Notes and queries

Notes and queries

This section contains anonymous questions and signed, peer-reviewed answers to questions on topics within the scope of the journal. Send questions or answers by mail to: Dr JA Kiernan, Department of Anatomy and Cell Biology, The University of Western Ontario, London, Canada N6A 5C1, or by e-mail to [email protected].

Hazel Horn Arkansas Children’s Hospital 800 Marshall, Mail Slot 820 Little Rock, AR 72202 [email protected]

Safe diaminobenzidine (DAB) disposal

Longitudinal sections of unŽ xed muscle

DAB (3,30 -diaminobenzidine hydrochloride), used to localize peroxidase-labeled antibodies and probes, is described as ``possibly carcinogenic’’ in some books and articles. We must assume it is hazardous, so what is a safe way to dispose of used solutions?

I have no trouble with transverse cryostat sections of un®xed skeletal muscle, but longitudinal sections frequently show severely damaged ®bers. How can this be prevented? It does not happen with frozen sections of ®xed muscle.

This procedure, similar to that of Lunn and Sansone (1990), is the one approved by our state water department for disposal of used DAB waste from our Dako autostainer. You might need to check with your own water department before using it.

Frozen skeletal muscle retains the capacity to contract after it thaws. This can lead to problems when examining longitudinal cryostat sections. The tissue literally tears itself apart as portions of the muscle retract toward the areas of the section that ®rst become attached to the microscope slide. No further contraction occurs with drying prior to histochemistry; the damage has already been done. Microscopically, this damage appears as multiple transverse tears in the myo®bers, with the broken ends severely retracted. In areas where the damage is not so severe, partial tearing is apparent within the sarcolemmal sheath. Muscle requires calcium ions (Ca2 ‡) to contract. A chelating agent such as ethylenediaminetetraacetate (EDTA) can combine with Ca2‡, incorporating the ions into a stable, biologically unreactive complex [Ca(EDTA)]2 ¡ ion. EDTA can be used to prevent contraction in un®xed sections of skeletal muscle. The following pretreatment is applied to the glass slides before using them to collect sections from the cryostat:

1.

2.

3.

4. 5.

6.

Prepare the following stock solutions: 0.2 M Potassium permanganate (31.6 g KMnO4 /liter) 2.0 M Sulfuric acid (112 ml concentrated acid/ liter) For each 10 ml of working DAB solution, add: 5 ml 0.2 M Potassium permanganate 5 ml 2.0 M Sulfuric acid Allow the mixture to stand for at least 10 h; this will allow the DAB to become noncarcinogenic. To decolorize the solution, add powdered ascorbic acid until the color disappears. Neutralize the decolorized solution with sodium bicarbonate. Test the neutrality (pH meter, pH paper or dipsticks). Discard by pouring down the drain with copious amounts of running water.

If anyone has a simpler method we would love to have it. By the way, our water department will not allow us to use the procedure using sodium hypochlorite (bleach) to destroy DAB.

1. 2.

3. Reference Lunn G, Sansone EB (1990) Destruction of Hazardous Chemicals in the Laboratory. Wiley Interscience. New York.

4.

Use only clean and grease-free slides. Dip the slides in a 3% w/v solution of Na2 EDTA in distilled water for 5±10 sec. (Na2 EDTA is ethylenediaminetetraacetic acid disodium salt, dihydrate.) Drain well and wipe excess Na2 EDTA solution from the backs and sides of the slides. Allow the slides to air dry at room temperature in a dust-free environment. 229

1. 2. 3.

4.

Notes: Use only slides with a uniform coating of EDTA. It may be necessary to increase incubation times. EDTA can be washed out of the sections by rinsing the slides in buffer for 15 min before proceeding with enzyme histochemistry. If a calcium-activated ATPase technique is to be performed, it is necessary to rinse the slides in barbital buffer, and pre-incubate in 0.04 M calcium chloride in barbital buffer for 15 min. It is also necessary to increase the concentration of calcium ions in the ATPase substrate medium to 150% of the original concentration. Ronnie Houston Bon Secours Health Partners Laboratories 5801 Bromo Road Richmond, VA 23226 [email protected]

Color preservation in pathology museum specimens Is there a way to ®x a large specimen (such as a heart or a slice of whole human kidney) so that the color will not be lost during long storage in a museum jar? The ®xation must also be suitable for microscopic study of small samples taken from the same specimen.

