In histology, pathology and cell biology, fixation is the preservation of biological tissues from decay due to autolysis or putrefaction. It terminates any ongoing biochemical reactions and may also increase the mechanical strength or stability of the treated tissue. Tissue fixation is a critical step in the preparation of histological sections, and its broad objective is to preserve cells and tissue components in a form that allows thin, stained sections to be prepared. This makes it possible to study tissue structure, which is determined by the shapes and sizes of macromolecules such as proteins and nucleic acids inside and around cells.
In their protective role, fixatives denature proteins by coagulation, by forming additive compounds, or by a combination of coagulation and additive processes. A compound that adds chemically to macromolecules stabilises structure most effectively when it can combine with parts of two different macromolecules, an effect known as cross-linking.
Tissue is fixed for several reasons. One is to kill the tissue so that postmortem decay, autolysis and putrefaction, is prevented.[1]
Fixation preserves biological material, tissue or cells, as close to its natural state as possible during preparation for examination. Several conditions usually must be met to achieve this.
First, a fixative usually disables intrinsic biomolecules, particularly proteolytic enzymes, which would otherwise digest or damage the sample.
Second, a fixative typically protects a sample from extrinsic damage. Fixatives are toxic to most common microorganisms, especially bacteria, that might exist in a tissue sample or colonise the fixed tissue. Many fixatives also alter the fixed material chemically to make it less palatable, either indigestible or toxic, to opportunistic microorganisms.
Finally, fixatives often alter cells or tissues at the molecular level to increase their mechanical strength or stability. This increased strength and rigidity helps preserve the morphology of the sample as it is processed for further analysis.
Even the most careful fixation alters the sample and introduces artefacts that can interfere with the interpretation of cellular ultrastructure. A prominent example is the bacterial mesosome, once thought to be an organelle of gram-positive bacteria in the 1970s but later shown by new electron microscopy methods to be an artefact of chemical fixation.[2][3] Standardisation of fixation and other processing procedures takes this into account by establishing which procedures introduce which kinds of artefacts. Researchers who know what artefacts to expect for each tissue type and technique can interpret sections accurately or choose methods that minimise artefacts in the region of interest.
Fixation is usually the first stage in a multistep process that prepares biological material for microscopy or other analysis. The choice of fixative and protocol may therefore depend on the additional processing steps and final analyses that are planned. Immunohistochemistry, for example, uses antibodies that bind a specific protein target, and prolonged fixation can chemically mask those targets and prevent antibody binding. In such cases a quick fix with cold formalin for about 24 hours is typically used. One hundred per cent methanol can also be used for quick fixation, and the time varies with the material. MDA-MB 231 human breast cancer cells, for instance, can be fixed for only 3 minutes with cold methanol at −20 °C. For enzyme localisation studies, tissues should be lightly pre-fixed only, or post-fixed after the enzyme activity product has formed.
There are generally three types of fixation process, depending on the sample that needs to be fixed.
Heat fixation is used for single-celled organisms, most commonly bacteria and archaea. The organisms are usually mixed with water or physiological saline to spread the sample evenly, and the diluted sample is spread onto a microscope slide. A diluted bacterial sample on a slide is commonly called a smear. After the smear dries at room temperature, the slide is held with tongs or a clothespin and passed through a Bunsen burner flame several times to heat-kill the organisms and make them adhere to the slide. A microincinerating device can also be used. After heating, samples are usually stained and imaged with a microscope.[4] Heat fixation generally preserves overall morphology but not internal structures. Heat denatures proteolytic enzymes and prevents autolysis. Heat fixation cannot be used for capsular staining, because heat shrinks or destroys the capsule, the glycocalyx, so that it cannot be seen.[5]
Immersion can fix histological samples ranging from a single cell to an entire organism. The tissue is immersed in fixative for a set period, and the fixative volume must be at least 10 times the tissue volume.[6] For fixation to succeed, the fixative must diffuse throughout the tissue, so tissue size and density, as well as the type of fixative, must be considered. Immersion is common for cellular applications but can also be used for larger tissues. A larger sample must be immersed longer for the fixative to reach deeper tissue.[7]
Perfusion is the passage of fluid through the blood vessels or natural channels of an organ or organism. In perfusion fixation, fixative is pumped into the circulatory system, usually through a needle inserted into the left ventricle. This can be done under ultrasound guidance or by opening the chest cavity.[8] The fixative is injected into the heart with a volume matching the typical cardiac output. Using the innate circulatory system, the fixative is distributed throughout the body, and the tissue does not die until it is fixed. A drainage port must also be added somewhere in the circulatory system to accommodate the added volume of fixative and buffer, typically in the right atrium. Fixative is pumped until it has replaced all of the blood. Perfusion preserves morphology well,[9] but the subject dies and the volume of fixative needed for larger organisms is high, which can raise costs. The volume can be reduced by clamping arteries that supply tissues not of interest. Perfusion fixation is commonly used for imaging brain, lung and kidney tissue in rodents, and it is also used in human autopsies.[7][10]
In both immersion and perfusion fixation, chemical fixatives preserve structures in a state as close to living tissue as possible, both chemically and structurally. This requires a chemical fixative.
