sectioning
Embedding medium Optimal Cutting Temperature (OCT) embedding medium, Tissue Tek, Miles Diagnostic Division Catalogue number 4583 (4 fl oz) Specimen molds Cryomold Disposable Vinyl Specimen Molds, Tissue Tek, Miles Diagnostic Division Standard size ® 25 mm x 20 mm x 5 mm Intermediate size ® 15 mm x 15 mm x 5 mm Biopsy size ® 10 mm x 10 mm x 5 mm Boxes of 100 molds Aluminium foil instead of Cryomolds Dry ice, liquid nitrogen or cryostat for freezing tissue blocks
1 Remove the tissue from the sucrose solution and blot off excess liquid 2 Select a cryomold slightly bigger than the tissue and fill it ~2/3 full with OCT, taking care not to introduce any air bubbles. Alternatively, fashion a mold out of aluminium foil 3 Place the tissue in the OCT and orientate it for cutting. Make sure there is no air trapped between the tissue and the OCT 4 Top up the mold with OCT to cover the tissue 5 Place the OCT / tissue-filled mold either on dry ice, on the surface of the liquid nitrogen or in the cryostat, and freeze it slowly from the bottom up 6 When fully frozen, the block can either be cut or stored at -70oC
1 Excess sucrose will form a casing around the tissue causing the tissue and OCT to separate during sectioning 2 Any air bubbles in the OCT or between the tissue and OCT will make sectioning more difficult 3 Aqueous solutions expand during freezing. Therefore freezing the block from the bottom allows the tissue to expand and avoids both crush artefacts and cracked blocks 4 If the block is to be cut immediately, freeze it in the cryostat in which it is to be cut, not on dry ice
5 - 6 mm sections are cut with a cryostat, recovered onto AES-coated slides and air-dried ready for ISH, or storage
Cryostat and disposable blades / knife and anti-roll plate Brushes AES-coated slides
1 If the block has been stored at -70oC, place it in the cryostat chamber or in a -20oC freezer for ~ 2 hours before sectioning 2 Cut sections at 5 - 6 mm 3 Smooth out the wrinkles in the section using brushes on the OCT around the edge of the section rather than the section itself 4 Recover the section by melting onto an AES-coated slide 5 Air-dry for 1 - 2 hours
1 The block should be the same / similar temperature to the cryostat in which it is to be cut. If it is too cold, the sections shatter 2 Although 5 - 6 mm sections are a good compromise between resolution and hybridisation signal, individual blocks may cut better either slightly thicker (decrease resolution) or thinner (decrease signal). The optimal section thickness for each block has to be determined empirically 3 For cutting individual sections, disposable blades are easier to use than a knife. For cutting serial frozen sections, it is better to use a knife and an anti-roll plate 4 Air-drying allows the sections to become firmly attached to the slide
After air-drying, the sections can be stored desiccated at -20oC for several months eg in black plastic containers for 25 slides. The S/N ratio gradually deteriorates over time
Fresh, unfixed tissues can be frozen in OCT and then sectioned using a conventional cryomicrotome (cryostat) before they are fixed.