Three steps are involved in preserving a pathological specimen for museum display: ®xation, color preservation or restoration, and specimen preservation. Fixation prevents autolysis and putrefaction and maintains the structural integrity of the specimen. After initial ®xation in phosphate-buffered formaldehyde and collection of small pieces for histological study, the large specimen is placed for an extended period in a museum ®xative formulation, either Wentworth’s or Kaiserling’s (Legualt and Huang 1979, Daws 1996). Wentworth’s ®xative is the easier to prepare and use (Drury and Wallington 1967): Formalin (37%): 100 ml Sodium acetate: 40 g Distilled water: 900 ml Replace the ®xative if it becomes discolored. It is essential that the tissues be fully ®xed (usually 3±4 weeks) in this ¯uid. The natural color of animal tissues is largely due to hemoglobin (red blood) or myoglobin (pink muscle). Fixation in formaldehyde immobilizes these 230

Biotechnic & Histochemistry 2002, 77(4): 229± 232

chromoproteins within cells, but with prolonged storage they degrade and become brown. The simplest method for restoring color to specimens is that of Kaiserling (Drury and Wallington 1967), which uses controlled treatment with ethanol. Fixed specimens are placed in 80% ethanol for 30 min to a maximum of 4 h until the change in color from brown to red is at its maximum. Care is needed because prolonged treatment with ethanol can result in irreversible bleaching of the specimen. The ®nal step is preservation and mounting of the specimen. Most techniques use a 20±40% aqueous glycerine solution containing formalin (less than 1%) or thymol as an antimicrobial preservative (Edwards and Edwards 1959, Drury and Wallington 1967, Daws 1996). Sodium acetate (10%) and dibasic sodium phosphate (ca. 0.1%) are often included to prevent acidi®cation; the pH should be 7.5±8.0. Sodium dithionite (3 g/kg of tissue) may be added to prevent oxidation and to maintain specimen color. Problems associated with this method include lowering of the ¯uid level owing to water loss from leaky or porous plastic containers and leaching of tissue pigments (e.g., blood and bile) into the ¯uid. Israel and Young (1978) recommend using liquid paraf®n (mineral oil) as a preservative solution. Tissues are removed from the 80% ethanol, rinsed quickly in absolute ethanol, blotted dry, and placed in the oil. The resulting specimens are brilliant, with no leaching of pigment from the tissues.

References Daws JJ (1996) Museum techniques. In: Bancroft JD, Stevens A, Eds. Theory and Practice of Histological Techniques. 4th ed., Churchill Livingstone, New York. pp. 699±704. Drury RAB, Wallington EA (1967) Carleton’s Histological Technique. 4th ed, Oxford University Press, New York. pp. 390±395. Edwards JJ, Edwards MJ (1959) Medical Museum Technology. Oxford University Press, London. pp. 77±82. Israel MS, Young LF (1978) Use of liquid paraf®n in the preservation of pathological specimens. J. Clin. Pathol. 31: 499±500. Legault JM, Huang SN (1979) Colour preservation of gross specimens for teaching and medical illustration. Arch. Path. Lab. Med. 103: 300±301.

Tony Henwood The Children’s Hospital at Westmead Locked Bag 4001 Westmead, NSW 2145 Australia [email protected]

Collagen type I staining

Processing dried out specimens

What is the most reliable antibody for detecting type I collagen in human and mouse tissue?

I have been asked to examine sections of dried ®sh (salted and unsalted) by light microscopy. How should I set about processing to obtain paraf®n sections?

If you want to save time, effort and money, why not try staining with picro-sirius red? The method, introduced by Puchtler et al. (1973), is no more dif®cult to do than a van Gieson’s stain. The reagents are inexpensive, and no harsh treatments such as antigen retrieval are needed. Stain hydrated sections for 30 min in:

Sirius red F3B (CI 35780; Direct red 80): 0.1 g Saturated aqueous picric acid solution: 100 ml (This solution can be kept and used repeatedly for at least 4 yr (Kiernan 1999)). Rinse in acidi®ed water (0.1% acetic acid), dehydrate in 3 changes of 100% alcohol, clear and cover.

Zimmermann (1976) showed that dried out ®xed tissues could be rehydrated for histopathological examination by soaking in Ruffer’s solution. This mixture was devised originally to enable the microscopic study of tissues from ancient Egyptian mummies (Ruffer 1921). Ruffer’s solution:

Water 50 ml Ethanol (100%) 30 ml 5% Aqueous sodium carbonate 20 ml (Alternatively, dissolve 0.6 g sodium carbonate in 42 ml water and add 18 ml alcohol, to obtain 60 ml of solution.)

