Tissue architecture and morphology must be preserved to enable histological or cytological investigations - a process known as fixation. An ideal fixative should stabilise and protect tissues and cells from damage in any onward processing steps. Prompt fixation is essential to achieve consistent results. Poor or delayed fixation can result in loss of antigenicity or diffusion of antigens into the surrounding tissue.
There is no one ideal fixative and the best preservation method may depend on the antigen in question. Usually there is a compromise to find between tissue morphology and antigen presentation. There are two main methods that cover most antigens between them – namely formalin fixed paraffin embedded (FFPE) and frozen sections.
Fixation, processing and microtomy can account for many artefacts in histology and immunohistochemistry. This paper is a good review of artifacts and causes for general histology.
Frozen sections are required for a number of antigens, but the tissues are not as easy to handle as paraffin blocks. Frozen sections might not have as good a tissue morphology and poorer resolution at higher magnifications compared to paraffin sections, however they are the fastest way of producing stainable sections. Tissues to be frozen may require special collection procedures and, as the freezing process does not fix the tissue, any pathogenic agents (such as potentially harmful micro-organisms) may not have been inactivated. Frozen sections can also be harder to cut. However, some antigens, such as T Cell Subset antigens, do not survive formalin fixation and paraffin embedding.
This method varies as fixation occurs after embedding and sectioning the tissue. Small pieces of tissue are snap frozen in pre-cooled isopentane in liquid nitrogen, or embedded in OCT and frozen to facilitate cutting. Blocks must be kept frozen during cutting and acetone is used to help fix the sections once mounted on slides (Choice of subbed slides is discussed here). The acetone fixation does not truly protect the sections against the detrimental effects of long incubations in aqueous solutions. Thorough drying of sections before and after acetone fixation can help prevent morphological changes such as chromatolysis and loss of nuclear membranes which can be an issue with acetone fixed frozen section through long IHC protocols.
Neutral buffered formalin (NBF), formal saline, paraformaldehyde (4% PFA) and 10% formalin in tap water are the most widely used fixatives and share a similar fixation action. Formalin fixation is popular as it seems to balance morphology and antigen preservation for a wide variety of samples. A down side of formalin fixation is that proteins can be cross linked, sequestering the desired antigen. There are a variety of antigen unmasking techniques that can be employed to improve the accessibility of the antibody to its antigen again, although not all antigens can be retrieved. Formalin induced autofluorescence can be an issue for researchers wishing to use immunofluorescent methods, however there are some reagents that can be used to tackle this issue. (See autofluorescence)
Shop Formalin Solutions
Mercuric chloride based fixatives, e.g. B5
These seem to offer the best fixation for lymph node biopsies, offering excellent cytoplasmic immunoglobulin demonstration, however surface membrane immunoglobulin is not readily demonstrated following B5 fixation. B5 fixation may also lead to higher levels of background staining.
Periodate lysine paraformaldehyde (PLP)
This fixative is designed to protect carbohydrate moieties associated with cell membranes against the damaging effects of tissue processing and embedding. This can be modified by the addition of 5% potassium dichromate solution (PLP/D) which tries to also preserve the lipid elements, however the morphology is not ideal with this fixation.
The time of fixation is dependent on the size of the specimen to be fixed. If sections are in the fixative for a long time, this may require adaption of the antigen retrieval steps.
Inadequate fixation can manifest itself during staining. Known as zonal fixation, it may be that antigens appear to fade towards the centre of the block. The first sections off a block may appear fine, but as sections from the centre of the block are reached, the antigen may appear to be marginalised round the outer edges of the section, however over fixation can also induce artefacts.
Where possible, tissues should be cut into smaller pieces to enable the fixative to penetrate to the centre before the outer edges are over fixed.
