The study of histology is to understand the microanatomy of cells, tissues, and organs and also to learn about their functions in structural terms.
Without question, the most important step to viewing biological tissue is fixation. The purpose of fixation preserves the structure of the tissue permanently in as life-like a state as possible.
Fixation should be carried out as soon as possible after the removal of the tissues or (if possible) in-vivo or soon after death (in the case of autopsy). A variety of fixatives is available for use depending on the type of tissue present and the features to be demonstrated. Check literature for optimum fixation methods for your specimen.
A. Aldehydes: Aldehydes fix samples by cross-linking proteins, especially between lysine residues.
1. Formalin is the most common. The standard solution is 10% neutral-buffered formalin. It does not harm the structure of the protein greatly, usually antigenicity is not lost.
2. Paraformaldehyde is a purer version of formalin and is often used in its place as a superior fixative. Penetration is faster than formalin or glutaraldehyde. It must be made from a powder no more than 1 week prior to use.
3. Glutaraldehyde causes deformation of alpha-helix structure, so it is usually not a good choice for immunocytochemistry. However, the fixation is non-reversible and it gives the best overall cytoplasmic and nuclear detail. 2.5% buffered glutaraldehyde is a common fixative for electron microscopy. Tissue should be less than 1mm cubed and the fixative not more than 3 months old.
B. Mercurials fix by an unknown mechanism. They contain mercuric chloride. Penetration is relatively poor and causes tissue hardness. They give excellent nuclear detail and are best used with hemapoietic and reticuloendothelial tissue. Since they contain mercury, they must be disposed of carefully.
1. B-5 fixative mercury pigment and acid formaldehyde hematein are 2 artifact pigments that must be removed before staining.
2. Zenker's fixative is recommended for lymph nodes, spleen, thymus, and bone marrow. It fixes nuclei very well and gives good detail. The mercury must be removed prior to staining or black deposits will result.
C. Alcohols are protein denaturants and are not routinely used for tissues, as they cause hardness and brittleness. Sprays of alcohol can be used to fix blood smears or PAP smears by physicians, but cheap hair sprays do just as well.
D. Oxidizing Agents cross link proteins, but cause extensive denaturation.
1. Potassium permanganate
2. Potassium dichromate
3. Osmium tetroxide fixes fats and lipids well and imparts a black coloration to them.
E. Picrates: Unknown method of action. Produces good nuclear detail, but now as much hardness as mercurials.
1. Picric acid is explosive in it's dry form. When in solution it stains everything yellow.
2. Bouin's solution contains picric acid. It is recommended for testis, connective tissue, GI tract, and endocrine tissue.
FACTORS AFFECTING FIXATION
A. Buffering solutions are used to regulate the pH of the fixative. The best fixation is usually carried out at a neutral pH (6-8). Lower pH can cause formalin-heme pigment that will appear as black as polarizable deposits in tissue.
B. Penetration depends on the diffuse ability of the fixative. The best penetrators are formalin and alcohol. Glutaraldehyde is the worst. No fixative will penetrate more than 2-3 mm of solid tissue or 0.5 cm of porous tissue in a 24 hour period.
C. The volume of the fixative is ideally at a 20:1 ratio of fixative to tissue. Agitation and frequent changes of fixative should help to insure good fixation.
D. An increase in temperature will increase the speed of the fixation. Be careful not to "cook" the tissue. Heat will coagulate proteins quickly, but also speeds autolysis.
E. The concentration should be adjusted to the lowest level possible. A high concentration is costly and will introduce artifacts similar to excessive heating. F. The time interval between tissue removal and fixation is critical. Longer periods of time will introduce drying artifacts, lost cellular organelles, nuclear shrinkage, and artifactual clumping. Most tissues should remain in fixative for 24 hours, then stored in 70% ETOH.
A. Quick freezing: It is most important that the tissue be frozen quickly, preserving enzyme activity and antigenicity as close to in-vivo as possible. Freeze artifacts may be introduced during the freezing process if the sample is frozen too slowly. The ice crystal formation size and amount is directly proportional to the speed of freezing. These artifacts can be seen as holes when thawed and are quite evident microscopically.
1. Cryogens: There are many methods of freezing sample for cryosectioning. Often the tissues are frozed in the cryostat itself. A precooled isopentane bath is also used routinely. My preference for freezing tissue (surrounded by a commercial embedding compound) is placed directly into liquid nitrogen. LN2 is about -195 degrees Celsius and will cause a gas bubble to form if tissue is placed directly into it. This gas layer will impede freezing causing artifacts if not surrounded by media.
