regarding the fixation and mounting of museum specimens

1. By : Dr SHASHIDHARA T S
Moderator : Dr ASHALATHA

2. INTRODUCTION
 Pathological museums are in part historical,
representing the pioneer work of diagnosticians and
therapists.
 Presenting records of past states not now encountered,
or conditions of great rarity; and finally they provide
the student with the basic material of his/her current
teaching.

3. BASIC MUSEUM TECHNIQUES
1. RECEPTION
2. PREPARATION
3. FIXATION
4. COLOR RESTORATION
5. PRESERVATION
6. MOUNTING
7. SPECIAL METHODS
8. PRESENTATION

4. RECEPTION
SOURCE :Most of the material is collected from
 Teaching hospitals which could be surgical resected
specimens –operating theatres
 Necropsy specimens- PM room
 Research laboratories
Specimens should be received with full details of the
patient/ lesion.

5. PREPARATION OF THE SPECIMEN
 One of the commonest causes of inferior quality
specimens is contact with tap water. The resultant
haemolys is greatly reduces their value.
 Specimens should be washed only with saline, and
should be kept in saline while awaiting demonstration
as drying again ruins the surface appearances.
 But as autolysis quickly sets in they should not remain
in saline for more than two hours.

7. FIXATION OF THE SPECIMEN
 The objective of fixation is to preserve cells and tissue
constituents in as close to life-like state as possible.
 Fixation arrests autolysis and bacterial decomposition
and stabilizes the cellular and tissue constituents.
 The fixatives used in museums all over the world are
based on formalin fixative technique, and are derived
from Kaiserling technique and his modifications.

8.  Kaiserling recommended that the initial fixation
should be in neutral formalin (KI) solution and then
transferred to a final preserving glycerin solution
(KIII) for long term display.
 Color preservation is also maintained with these
solutions.

9. PRINCIPLES OF FIXATION
 Specimens containing bile or stained by bile must be
fixed and stored apart from others.
 Specimens undergoing fixation must not touch other
specimens, or the sides of jars; they must either lie on
washed fluffiless lint or should be suspended by linen
thread.

10. • Flat flaps of tissue like stomach, intestine etc. should
be fixed to cork board and left in formalin so that they
are not crumpled and irregularly fixed.
• Unopened cystic cavities should be injected with
fixative if opened they should be packed with cotton-
wool.
• Solid viscera should be fixed by vascular injection
example; brain through basilar artery.
 The lungs and limbs are particularly suitable for
fixation by vascular injection.

11. FIXATION TECHNIQUE
 The technique most widely used is modification of
method described by Kaiserling (1897)
 The original technique employed 3 solutions : first for
fixing, the second for restoring color, and the third a
mounting fluid.

12. Kaiserling No.I –Fixing fluid
Formalin (40%) - 400 ml
Potassium Nitrate - 30 gm
Potassium acetate - 60 gm
Water up to - 2000 ml
 Fixation in Kaiserling No.1 solution for 24 hrs to few
weeks depending on the size of specimen.

13. Kaiserling No.II Solution
 Specimen is placed in 80% Ethyl alcohol solution for
optimal period of 1 hour (up to 4hrs depending upon
the size of the specimen) if the specimen is discolored.
 If the specimen is left for too long in alcohol- the color
will fade and this effect is irreversible.
 NOTE :This step is not necessary when using a sodium
hydrogen sulphite mounting fluid.

14. COLOR RESTORATION
 The fixed specimen is now transferred to a jar
containing industrial methylated spirit until the color
is fully restored.
 The alcohol penetrates the tissues rapidly.
 If the specimen floats, it should be lightly covered with
surgical gauze, and the vessel closed to prevent
evaporation.
 Color restoration is usually complete in two to eight
hours, depending on the size and character of the
specimen.

15.  Pulvertaft (1936) described a method of restoring the
color to tissues by addition of reducing agent (sodium
hydrogen sulphite) to the mounting fluid.
 Specimens mounted by this technique showed
remarkably little fading even after 25yrs.