The animal is killed and the organs dissected free as quickly as possible, before freeze-embedding
Anaesthetic Methoxyfluorane, Abbot Laboratories, Sydney, - or similar inhalation anaesthetic. Alternatively, an injectable anaesthetic such as Avertin can be used n-Hexane reagent grade, BDH Dry ice Embedding medium Optimal Cutting Temperature (OCT) embedding medium, Tissue Tek, Miles Diagnostic Division Catalogue number 4583 (4 fl oz)
Specimen molds Cryomold Disposable Vinyl Specimen Molds, Tissue Tek, Miles Diagnostic Division Standard size ® 25 mm x 20 mm x 5 mm Intermediate size ® 15 mm x 15 mm x 5 mm Biopsy size ® 10 mm x 10 mm x 5 mm Boxes of 100 molds Aluminium foil
Make an aluminium foil mold which is slightly larger than the vinyl Cryomolds. In a fume hood, place the mold onto dry ice and fill it with a slurry of crushed dry ice and n-hexane. Add enough n-hexane to just cover the surface of the dry ice. Make the slurry ~ 30 minutes before harvesting the organs - it needs time to cool down to ~ -80oC
1 Kill the animal and dissect the organs free as quickly as possible 2 Select a cryomold slightly bigger than the tissue and fill it ~2/3 full with OCT, taking care not to introduce any air bubbles. Alternatively, fashion a mold out of aluminium foil 3 Place the tissue directly into the OCT and orientate it for cutting. Make sure there is no air trapped between the tissue and the OCT 4 Top up the mold with OCT to cover the tissue 5 Place the OCT / tissue-filled mold on the surface of the n-hexane-dry ice slurry and freeze it from the bottom up. Make sure that the n-hexane does not seep into the block during freezing 6 When fully frozen, the block can be stored at -70oC for months 8 Before the tissue can be sectioned, the block has to be warmed up to ~ -20oC. This will take ~ 2 hours in the chamber of the cryostat or in a -20oC freezer
1 n-hexane is a neurotoxin. Use it in the fume hood 2 Any method can be used to kill the animal. If the brain, brain stem or spinal cord are needed, don't use cervical dislocation 3 Cellular mRNA will continue to degrade, after the animal is killed, until the tissue is frozen. The faster the organs are frozen, the better the hybridisation signal will be. This is particularly noticeable for organs rich in RNases such as pancreas 4 There is no need to wash the tissue in PBS first. The PBS will also tend to form a 'casing' around the tissue and cause it to separate from the OCT during sectioning. Blot off any excess blood 5 Aqueous solutions expand during freezing. Therefore freezing the block from the bottom allows the tissue to expand and avoids both crush artefacts and cracked blocks
5 - 6 mm sections are cut with a cryomicrotome (cryostat), recovered onto AES-coated slides and frozen back onto the slides. The frozen sections are allowed to fully adhere to the slides before being paraformaldehyde-fixed, ready for ISH or for storage
Phosphate buffered saline as Phosphate Buffered Saline Tablets Dulbecco 'A' (Magnesium and Calcium-free), Oxoid, Unipath, Catalogue No BR14a, boxes of 100 tablets Paraformaldehyde Extra Pure Paraformaldehyde powder, Merck, Catalogue No 4005.1000 (500g or 1 kg). Alcohols Reagent grade 70%, 95% and absolute alcohols, BDH Sterile MilliQ water
Cryostat and disposable blades / knife and anti-roll plate Brushes AES-coated slides Dry ice Slide racks Plastic slide racks NOT used for routine staining. Black plastic rack for 25 slides, H.D Scientific, Catalogue number VI 991-99 Trough to fit slide racks Plastic troughs NOT used for routine staining. Black plastic troughs to fit slide rack for 25 slides, H.D Scientific, Catalogue number VI 992-99 Clean plastic containers for graded alcohols which will hold a 9 cm x 7.5 cm x 3 cm black plastic slide rack, eg 500 ml freezer containers for food
1 If the block has just been made by freezing on n-hexane and dry ice, or it has been stored at -70oC, place it in the cryostat chamber or in a -20oC freezer for ~ 2 hours before sectioning 2 Make up 200 ml of fresh 4% (w/v) paraformaldehyde in PBS as per 3.2.1 pg 14 3 Make up 200 ml of PBS. Dissolve 2 tablets of PBS in 200 ml of sterile MilliQ water and re-autoclave