Cross-linking fixatives create covalent chemical bonds between proteins in tissue. This anchors soluble proteins to the cytoskeleton and lends additional rigidity to the tissue. Preservation of transient or fine cytoskeletal structure, such as contractions during embryonic differentiation waves, is best achieved by pre-treatment with microwaves before adding a cross-linking fixative.[11][12]
The most commonly used fixative in histology is formaldehyde, usually as 10 per cent neutral buffered formalin (NBF), which is about 3.7 to 4.0 per cent formaldehyde in phosphate buffer at pH 7. Because formaldehyde is a gas at room temperature, formalin, a solution of about 37 per cent w/v formaldehyde gas in water, is used to make the fixative. Formaldehyde fixes tissue by cross-linking proteins, primarily the residues of the basic amino acid lysine. Its effects are reversible by excess water, and it avoids formalin pigmentation. Paraformaldehyde is also common and depolymerises back to formalin when heated, which makes it an effective fixative. Its benefits include long-term storage and good tissue penetration, and it is particularly good for immunohistochemistry. Formaldehyde vapour can also fix cell smears.
Another popular aldehyde is glutaraldehyde. It acts similarly to formaldehyde and deforms the alpha helices of proteins. Glutaraldehyde is a larger molecule than formaldehyde, however, so it permeates membranes more slowly, and fixation of thicker tissue can be difficult unless the sample is trimmed. One advantage of glutaraldehyde is that it can give a more rigid or tightly linked product, because its greater length and two aldehyde groups allow it to bridge more distant pairs of protein molecules. It causes rapid and irreversible changes, is well suited to electron microscopy, works well at 4 °C, and gives the best overall cytoplasmic and nuclear detail. It is not ideal for immunohistochemistry.
Some protocols combine formaldehyde and glutaraldehyde so that their strengths complement one another.
These cross-linking fixatives, especially formaldehyde, tend to preserve the secondary structure of proteins and may also preserve most tertiary structure.
Precipitating, or denaturing, fixatives act by reducing the solubility of protein molecules and often by disrupting the hydrophobic interactions that give many proteins their tertiary structure. Precipitation and aggregation of proteins is a very different process from the cross-linking produced by aldehydes.
The most common precipitating fixatives are ethanol and methanol, which are commonly used for frozen sections and smears. Acetone is also used, and in the acetone-methyl benzoate-xylene (AMEX) technique it has produced better histological preservation than frozen sections.
Protein-denaturing methanol, ethanol and acetone are rarely used alone to fix blocks unless nucleic acids are being studied.
Acetic acid is a denaturant sometimes combined with other precipitating fixatives, as in Davidson's AFA.[13] Alcohols alone cause considerable shrinkage and hardening of tissue during fixation, while acetic acid alone is associated with tissue swelling, and combining the two may preserve morphology better.
Oxidising fixatives react with the side chains of proteins and other biomolecules and form cross-links that stabilise tissue structure. They cause extensive denaturation despite preserving fine cell structure, and they are used mainly as secondary fixatives.
Osmium tetroxide is often used as a secondary fixative for electron microscopy. It is not used for light microscopy because it penetrates thick tissue poorly.
Potassium dichromate, chromic acid and potassium permanganate are used in certain specific histological preparations.
Mercurials such as B-5 and Zenker's fixative have an unknown mechanism that increases staining brightness and gives excellent nuclear detail. They are fast but penetrate poorly and produce tissue shrinkage. Their best application is fixation of haematopoietic and reticuloendothelial tissues. Because they contain mercury, disposal requires care.