16. Original Kaiserling No. III solution
 Glycerine 500 ml.
 Arsenious acid 1% 200 ml.
 Pot. acetate 250 g.
 Thymol 2.5 g.

17. Pulvertaft – Kaiserling mounting fluid III
Glycerine - 300 ml
Sodium acetate 10% (pH 8) -100 g
10% Formalin - 5 ml .
Tap water - 1000 ml
 Camphor / Thymol can be added to prevent growth of
moulds.
 Immediately before sealing 0.4% sod. hydrosulphite is
added. The amount of hydrosulphite should not normally
exceed 0.4%.
 If color restoration must be rapid, 0.6% may be added, but
this is to be avoided, as a white precipitate may form.

18.  If the solution is not crystal clear, it is usually due to
impurities in the sodium acetate.
 Such solutions should be filtered through paper pulp
under negative pressure.
 If this fails, 50ml of saturated solution of camphor in
alcohol should be added to 1 liter of the solution,
refilter as before.

19.  Carbon monoxide has also been employed as a color-
retaining agent. Schultz(1931) introduced the
technique, which gives brilliant color contrast, but
entails the risks of poisoning and explosion and also
colors are unrealistic.

20.  Israel and Young (1978) used pure liquid paraffin as
the final mountant after color restoration with alcohol.
 This procedure reduces the discoloration of the
mounting fluid by pigments in the specimen.

21. HOLLOW VISCERA
 Cut hollow organs should be padded out with cotton wool,
but if uncut they can be pressure inflated. For example
 Through urethra into the bladder,
 Through urethra into Pelvicalyceal system,
 Through trachea into lung, and
 by direct injection in the case of cysts.
The fixative can be injected into such organs with a
Higginson syringe or with a conventional hypodermic
syringe.

23. HEART
 Specimens of heart -usually been cut before being
sent to the museum , in order to maintain the
natural shape it is important to pad out all cavities
and major vessels with cotton wool before fixation.
 A heart received fresh and uncut is placed in an
adequately large container of fixative and
additional fixative perfused through the coronary
ostia with a syringe, heart will revert to its natural
shape.
 This is the ideal method for fixing hearts for
display.

24. BRAIN
 Soft in consistency , difficult to handle in a fresh
state, so it is necessary to fix the brain before
cutting.
 Also because of the softness, if the specimen is
allowed to rest on the base of the container , even
if supported with cotton wool, distortion will still
occur.
 It is therefore preferable to perfuse the brain
through the basilar and cerebral arteries at its base
and it should then be suspended by the basilar
artery within the fixative -at least a week
 It can be easily bisected or sliced with a brain knife

25.  After fixation when the specimen is ready to be
mounted, pH should be determined.
 If the pH is greater than 6.5, specimen is placed
directly in solution III (mounting fluid).
 If pH is less than 6.5, specimen is placed in solution II.

26. Factors affecting fixation
There are a number of factors that will affect the
fixation process:
 Buffering
 Penetration
 Volume
 Temperature
 Concentration
 Time interval
 Position of tissue

27. Buffering
 Fixation is best carried out close to neutral pH, in
the range of 6-8.
 Hypoxia of tissues lowers the pH, so there must be
buffering capacity in the fixative to prevent
excessive acidity.
 Acidity favors formation of formalin-heme
pigment that appears as black, polarizable
deposits in tissue.
 Common buffers include phosphate, bicarbonate,
cacodylate, and veronal.
 Commercial formalin is buffered with phosphate
at a pH of 7.

28. Penetration
 Penetration depends upon the diffusability of each
individual fixative, which is a constant.
 Formalin and alcohol penetrate the best, and
glutaraldehyde the worst.
 Mercurials and others are somewhere in between.

29. Volume
 The minimal acceptable volume of fixation fluid is
about 15 to 20 times the volume of the specimen.
• The use of small volumes of fixation fluids for larger
specimens is the most frequent cause of poor tissue
preservation.