1 Cut sections at 5 - 6 mm 2 Smooth out the wrinkles in the section with brushes. Use the OCT around the edge of the section rather than the section itself 3 Recover the section by melting onto an AES-coated slide 4 Re-freeze the melted section as fast as possible using dry ice 5 Place the slides in a rack in the cryostat chamber and leave for ~ 30 minutes 6 Fix in 4% paraformaldehyde in PBS for 30 minutes at 22oC 7 Rinse in PBS 3 minutes 22oC 8 Rinse in sterile MilliQ water 3 minutes 22oC 9 Dehydrate through graded ethanols : 70% alcohol 3 minutes 22oC 95% alcohol 3 minutes 22oC Absolute alcohol 3 minutes 22oC Absolute alcohol 3 minutes 22oC 10 The sections can either be air-dried, ready for ISH, or they can be stored
1 The block should be the same / similar temperature to the cryostat in which it is to be cut. If it is too cold, the sections shatter 2 Although 5 - 6 mm sections are a good compromise between resolution and hybridisation signal, individual blocks may cut better either slightly thicker (decrease resolution) or thinner (decrease signal). The optimal section thickness for each block has to be determined empirically 3 For cutting individual sections, disposable blades are easier to use than a knife. For cutting serial frozen sections, it is better to use a knife and an anti-roll plate 4 mRNAs can degrade when the section is melted. Any delay in re-freezing reduces the subsequent hybridisation signal 5 The 30 minute interval between sectioning and fixation ensures that the sections become firmly stuck to the slide 6 Unfixed frozen sections can be stored desiccated at -20oC but cellular mRNAs are very vulnerable should the sections thaw before fixation - not recommended 7 Using the black plastic troughs, 200 ml (150 ml minimum) of solution completely covers the slides in the black plastic slide racks. The troughs are used for fixation and PBS and water washing of sections - a minimum of 3 8 Any contaminating RNases in the fixative solution will be fixed along with the tissue. In all subsequent steps, care must be taken to use sterile solutions. Where possible, make them up with sterile MilliQ water and then re-autoclave 9 The PBS wash removes excess fixative and the MilliQ water wash removes excess salt
After ethanol dehydration, the sections are ready to be hybridised. Alternatively, they can be stored over absolute ethanol at -20oC for several months. The S/N ratio gradually deteriorates over time
5 J. R. Baker (1966) Cytological Technique : the principles underlying routine methods 6 A. G. E. Pearse ( 1961) Histochemistry. Theoretical & Applied 7 J. H. Pringle, P. V. Senior and A. Warford (1989) In situ hybridisation to cellular mRNA using single-stranded cRNA probes labelled with 35S In situ hybridisation course. Course manual. Revised 1990 8 R. H. Singer (1986) Biotechniques 4:230 9 A. T. Haase, L. Stowring, J. D. Harris, B. Traynor, P. Ventura, R. Peluso and M. Brahic (1982) Virol 119:399-410 10 A. R. Jilbert (1986) Histochem 85:505 11 J. P. Coghlan, P. Aldred, J. Haralambidis, H. D. Niall, J. D. Penschow and G. W. Tregear (1985) Hybridization histochemistry Anal Biochem 149:1-28 12 J. B. Lawrence and R. H. Singer (1985) Quantitative analysis of in situ hybridization methods for the detection of actin gene expression Nucleic Acids Res 13:1777-1799 13 P. V. Senior, D. R. Critchley, F. Beck, R. A. Walker and J. M. Varley (1988) The localization of laminin mRNA and protein in the postimplantation embryo and placenta of the mouse: an in situ hybridization and immunocytochemical study Development 104:431-446 14 J. D. Penschow, J. Haralambidis, S. Pownal and J. P. Coghlan (1989) The location of gene expression by hybridisation histochemistry using oligodeoxyribonucleotide probes Methods in Neurosciences 1 (Gene Probes):222 - 238 15 B. E. Faulkner-Jones, D. S. Cram, J. Kun and L. C. Harrison (1993) Localisation and quantitation of expression of two glutamate decarboxylase genes in pancreatic b-cells and other peripheral tissues of mouse and rat Endocrinol 133:2962 - 2972 16 M. Jamrich (1984) EMBO J 3:1939 - 1943