Picrates penetrate tissue well and react with histones and basic proteins to form crystalline picrates with amino acids, precipitating all proteins. They are good fixatives for connective tissue, preserve glycogen well, and extract lipids, giving better results than formaldehyde in immunostaining of biogenic and polypeptide hormones. They cause loss of basophils unless the specimen is washed thoroughly after fixation.
Hepes-glutamic acid buffer-mediated organic solvent protection effect (HOPE) gives formalin-like morphology, excellent preservation of protein antigens for immunohistochemistry and enzyme histochemistry, good RNA and DNA yields, and no protein cross-linking.
Fixation became a defined laboratory step in the nineteenth century, when microtomes and improved microscopes made thin sections routine and when pathologists needed to preserve tissue before embedding.
Formalin was introduced as a fixative in the 1890s, and neutral buffered formalin became the standard because it limited the formation of formalin pigment and preserved both morphology and antigens reasonably well.
Glutaraldehyde was developed as a fixative for electron microscopy in the 1960s, when its two aldehyde groups were recognised as giving stronger cross-linking than formaldehyde.
HOPE and other non-cross-linking fixatives were developed in the late twentieth and early twenty-first centuries to improve nucleic acid and antigen preservation.
Standardisation bodies and pathology societies have since published fixation guidelines, because pre-analytical variation remains a major source of diagnostic error.
A persistent controversy concerns how much fixation is enough, because under-fixation produces autolysis and poor sections while over-fixation masks antigens and impairs immunohistochemistry.
Antigen retrieval, using heat or enzymes, was developed to reverse some formaldehyde masking, but it does not work equally well for all epitopes.
The mesosome artefact showed that fixation can create structures that do not exist in living cells, and it remains a cautionary example in ultrastructure research.
Fixative choice is often a compromise between morphology, antigen preservation, nucleic acid integrity and safety, and no single protocol is optimal for every downstream assay.
Formaldehyde is a carcinogen and a workplace hazard, which has driven interest in safer alternatives and in closed automated processing systems.
Fixation quality directly affects diagnosis. Under-fixation can produce nuclear clearing, cytoplasmic vacuolation and tissue breakdown that mimic necrosis or autolysis, while over-fixation hardens tissue, causes tearing and curling during sectioning, and weakens immunohistochemical signal.
In tumour pathology, delayed or incomplete fixation can affect grading, margin assessment and the detection of lymphovascular invasion. For specimens destined for molecular testing, prolonged formalin fixation also reduces the yield and integrity of DNA and RNA.
In transplant pathology, fixation influences the recognition of ischaemia-reperfusion injury, rejection and infection. Biopsy specimens are usually small, so fixation time is easier to control, but drying must still be avoided.
In autopsy practice, the degree of autolysis depends on the interval between death and fixation, ambient temperature and tissue type, so the fixation protocol must be adjusted rather than copied from surgical biopsy practice.
Operational quality assurance includes recording the time of removal, the start and end of fixation, the fixative volume, the pH and the lot number. Specimens should be sliced or trimmed so that fixative reaches the centre within a reasonable time.
Special tissues require specific handling. Bone and calcified tissue usually need decalcification before sectioning, and the choice of acid or EDTA affects antigen and nucleic acid preservation. Adipose tissue remains difficult to section because lipid is dissolved during dehydration, and lymph nodes need slicing or needle puncture because the capsule impedes penetration.
Frozen sections are fixed after cutting, so the fixative must penetrate quickly without destroying antigens. Alcohols and acetone are common, but they shrink cells, and timing must be adjusted to avoid drying or antigen diffusion.
Storage and transport also matter. Fixed specimens should be kept in sealed containers labelled with the fixative and the start time, and they should not be allowed to dry, freeze or shake excessively, because ice crystals destroy cells.
Formaldehyde is a carcinogen and a respiratory irritant, and glutaraldehyde is a sensitizer that can cause occupational asthma and dermatitis. Osmium tetroxide and mercurials require special waste handling, and picric acid must be kept moist because dry picrate can explode.
In teaching, well-fixed specimens show structure and disease clearly, whereas poorly fixed specimens teach students to mistake autolysis for necrosis. Fixation is therefore both a technical and an educational responsibility.
No fixative is ideal for every purpose, and the choice is always a compromise among morphology, antigen accessibility, nucleic acid integrity, cost, safety and turnaround time. Understanding these competing goals is the basis of rational fixation practice.
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