Tissue removal and fixation
Introduction
Unlike plant tissue, cutting animal tissue is much more difficult. Animal tissue is not so easy to cut. The material is too soft and deforms easily. Moreover, coupes (slices) may not exceed 10 μm (1 μm = 1/1000 mm), but preferably even less, thick if they are to be usable for study. Like plant tissue has cell walls, animal tissue has no cell wall, therefore it is not rich in contrast. Therefore, staining of the specimen will always be necessary. In order to stop the autolysis (degradation) processes, animal tissue will always need to be fixed. The structure of the tissue remains roughly the same, but chemically it changes a lot. The tissue becomes a little harder, but cutting is still not possible. The tissue is stiff. Moreover, these are often small pieces of material that are almost impossible to hold. A good solution for cutting is the use of paraffin.
This substance can be processed very well but has a big disadvantage: it is not soluble in water and almost all animal tissues consist for a large part of water. For this reason, in several steps (50% ethanol, 70% ethanol, 85% ethanol, 96% ethanol and 100% isopropanol) the water is displaced by alcohol (dehydration). After all the water has been displaced by alcohol, the alcohol is in turn displaced by an intermedium. Examples of intermediates are: xylol, butanol, benzol, toluol et cetera. These substances are soluble in paraffin. After the intermedium has been displaced by paraffin, the tissue can be moulded into a block of paraffin. The moulded tissue can now be cut into thin sections on a microtome. The wafer-thin sections are brought from a hot water bath to a microscope slide or sometimes glued with a protein/glycerol mixture.
Histology
What is histology? Histology literally means: studies the microscopic anatomy of biological tissues. In order to be able to study animal tissues such as lung, heart or kidneys at cellular level with a light microscope, it is necessary to first fix the tissue (stop chemical processes), cut very thin slices of it, attach them to a solid surface, stain them in special ways and also make them "preservable". A so-called preservable section is made. For this purpose, a sample is made, which can be studied later, is called histological technique. This whole process is always done in a fixed pattern.
1. Tissue removal
2. Fixation
3. Dehydrating
4. Embedding in paraffin
5. Cutting sections on a microtome
6. Mounting sections to an object glass
7. Hydrating
8. Staining
9. Make the sample preservable.
1. Tissue removal
Tissue has to be 'fresh'. After about 1 hour autolysis (degration) starts and it is less or no longer usable for histological purposes. In order to obtain usable tissue, one can use freshly caught fish, a visit to a slaughterhouse or, for example, breeding mice yourself. Mice will be frequently used on this website. The reason for this is that mice are easy to obtain, the tissue is often similar to human tissue and because mice are widely used in science, a lot of information can be found on the internet in both Histology and Pathology. When the goal is to prepare newborn mice it is advisable to separate the pregnant mouse from the other mice. Immediately after birth, the male will bite the young mice to death.
It does not make sense to completely fix a dead animal. The histological techniques described here require pieces of tissue (muscle, skin, heart, liver, kidney, brain, etc.) from a few mm to about two cm maximum. Larger pieces cannot be fixed well (not fast enough) and will therefore partially degrade. An exception to this is the fixation of a complete embryo of a small animal such as a rat or mouse, and even a newborn mouse can be completely fixed with a special technique. Further on in this histological chapter this will be discussed and in the slides pages there are several examples available that show the fixation of a newborn mouse. If a slice is cut out of an organ, make a sketch of the location and direction of the slice. What will eventually be visible in a sample depends very much on that. Make a work table and prepare all materials and tools.
Killing small animals, like a mouse here, is easy by administering an ether-soaked pellet of cotton wool in a sealed container. The killed mouse is briefly immersed in 50% ethanol. This is to work more sterile and to prevent hairs from being taken in during the incision. The mouse is neatly clamped onto a piece of firm foam with needles. Wear plastic gloves throughout the entire period to prevent possible bacterial infections. First make a solution of 9 grams of table salt (sodium chloride) in 1 litre of water. This solution is called "physiological saline solution". It reasonably corresponds to the "salinity" of tissue fluid. Organs and tissue can be rinsed with it. Never use tap water for rinsing fresh examination material. Due to the large differences in osmotic value, the cells will burst open. Use a scalpel, tweezers and bandage scissors. Pick up the skin from the thoracic or abdominal cavity and make an incision. The muscle layer will then become visible. Do the same with it (in some animals, such as fish, this does not work. Then try to stab straight through to the abdominal cavity). At the chest cavity the ribs will have to be cut. This can be done in smaller animals with bandage scissors (e.g. mice). In larger animals a pair of tin or pruning scissors is desirable. At the thoracic cavity a kind of lid can be loosened by cutting left and right. In the abdominal cavity, the muscle layer can be pushed open or flaps can be cut by making a transverse opening. In the thoracic cavity the lungs and the heart can be found. The abdominal cavity usually contains the liver, stomach, kidneys and intestines.
Use tweezers to grab an organ to a blood vessel or other offshoot (the lung e.g. near a trachea) and cut it free. If desired, always rinse with physiological saline solution in order not to lose sight through blood. A blood smear can of course be made immediately from the fresh blood. Use a new scalpel knife to cut out the desired pieces from the organs. Do not do this with scissors, because the tissue will be squeezed to pieces. Immediately transfer the pieces of organ into a fixative to prevent further degradation. It is important to have several small jars of fixative ready. Do not put two types of organs in one jar because after fixation the difference is hardly visible anymore because almost all tissues take on a grey-brown color. Label everything with tissue type and date group. Do not forget to thoroughly clean the work table and tools and disinfect them with a disinfectant (e.g. ethanol).