30. Temperature
 Increasing the temperature will increase the speed of
fixation.
 Hot formalin will fix tissues faster.

31. Concentration of fixative
 Concentration of fixative should be adjusted down to the
lowest level possible.
 Too high a concentration may adversely affect the tissues
and produce artifact similar to excessive heat.

32. Summary
 FIXATION :
 Confers chemical stability on the tissue
 Hardens the tissue (helps further handling)
 Halts enzyme autolysis
 Halts bacterial putrefaction

33. PRESERVATION
 The specimen together with a duplicate label, is
wrapped in gauze or muslin and the label attached
with a piece of linen thread.
 Specimens are preserved in large rectangular
earthenware tanks.
 The fluid used may be Kaiserling fixing fluid I for a
period of 6 months.
 After which time specimens should be treated with
80% alcohol to restore the color.

35. MOUNTING
 Specimens are trimmed to the desired size and
shape so that it fits into the jar. All unwanted and
non representative tissues removed by careful
dissection.
 If after removal of cotton wool packing from
cavities, the specimen will not remain in a natural
position by normal mounting methods, such
cavities should be filled with arsenious acid-
gelatin.
 Regular cuts are given keeping in anatomical
position.

36.  Specimens which are friable may be covered with a
thin layer of arsenious acid-gelatin(Wentworth 1947)
and it may also be used locally to hold fragments such
as blood clot in position.

37.  Bile stained specimens are soaked in saturated
solution of calcium chloride for 24hrs to avoid
discoloration of mounting fluid.
 This will only reduce the degree of coloring, and
frequent changes of fluid is necessary to keep the
discoloration at the minimum.

38. ROUTINE MOUNTING PROCEDURE
 Museum jars or boxes
 Centre plates
 Stitching specimens to centre plate
 Fixing the centre plate
 Filling and sealing

39. MUSEUM JARS OR BOXES
 Perspex boxes are used almost universally.
 They are available commercially or may be made in the
laboratory.
 The method employed commercially to join the sides
is far superior to the cementing process that is done in
the laboratory.

40.  The specimen should be laid on a flat, waterproof
bench.
 The position in which they are to be mounted should
be anatomically correct.
 Specimen is then measured, allowing a ½ inch
clearance at the bottom is to enable a label to be fitted
without obscuring part of the specimen.

41.  Depth of the specimen is measured and approximately
¼ inch is added for the centre plate.
 Perspex sheet can be moulded or bent to satisfy the
requirements of individual specimens.

42. Bending of the
corners was effected with the flame from a Pasteur pipette

43. Centre plates
 Advantage of the perspex sheet is its flexibility when
heated.
 Specimens can be stitched to a flat sheet of perspex.
 Commercial boxes may be available with already fitted
center plates.
 Colored opaque plates may be used to enhance the
color of the specimen or to attach specimens on both
sides.

45. Stitching specimens to centre plate
 The specimen is arranged in the desired position, and
crosses are made on the centre plate with a scribe
where stitches are to be placed.
 With solid specimens the number of stitches will
depend on the weight and consistency of the tissue: for
example, half a kidney is adequately supported with a
stitch at each pole.

46.  Hollow or cystic organs, or organs with attached
structures, may require stitches to hold the specimen
in the correct position in addition to providing
support: for example, the oesophagus and stomach
may require up to 12 stitches.

47.  Attached structures may need to be stitched to the
main organ or to each other to hold them in position.
 Stitches must not be placed through pathological
lesions.
 When the centre plate has been marked, holes 1/16
inch in diameter are drilled at those points.

48.  If linen thread is to be used, one hole is drilled at each
point; if nylon thread is used two holes are necessary.
 Nylon thread has the advantage of being almost
unbreakable but is so hard it tends to cut through
specimens and for this reason linen thread should be
used for all specimens except bone.

49.  Lengths of linen thread are cut and a small clear glass
bead is threaded on and tied in the centre; the bead
should be slightly larger than the hole in the centre
plate since it acts as a retainer for the tie.