If the sample is known to contain bone, then a decalcification step here may be easiest before embedding. However, tissue blocks can be run back through xylenes & alcohols to decalcify if needed. There are a number of methods for decalcification, which can also affect the sample for downstream assessment. EDTA is a slow chelating method of decalcification, but seems to leave the tissue morphology intact, although too long an exposure can start to destroy nuclear detail, but preserves the DNA/RNA. Strong acid based methods are quicker, but can damage DNA/ RNA, so are not suitable if it is planned to test for these elements later. Some labs may use a combination of these methods for speed. If fixation is not complete, a separate decalcification step may demonstrate swelling of the tissue and a later lack of staining as antigens are lost to necrosis. Some fixative solutions also contain decalcifying reagents too to help with this issue. It is worth checking if there are any contraindications on decalcification if you are doing any clinical diagnostic testing. It is worth validating your decalcification method to see if, and how, it may affect results with any methods and antibodies being used.
The tissue specimens that have been fixed require embedding in media to facilitate cutting.
With the above fixative solutions, sections are most commonly dehydrated through a series of graded alcohols. Tissue processing is finished by clearing the tissue in an organic solvent such as chloroform, xylene or toluene before impregnating with paraffin wax, such as Ralwax I and cooled to make a sectionable block. The paraffin wax should be of good quality so that a ribbon can be cut on a microtome, ready to put on slides.
Choice of subbed slides is discussed here, but paraffin sections need to fully dry on the slide - this can be by hot plating at 60°C for 30 minutes in busy labs – however some antigens, such as PCNA, can be lost at the higher temperature so should be dried at 37°C, or sections can be dried overnight at room temperature.
This paper discusses some of the effects of fixation and slide preparation on different antigens.
Paraffin wax is not suitable as an embedding media for some microscopical techniques however, such as where ultrastructural studies require very thin sections for study under an electron microscope.
As there has been a dehydration protocol during tissue processing for paraffin embedding, tissues must be brought back to the aqueous phase before staining can commence. (See dehydration and rehydration)
Resin embedded sections can offer improved resolution of cellular detail, less tissue shrinkage, lower temperatures in the embedding process and removes the need for a decalcification step for calcified material. Sections can be fixed as above.
There are a number of resins that can be used, however the properties of resins are complex with a wide range of physical & chemical properties possible. These fall roughly into 2 groups for histology:- Acrylic and Epoxy resins. For light microscope techniques, acrylic resins are best, but there are many variations both in chemistry and polymerisation method that can affect the tissues’ suitability for staining or even sectioning if the resin is too hard. Polymerisation of resins can depend on the monomer, softener/ plasticiser, cross linker, catalyst and accelerator (be this heat, light, radiation or chemical) and affect the suitability of tissues for further experiments. For instance, heat is not recommended to initiate tissues for enzyme histochemical studies.
Popular acrylic resins include those based on Glycolmethacrylate (GMA), Polyhydroxy Aromatic Dimethacrylate and Methyl methacrylate. Lowicryl resins are more suited to Electron Microscopy than Light Microscopy techniques. The ability of acrylic resins to polymerise at low temperatures has enabled avoidance of prolonged heating that can be detrimental to some antigens.
Glycolmethacrylate resins can be good for general Light Microscopy, but are not recommended for IHC techniques, as the polymerised glycolmethacrylate seems to form an irremovable barrier to subsequent procedures.
A Modified methyl methacrylate resin can produce comparable staining to paraffin sections with improved morphology and resolution. The resin can be removed prior to staining by treating with warmed (37°C) Xylene for 30 minutes.
When known, for immunohistochemistry, the requirements of the primary antibodies to be used might give an idea of the most appropriate fixative or embedding method to use.
Traditionally, frozen sections have been preferred for immunofluorescence, due to the formalin induced fluorescence often associated with paraffin sections, however there are now kits available to block this type of autofluorescence.
For new antibodies, a variety of fixatives, embedding materials and antigen unmasking techniques should be tested, to see which offers the best antigen and tissue preservation.
Some antibodies can be used on either paraffin sections (with or without pre-treatment) or frozen sections. For multiple labelling, all antibodies need to be compatible with the same fixative and embedding method. There is a little more flexibility on antibodies that do, or do not, require antigen unmasking in the same method, but please contact us to discuss these combinations on an individual basis.
For in situ hybridization, frozen sections or paraffin sections can be used, although further pre-treatments may be required to prepare them for hybridization. Further fixatives can be used, but it is recommended to avoid solutions that may contain RNases.