Museum techniques

regarding the fixation and mounting of museum specimens

1. MUSEUM TECHNIQUES 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.

6. 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.

7. 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.

8. 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.

9. 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.

10. 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.

11. 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.

12. 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.

13. 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.

14. 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.

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

16. 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.

17. 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.

18. 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.

19. 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.

20. 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 .

21. 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.

22. 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

23. 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.

24. 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

25. 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.

26. 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 .

27. 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.

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

29. 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.

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

31. 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.

32. 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.

33. 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.

34. 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.

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

36. 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.

37. 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.

38. 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.

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

40. 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.

41. 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.

42. 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.

43. 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.

44. 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.

45. 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.

46. 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.

47. 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.

48. 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.

49. 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

50. 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 .

51. 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.

52. 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.

53. 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.

54. 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.

55. SPECIAL METHODS

56. 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.

57. 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.

58. 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.

59. 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.

60. 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.

61. 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.

62. 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.

63. 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.

64. 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.

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

66. 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.

67. 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.

68. 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.

69. 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.

70. 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 7 th edition 2013 chapter 4; pg 69-73

71. THANK YOU