Skip to main content

10% Formal Calcium (4% Formaldehyde Calcium)

10% Formal Calcium

(4% Formaldehyde Calcium)


10% Formal calcium – Baker

Strong formalin100mL
Tap water900mL
Calcium chloride20g

10% Formal calcium – Lillie

Strong formalin100mL
Tap water900mL
Calcium acetate20g


The only difference between the two formulae above is in the calcium salt used. Calcium acetate gives a more alkaline solution than calcium chloride. Apart from that they are much the same. Baker originally suggested adding calcium chloride to 10% formalin as a means of improving the preservation of phospholipids, but both the variants are suitable for routine fixation as well. The only concern would be in those cases which involve microscopic calcium deposits, but even then it is unlikely to be a serious concern. If this solution is used routinely, this aspect should be checked practically.

It is recommended that the formalin used be neutralized with marble chips or crushed oyster shells before the calcium salt is added. This may be done by diluting the formalin with tap water, adding the calcium carbonate, and leaving overnight. The next day the diluted, and now neutralized, formalin may be placed into a container and the calcium salt added.

Although this is a simple fixative, it is not that popular for routine use. Erythrocytes will not usually be damaged, but formalin pigment may still be produced. Although the diluted formalin has been neutralized with calcium carbonate, formic acid will continue to be produced and may cause formalin pigment, especially in bloody tissues, although calcium acetate in Lillie’s variant gives some protection from this.

Bancroft and Stevens note that 7 grams of cadmium chloride per liter may be added to the Baker formula, without specifying the purpose.


This fixative should be applied overnight as a minimum, but fixation is not complete until applied for a few days, and a week or two is not too long. For thorough fixation, the proteins in the tissue need to be crosslinked, but it is well known that simple formalin mixtures discolor tissues well before they crosslink the proteins. For that reason, visual observation does not give any indication as to the degree of fixation, and discoloration should never be used as an indicator that it is complete.

It is essential that the time in fixative be noted, and sufficient time be allowed for the chemical reactions to occur. Time measured in a few hours is not adequate, and the lack of crosslinking in tissues treated for such a short time will not give adequate protection to the tissue from the fixation effects of dehydrating ethanols. Smaller pieces of tissue do not fix appreciably faster than larger pieces. Fine needle biopsies require the same length of time as 3 mm thick slices of solid organs.

On a practical basis, in a diagnostic laboratory, the slowness of simple formalin fixation is a distinct drawback. If the constraints of making a diagnosis are a major problem, then consideration should be given to increasing the temperature of the fixative solution. It must be emphasized that, although increasing the temperature of the fixative may increase the speed of fixation, it will cause a reduction in the quality of the final stained section and there may also be effects on some staining or immunohistochemical reactions. The temperature increase should be kept to a minimum consistent with accomplishing the goal of faster fixation, since heat itself is a means of fixation. If increased temperatures are used, then formaldehyde fumes will be generated, more so as the temperature is increased, so fixation should be done in a fume chamber to protect personnel.

Complete organs are often placed in containers filled with a simple formalin variant. Although formalin penetrates fairly rapidly, it will not reach the center of a large mass for some time and the delay may permit some autolysis in the middle of the tissue. Large organs should be described as soon as possible, then sliced into 1 cm or so slices to allow the fixative to gain access. Similarly, intestines should be opened and cleaned of any contents, then returned to a container large enough to permit the fixative to contact all the tissue surfaces, or pinned out on a board with rust-free pins and placed into the fixative.

Tissues may be stored in this solution for long periods, but the fixative should be changed at least every six months.


There is no special aftertreatment.

Secondary fixation

Other fixatives may be applied after formalin fixation, and some of their characteristics will be obtained. It must be realized that secondary fixation in any fixative will not give the same results as would have been obtained if the secondary fixative had been applied to fresh tissue.

Safety Note

Prior to handling any chemical, consult the Safety Data Sheet (SDS) for proper handling and safety precautions.


  1. Culling, C.F.A.,
    Handbook of Histopathological and Histochemical Techniques, Ed. 3,
    Butterworths, London,, UK.
  2. Bancroft, J.D. and Stevens, A.,
    Theory and Practice of Histological Techniques, Ed. 2,
    Churchill Livingstone, Edinburgh,, UK.