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Tissue Preservation

in Histology

Preservatives are separate and distinct from fixatives, even though they may contain similar chemicals. The primary purpose of a fixative is to chemically alter the tissue proteins so that microscope preparations may be made. The primary purpose of a preservative is to keep, i.e. preserve, the tissue in the same state it was when first immersed in the preservative, that is without any deterioration. Some are used after the tissue has been fixed for long term storage or display, others with unfixed tissue to ensure viability of individual tissue components, still others as a coverslipping medium.

Preservatives used for coverslipping differ from mounting media because they do not solidify and require that the medium be sealed against evaporation and leakage by hardening media such as asphalt varnishes, paraffin wax or nail polish. Hardening mounting media are listed separately.

It should be noted that the method which preserves tissues the most effectively is to process them through paraffin. They will then be stable for very long periods of time, well over 100 years, as shown by the preservation in blocks processed at the end of the 19th century when histotechnology was in its infancy. Those blocks can still be sectioned and stained, the results being indistinguishable from those of well processed tissues from the present.

Formalin-Based Preservatives

10% Formalin

Simple formalin solutions, the same as are used as fixatives, are convenient and effective as preservatives if a few precautions are taken. Chief among these is to change the formalin solution regularly every six months or so. This will allow the tissue to be stored for several years and, as a bonus, it ensures complete and thorough fixation.

Long term storage in simple formalin usually results in formalin pigment and, should the tissue develop this, it will need to be removed before staining. The formation of formalin pigment can be overcome to a large degree by using neutral buffered formalin as the storage fluid, since formalin pigment is produced more rapidly at acid pH, but it is essential that the NBF be regularly changed as formalin pigment may still be formed on long term storage if it is not.

Long term storage in formalin may cause some relatively minor changes to the staining. Basophilia may be reduced, and eosin may not be as avid. These can usually be overcome by increased staining times. These effects may be aggravated by failure to change the formalin regularly.

Although 10% formalin variants are convenient for storage, it is quite feasible to use much lower concentrations of 1% or even less. Once the tissue has been properly fixed by being in a 10% variant for some time, the preservative does not need to be so strong that fixation continues, merely strong enough to inhibit bacterial action.

Kaiserling Fluids

There are four preservative formula under Kaiserling’s name. Two of these (Kaiserling I and II) are used sequentially for the preparation of medical museum displays. The other two are general preservative solutions, although all four may be used for that purpose.

It is not an uncommon practice to substitute 10% formalin for Kaiserling I as it does much the same job, although tissue must be left in it for fixation to be complete – a few months at least. One effect of using either 10% formalin or Kaiserling I is that the natural colour is lost during fixation. This may be restored in two ways:

  • Soak the fixed tissue in 80% ethanol until the colour returns, then immerse in Kaiserling II in a sealed jar.
  • Immerse in Kaiserling II in a sealed jar after adding a small amount of potassium hydrosulfite. The colour slowly returns over a few days.

For those interested in this procedure please consult a text on medical museum technology.

Kaiserling I

Strong formalin200mL
Potassium nitrate15g
Potassium acetate30g

Kaiserling II

Potassium acetate100g

Kaiserling 1896

Strong formalin430mL
Potassium nitrate5g
Potassium acetate17g

Kaiserling 1899

Strong formalin160mL
Potassium nitrate9g
Potassium acetate18g

Alcohol-Based Preservatives

Spirits of Wine

The term spirits of wine refers to strong brandy, i.e. strong ethanol of about 80% or more, as it is produced from a distillery. The actual ethanol concentration depends on how many times it has been distilled, but 80% ethanol is not unusual. During the 1800s and earlier this was readily available. It was one of the first means to preserve gross specimens and to fix tissues for microscopic examination. Today, of course, the use of brandy would be prohibitively expensive. It would be more reasonable to use 95% industrial ethanol or methylated spirits. Due to the distortion that ethanol causes during long term storage, there is no reason to use it.

70% Ethanol

One of the commonest fluids recommended for long term storage is 70% ethanol, although it makes little difference whether it is made from pure ethanol or methylated spirits. Tissues can remain in this fluid for very long times. The fluid should be changed from time to time. Ethanol is a solvent and will extract some materials from the tissue, including some lipids. Using 70% concentration rather than 95% or absolute ethanol minimizes this. Storage in ethanol initially restores the colour of formalin fixed tissues, but this is temporary and the colour eventually fades again. It may not then be restored a second time.

A quick means of making a known concentration of ethanol from a stronger concentration without having to do any calculations is to take the volume equivalent to the concentration wanted and add water to the volume equivalent to the stronger concentration used. For example, to make 70% ethanol from 95% ethanol, take 70 mL of 95% ethanol and add enough water to bring the volume to 95 mL. You will then have 95 mL of 70% ethanol. This works for any dilution and all that is required is a large enough graduated cylinder.


Glycerol (glycerine) is easily available at a reasonable price and has little chemical effect on tissues. It raises the refractive index somewhat, but that is not necessarily a defect as it may enable structures to be seen within the tissue – lymph nodes, for instance.

At its simplest glycerol can be used as a simple solution in water or as 100% glycerol. The tissues can be dehydrated with increasing concentrations of glycerol in a manner similar to dehydration with increasing concentrations of ethanol, stopping when the final concentration wanted is reached. The tissue can then be stored for extended periods. There is no real need to replace the glycerol, but it should be appreciated that concentrations less than 100% may preserve less well than absolute glycerol due to the presence of water. At higher concentrations of glycerol some lipids may be removed. Triglycerides are glycerol and fatty acid compounds, after all. If the glycerol becomes dirty or greasy, then the solution and container should be changed.

There are many formulations of preservatives containing glycerol and they may contain ethanol, formaldehyde, and acids. Many of these were designed for coverslipping rather than displaying specimens, which is better done using Kaiserling fluid, but since they do not harden the coverslip will need to be sealed if a degree of permanence is required.

The lists of formulas below is by no means exhaustive. There are dozens of such mixtures and those interested should read the text listed as the reference.

Formula NameMaterial (mL)
Water95% EthanolGlycerolStrong FormalinGlacial Acetic AcidConcentrated Nitric Acid
Gatenby & Painter502525


  1. Gray, Peter. (1954)
    The Microtomist’s Formulary and Guide.
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.
    Berliner Klinische Wochenschrift, (1896), v. 33, pp.775, Berlin.
    Virchows Archiv für pathologische Anatomie und Physiologie und für
    klinische Medizin
    , (1897), v. 147, pp.389. Berlin.
    Verhandlungen der Deutschen Pathologischen Gesselschaft, (1899), pp.203, Jena.