50.  The centre plate is thoroughly washed in a detergent,
and dried on a fluff less cloth.
 The specimen is stitched on by passing first one end of
a tie and then the other through the centre plate and
the specimen, pulling on both ends until the glass
bead is tight against the centre plate.

52. Fixing the centre plate
 The centre plate, with specimen attached, is put into
the box and marks are made with a grease pencil if
stops are required to hold the centre plate in position.
 If the box is of the correct depth there will be no
movement of the specimen, but if a deeper box has
been used, two rectangles with polished edges, are
cemented to the wall of the box to keep the centre
plate in position.

53. Filling and sealing
 When the specimen is in position, museum fluid, to
which 0-4 per cent sodium hydrogen sulphite has been
added, is run in to within 1/2 inch of the top.
 Air-bubbles trapped between the specimen and centre
plate are released with a broad bladed spatula.

55.  A hole 1/8 inch diameter is drilled in one corner of the
lid.
 The top of the box is wiped dry and Perspex cement
applied with a Pasteur pipette.
 After 30seconds the lid is laid lightly in position.
 After a further 30 seconds, a lead weight is applied and
left for at least 1 hour, preferably 2-3 hours

56.  A short length of Perspex rod, 1 inch In diameter, is
tapped lightly into the hole in the lid and the
specimen left for 24-48 hours to remove residual air-
bubbles.
 When the last bubble is removed, the Perspex plug is
placed and tapped firmly into position and, when
dried, a small amount of Perspex cement is applied.
 NOTE: It will be found that large specimens develop
air-bubbles over a period of 2-3 weeks after mounting,
these may be removed by drilling a fresh hole, filling
up and replugging.

57. MOUNTING IN GLASS JARS
 The specimens are mounted as described above except
that holes are drilled with an engravers tool (a metal
rod with a diamond-shaped end) in a hand drill, using
camphor dissolved in turpentine as a lubricant.
 Glass jars are sealed with an asphaltum-rubber
compound(Picein).
 The jars look neater if, after sealing the edges are
painted black enamel or asphaltum varnish.
 NOTE:If the glass jars are completely filled with
mounting fluid, they will crack with atmospheric
changes.

58. GELATIN EMBEDDING
 Delicate structures(for example the circle of Willis)
which are difficult to stitch, may be embedded in a
thin layer of arsenious acid-gelatin on a centre plate,
and then mounted by the routine method.
 A trough is formed by applying sellotape around the
edge of the centre plate and filled with gelatin. The
sellotape is removed after the gelatin has set.

60.  This method was popular with glass containers since
the gelatin protected the specimen and avoided the
sudden release of a glycerin solution.
 It has the disadvantage that the gelatin tends to
become yellow with age, and also to undergo
liquefaction with the resultant formation of air-
bubbles.

61. EMBEDDING IN SOLID PLASTIC
BLOCKS
 Embedding in a solid block of plastic would appear to
offer the ideal method of presenting museum
specimens, but unfortunately till now no method is
available which preserves the color of soft tissues.
 Such methods while adequate for hard tissues (certain
insects, plants, and so on) are useless for the normal
pathological museum specimen.

63. MACERATION
 Maceration is used to demonstrate bony
lesions such as osteogenic sarcomas , osteomas
and tuberculosis.
 This method enables preservation of even the
finest bony spicule.
 Trim off the excess soft tissues. Boil in tap water
or very dilute (N/100) sodium hydroxide.
 A gross method for hard compact bone is
autoclaving in N/10 sodium hydroxide for 5
minutes.

65.  DEGREASING AND BLEACHING
 After the removal of soft tissue by any of the above
methods, bone is immersed in chloroform for 3-4 hrs
in order to remove fat.
 Specimens are dried in an incubator and bleached in
hydrogen peroxide.
 MOUNTING
 Macerated bones are mounted dry , either on a central
plate or on Perspex supports designed for individual
specimens.
 The specimen is fixed with nylon wires.