2. Fixation
Why fixation? Fixation stops the complicated metabolic processes and prevents further tissue degradation. The purpose of fixation is to secure in as identical way as possible the structure that the tissue had when it was still alive. Tissue freezing is not an option because living tissue consists of about 80% water and therefore all cells would burst (water expands when frozen) and by the formation of ice crystals the sharp crystals cut right through the structure of the cell. Special techniques can be used to make freezing sections, but this will not be discussed here. A chemical intervention is therefore necessary. In order to give the fixative the chance to work properly and as quickly as possible, it is important that the tissue pieces are not too large.
Blocks of 4 to 8 mm are easy to fixate or a slice of a few millimetres thick and may be 2 or 3 centimetres long. It is also better to hang the tissue in the fixating fluid instead of leaving it on the bottom. This can be achieved by using cassettes or placing a tuft of cotton wool at the bottom of the jar on which the tissue will lie. In any case, the aim is to allow the fixer fluid to penetrate the tissue from all sides as quickly as possible. One way to increase this speed is to pre-fix tissue (especially the larger pieces) warm between 35-40°C and then cold-fix it further[1]. A major disadvantage of fixation is that the tissue shrinks or swells. Experience shows that the more water a tissue contains, the greater the volume change will be. The shrinkage can be so significant that the epithelial layer of the underlying connective tissue tears. Subsequent treatments such as dehydration also usually lead to tissue shrinkage. In total, the shrinkage can be as much as 20 to 25% of the original size. There are many types of fixation fluids that all have their own advantages and disadvantages.
- Ethyl alcohol or
Ethanol[1] is a fixative that acts quickly but abruptly drives the water out of
the tissue. It is suitable for small pieces of tissue (3-4 mm thickness is
fixed in 2-4 hours). The shrinkage is large and the outside of the tissue
hardens quickly and is negative for later cutting. The composition of proteins
is not affected by ethanol fixation. Substances enclosed in proteins such as
mucus and glycogen remain intact. Lipids (fat) and hemoglobin (red dye of
erythrocyte) are rinsed out. Ethanol is sometimes used in a blood smear but
mainly in botany.
- Formaldehyde[1] is
a strong smelling gas that dissolves well in water. The solution in water is
called formalin and contains 37-40% formaldehyde (syn. formol) (it was formerly
called 'strong water'). The solution penetrates the tissue well and is also
suitable for fixing larger pieces. In contrast to all other fixatives, a piece
of tissue in formaldehyde can stay for months without clearly changing the
structure. If we want to preserve tissue longer, it is advisable to add a small
chunk of marble to the formalin. The reason for this is that under the
influence of sunlight and substances pulling out of the tissue, formic acid is
formed spontaneously. A concentration of 4% formaldehyde (formol) in water is
usually used as a fixative. Look in the tab 'solutions' for a handy table.
- Zenker[1] is a
combination fixative that consists of: Sublimate (mercury 2 chloride),
potassium dichromate, sodium sulfate, glacial acetic acid and distilled water.
Look at 'making solutions' for the correct mixing ratios. Single sublimate
causes the cytoplasm to shrink sharply but in combination with formalin,
glacial acetic acid, trichloroacetic acid or chromium salts it is one of the
most widely used fixatives. A disadvantage of this fixative is that there is
always a precipitate of mercury that has to be removed from the preparation.
After fixation in Zenker, the preparation is easily accessible for many dyes
and even necessary for some stainings, e.g. Mallory or AZAN.
- Bouin[1] is a
combination fixative consisting of: Picric acid, formol and glacial acetic
acid. This fixative is widely used and is excellent for overview staining or
cytological studies and embryos. Fats are not preserved. The shrinkage is only
2.5%. The fixation time is between 2 and 24 hours depending on the size of the
tissue. A major advantage of this fixative is that the tissue can stay for
months without damaging the tissue. Most specimens on this website are
therefore fixed with Bouin. It can be purchased ready to use but is also easy
to make yourself. For correct mixing ratio view the tab 'making solutions'.
- Stieve[1] is a
combination fixative that consists of: Sublimate (mercury 2 chloride), formol
and glacial acid. It penetrates the tissue very quickly and is therefore also suitable
for fixing larger pieces of tissue. After fixation, the tissue is placed
directly in 96% ethanol without the usual lower ethanol range. Trichloroic
acetic acid can also be used instead of glacial acetic acid.
- Susa according to
Heidenhain[1] is a combination fixative that can be made from two shelf-life
stock solutions. It consists of: Sublimate (mercury 2 chloride), formol,
glacial acetic acid, salt, trichloroic acetic acid and water. The mixture
penetrates the tissue very well. Fixation time between 1 and 24 hours. After
fixation, the tissue is removed and fixated in 90% ethanol which must be
refreshed several times. A disadvantage of this fixative is that there is
always a precipitate of mercury which has to be removed from the sample.
The duration of fixation strongly depends on: the
fixative used, the type of tissue and the temperature of the fixative. Most
preparations described on this website are fixated with formaldehyde, Zenker or
Bouin. The duration varies from a few days to about a week. The fixation times
used are mentioned in all preparations and partly empirically determined. For example, a piece of spongy lung tissue
will need a shorter fixation time than the much firmer liver tissue. If, for
example, you want to fixate a newborn mouse as a whole, it is advisable to cut
the animal at the sides. A few small incisions through the skin is sufficient.
The extremities can also be removed so that the fixative fluid can penetrate
there as well.
References:
[1] Prof. Dr. Peter Böck (1989, 17., neubearbeitete auflage), Romeis Mikroskopische Technik, München. Verleger Urban & Schwarzenberg. Hoofdstuk 4, 'Fixierung histologischer Präparate'.