With this technique, all collagen is stained red, but only Type 1 is birefringent with crossed polars (Junqueira et al. 1979). The yellow and orange ®bers stand out brightly on a dark background. The thinnest (reticular) ®bers have green birefringence. It looks like ¯uorescence, but it never fades and the slides can be kept forever. Basement membranes, in which the predominant collagen is type IV, are stained red, but are not birefringent. If picro-sirius red is preceded by nuclear staining with Weigert’s iron-hematoxylin, this should be overdone, because much of the color is extracted from nuclei during the 30 min immersion in picric acid solution.

Rehydrate for 8 h or longer, depending on the size of the specimen, then proceed with dehydration, clearing and embedding. At a National Society for Histotechnology Workshop, Zimmerman (1993) demonstrated rehydrated Egyptian mummy lung with ¯exible, normal looking tissue. The connective tissue framework, bacteria and parasites could be seen in appropriately stained sections, but nuclei and other intracellular structures were lost to autolysis. In contrast, normal and abnormal cells are quite well preserved in desiccated and rehydrated ®xed specimens (Zimmermann 1976).

References

References

Junqueira LCU, Bignolas G, Brentani RR (1979) Picrosirius staining plus polarization microscopy, a speci®c method for collagen detection in tissue sections. Histochem. J. 11: 447±455. Kiernan JA (1999) Histological and Histochemical Methods: Theory and Practice, 3rd ed. Oxford: Butterworth-Heinemann. p. 150. Puchtler H, Waldrop FS, Valentine LS (1973) Polarization microscopic studies of connective tissue stained with picro-sirius red FBA. Beitr. Path. 150: 174±187.

Ruffer MA (1921) Studies in the Paleopathology of Egypt. University of Chicago Press. Chicago. Zimmermann MR (1976) Rehydration of accidentally desiccated pathology specimens. Lab. Med. 7: 13±17. Zimmerman MR (1993) Paleopathology: rehydration techniques, histologic processing and examination overview of paleopathology. Workshop #89, National Society for Histotechnology Symposium/Convention, October, 1993, Philadelphia, PA.

John A. Kiernan Department of Anatomy and Cell Biology The University of Western Ontario London, Canada N6A 5C1 [email protected]

Gayle Callis Veterinary Molecular Biology, Marsh Lab Montana State University, Bozeman 19th and Lincoln St Bozeman MT 59717-3610 [email protected] Notes and queries

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Suppressing auto uorescence What can I do to prevent or reduce troublesome auto¯uorescence in 40 mm frozen sections of formaldehyde-® xed brain that are immunostained with a ¯uorescein-labeled secondary antibody?

The traditional way to quench auto¯uorescence is to pretreat the sections with a solution containing heavy metal ions; osmium tetroxide is the most effective (Ornstein et al. 1957). Ornstein recommended 1% OsO4 for 10 min, but a 0.2% solution for 5 min is also effective (Stoddart and Kiernan 1973). A long wash (at least 4 h in running tap water for slides bearing hydrated paraf®n sections) is then needed to remove the OsO4 . Other techniques use certain dyes, including Sudan black B applied after immuno¯uorescence staining (Schnell et al. 1999, Baschong et al. 2001). Recently, a method has been proposed that avoids the use of chemical quenching. The sections, mounted on slides, are irradiated for 12±48 h with visible and near-UV light from ¯uorescent tubes. The irradiation, which causes fading of the unwanted auto¯uorescence, is done before immunostaining. The original paper (Neumann and Gabel 2002) provides detailed instructions for this ingenious and simple procedure. References Baschong W, Suetterlin R, Laeng RH (2001) Control of auto¯uorescence of archival paraf®n-embedded tissue in confocal laser scanning microscopy (CLSM). J. Histochem. Cytochem. 49: 1565±1571.

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Neumann M, Gabel D (2002) Simple method for reduction of auto¯uorescence in ¯uorescence microscopy. J. Histochem. Cytochem. 50: 437±439. Ornstein L, Mautner W, Davis BJ, Tamura R (1957) New horizons in ¯uorescence microscopy. J. Mount Sinai Hosp. 24: 1066±1078. Schnell SA, Staines WA, Wessendorf MW (1999) Reduction of lipofuscin-like auto¯uorescence in ¯uorescently labeled tissue. J. Histochem. Cytochem. 47: 719±730. Stoddart RW, Kiernan JA (1973) Histochemical detection of the a -D-arabinopyranoside con®guration using ¯uorescent-labeled concanavalin A. Histochemie 33: 87±94.

John A. Kiernan Department of Anatomy and Cell Biology The University of Western Ontario London, Canada N6A 5C1 [email protected]

Answers needed: Is there a silver method for staining Alzheimer’s neuro®brillary tangles that can be relied on to work every time? Is it possible to obtain red staining of peroxidaselabeled secondary antibodies without using a proprietary product whose composition is a trade secret?