2. Cryoprotectants: To aid in the assurance that ice crystals do not form, a cryoprotectant is sometimes employed. Some common ones are 25-30% sucrose, glycerol and PVP.
B. Cutting frozen sections is not difficult, but the following recommendations make for a more satisfactory product.
1. Keep the microtome clean and lubricated.
2. Knife must be sharp, clean and nick-free.
3. Knife and chuck must be secure and tight.
4. Clean your knife between sections.
5. Temperature must be appropriate for the type of tissues you are cutting.
The anti-roll plate is a swing away plastic plate that rests against the knife. It must properly be adjusted to function correctly. Most need to be parallel to the knife edge. The temperature of everything is important in the cryostat. Touching the knife, the block, or the anti-roll plate will warm them. If the anti-roll plate is not touching the knife it will warm slightly and cause the sections to stick.
Tissue Processing: Once the tissue is fixed it must be surrounded and infiltrated with a matrix to increase stability for sectioning.
Dehydration: Tissues in aqueous solutions cannot usually be infiltrated with most medias. Water removal must be achieved gradually through a series of graded dehydrant (alcohol or acetone). Typically 50% 70% 95% 100% solutions are used.
At the microscopic level, forces between the liquid/gas interface will cause enough pressure to distort a biological sample, therefore it is not a good idea to let samples dry out during processing.
Clearing: After dehydration samples can be placed in a nonaqueous liquid miscible with the embedment media.
1. Xylenes are the most common clearing agent for paraffin embedment.
2. Toludine is more tolerant of residual water, but is 3x more expensive.
3. Chloroform is slow and a health hazard.
4. Methyl salicylate is expensive, but smells nice (also known as oil of wintergreen).
5. Some newer xylene substitutes are available and a must if sectioning beta-galactosidase (beta-gal) stained tissue. Most of these are limolene based. Limolene is a volatile oil found in citrus peels (smells wonderful). They are less of a helath hazard and I have found them to be quite good, as long as the tissues are well-fixed.
Embedment media: In order to cut biological tissue very thinly (to send photons or electron through), it must be embedded in a hard substance for support. The thinner the sample needs to be, the harder the embedment media must be. Infiltration of the media in it's liquid form is crucial to good sectioning. A vacuum can sometimes be applied for difficult tissues.
1. Paraffin is the most common media and can be purchased for use at different melting points.
2. Plastics are also available for thinner sections or special staining.
After embedment, samples are sliced to the desired thickness on a microtome. For light level viewing, the thickness is usually 1-10 microns. The most important feature of the microtome is a very sharp knife.
A. Knives can be made of different materials, but usually you are limited by the availability of adaptors to your own microtome and choice of embedment media.
1. The standard thick metal variety has been around for a long time. These are very sturdy and can be resharpened. They are wedge shaped and cost around $200.
2. Thin and disposable knives are nearly as good, you are assured a good edge every time and there is less worry when a nick in the knife occurs. They cost around $1 each and are easily disposed of. .
3. Glass knives can be used for plastic embedded tissue to cut thinner sections. The glass needs to be cut on a special machine, but cost is less than $0.10 per knife. They are the sharpest knives you can buy, but very brittle and not very durable.
B. Cutting sections is not difficult. Cutting good sections is a little more tricky. It is important to have properly fixed and embedded tissues or artifacts can be introduced into your sections.
Some common artifacts are: tearing or ripping of the tissue, chatter, holes, foling, and compression.
Paraffin Microtomy: After a ribbon of sections has been created, the ones that you want to mount on a slide should be laid out on a warm water bath (45-55 degrees Celsius) and picked up onto a slide. The slides should be warmed flat on a warming tray until all moisture is gone between the slide and the section.
For subsequent staining, it is a good idea to use high-quality slides. In the past, adherents like "subbing" the slides were used. Now some slides are made with ionically charged surfaces. They are streak-free and stress-free. Most tissues will stick until you wipe them off.
Plastic Microtomy: This is usually done for thinner sections and higher resolution. Glass knives are usually made just before use. A water trough is sometimes attached to catch the section. Epoxy resins can then be laid out on a drop of water and placed in a warm xylene environment to dry. Methacrolates are water soluble and will usually stretch on the water without help. These techniques require a bit more skill at the microtome.
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