68. CALCULI
 Calculi are cut in half with a fine fretsaw , and cut
surfaces polished with sand paper .
 Dry mounting: Cut surface of calculi shows
laminations which can be dry mounted in closed
jars so that it remains dust free.
 Gelatin mounting which can be kept under
Formalin.

69.  Marks are made on Perspex sheet around
positioned stones and holes drilled so that the
stone fits into the gap , which is placed in
position using Perspex cement.
 Then the Perspex sheet is mounted in a glass
jar and the lid is sealed.

70.  Alternative method is to make depressions in
thermocol slab , of the size of the calculi so that the
cut surface and external surface of calculi is stuck on it
with glue.
 Once dry, the thermocol sheet is placed in a clean glass
jar and sealed.

72. AMYLOID
 IODINE TECHNIQUE
 Place slices of formol fixed tissue in Lugol’s iodine to
which 1% sulphuric acid has been added, leave it for 1-
2 hours.
 Wash in running water.
 Mount in liquid paraffin.

73.  CONGO RED TECHINIQUE
 After fixation in Kaiserling I specimens are immersed
in 1 % Congo Red for one hour and are then
transferred to a saturated solution of lithium
carbonate for two minutes.
 They are differentiated in 80% alcohol.
 Normal arteries and veins tend to retain the color.
 Specimens are mounted in Kaiserling III.

74. HAEMOSIDERIN
 Specimens are fixed in Kaiserling I, precautions being
taken to prevent contact with iron rust.
 They are readily stained with a solution of equal parts
of 10% HCI and 5% aqueous potassium ferrocyanide,
and are then washed in running water for 12 hours.

75.  Freshly prepared specimens should be mounted in 5%
formol saline, which prevents color diffusion except
with specimens of haemochromatosis, where for some
reason the color diffuses and the fluid becomes milky
within three months.
 Fat may be demonstrated by staining Sudan III, and
masses of malignant or other tissue by Ehrlich's
haematoxylin, often an excellent way of demonstrating
the limits of lesions.

76. PRESENTATION
 Museum specimens should be clearly labeled and a
system of cataloguing should be employed which
allows easy and rapid access.

77. LABELING
 It may be a rectangle of Perspex sheeting, 1/16 inch in
thickness which is cemented in the centre at the
bottom of the outside of the box, or at the bottom of
the centre plate.
 The advantage of the former method is that alterations
can be easily carried out.

78.  Labels can be of different color; for example red for
undergraduate students, yellow for post graduate
teaching and white for duplicate specimens.
 Alternatively rectangles may be painted on the boxes
with white or colored cellulose paint and on these the
number or diagnosis is painted.

79.  Where specimens are used for examinations, it is best
to employ a numerical system of labeling.
 A good method is a modification of the decimal
system where by the first two figures of a four figure
number indicate the type of conditions; for example,
01, normal organ or congenital lesion; 02, traumatic
and mechanical; 03, acute infection.

80.  The remaining figures divide the subsection, and re-
divide it again.
 Four, five or six figures may be used for this purpose.
 By prefixing the number with a letter the section of the
museum is shown; for example, D, breast; E, nervous
system; F, endocrine.
 CATALOGUING
 A loose-leaf system is essential in order to enable new
specimens to be easily listed.

82. References
 Pulvertaft RJ. Museum Techniques: A Review. J Clin Pathol
1950;3:1-23.
 Proger LW. Perspex Jars for Pathological Museums. J Clin
Pathol 1958;11:92-5.
 Culling CF, Allison RT, Barr WT. Cellular pathology
technique. S.l.London: Butterworth and Cooperations;
1985.
 Indianjournal of pathology and microbiology - 55 (2) , april
- june 2012 pg;261.
 Journal of anatomy soc. India; pg 41-43 (2012)
 Bankroft’s theory and practice of histological techniques
7th edition 2013 chapter 4; pg 69-73