QUESTIONS AND ANSWERS

Staining, histochemistry and histotechnology FAQ

(Frequently Asked Questions)  Version  2.0    November 2019

compiled by

Click here for a note on revisions in Version 2.0
J. A. Kiernan,
Department of Anatomy and Cell Biology,
The University of Western Ontario,
London, Canada

Biological Stain Commission  - Home Page

Introduction

The questions and answers are grouped into six categories. Click on a a link below to go directly to a category.

Fixation, freezing etc.Processing, decalcifying, embeddingSectioning, slide adhesives, mounting
Staining methods, histochemistryImmunohistochemistryMiscellaneous questions and answers
Each category contains a number of items. An item is flagged with a title line that begins with two asterisks; for example:

              ** Glycogen, fixation.

(These titles are listed in the Table of Contents, below.)

In an attempt to cut down on spam, all the email addresses in this document have the @ sign replaced with [AT]. If you need to send an email to one of these addresses, edit it to restore the @.

Each item begins with a Question (sometimes more than one, if they are closely related), which is followed by one or more Answers. Items vary in length. Most consist of one or two screenfuls of text. A few topics are differently treated, to achieve a more effective way to answer some questions.

It should be noted that the intention of this FAQ is to explain things, not to provide a compendium of favorite recipes. There are textbooks, and also other web sites, that provide detailed instructions for making solutions and performing techniques. The properly educated technician, pathologist or research worker understands the reason for each step in a procedure. Simple modifications will always be needed to adapt "standard" methods to particular applications. Often, a little study and a lot of thought can shorten the trial-and-error approach to staining

Visitors to this web site are encouraged to submit new questions (or, better still, questions with answers) to be considered for inclusion in later releases of this FAQ. Corrections and other suggestions will also be welcome. Questions and comments may be sent to the compiler by email:  kiernan[AT]uwo.ca.

Acknowledgements

I thank the many people who have answered my questions about staining and related methodology at the Universities of Birmingham (UK), Cambridge (UK) and the University of Western Ontario (UWO, London, Canada). I also thank those students and colleagues, especially at UWO, who asked my advice and made me think and investigate. In recent decades the HistoNet listserver has been another valuable source of questions and answers, and I am grateful to many of its contributors who have kindly allowed me to reproduce their wisdom here.  The Questions in this FAQ are all anonymous, but the sources of the Answers are all acknowledged.

Permission was requested and granted for all the Answers provided by people other than myself. This involved much exchanging of emails, which may not always have been received and answered. It is therefore possible that I have erred by including a few Answers without written permission.  If, gentle reader, you see yourself quoted without consent in this FAQ, please email me (kiernan[AT]uwo.ca). I will immediately expunge the offending Answer and try to find another, perhaps your suggested change, to replace it.  Since the launch of this FAQ on the  BSC's web site in 2006,  questions and answers have been added and modified (Versions 1.0 to 1.6), but  no  contributor has asked  me to remove an item.  This  revision  (Version 2.0)  was  prepared in 2019 for inclusion in a  new  presentation  of the  BSC's  web site.  



Table of Contents.


FIXATION, FREEZING ETC

    **  Carbodiimides as fixatives
    **  Carnoy & alcoholic fixatives
    **  Perfusion fixative for electron microscopy
    **  Fixation of frozen sections.
    **  Non-formaldehyde commercial fixatives
    **  Glutaraldehyde and immunohistochemistry
    **  Isopentane: alternative names
    **  Lidocaine in perfusion fixation
    **  Michel's fluid for transporting cells or specimens
    **  Microwave ovens: Advice for new users
    **  Paraformaldehyde: why won't it dissolve?
    **  Saccomano's fixative
    **  Zinc-containing fixatives: What has been published?
    **  Alternatives to mercury-containing fixatives

 PROCESSING, DECALCIFYING, EMBEDDING

    **  Solvent to replace xylene AND alcohols
    **  2-butoxyethanol ("Clereum") dehydrating or clearing agent
    **  Decalcification: Acid or EDTA?
    **  Testing for completeness of decalcification
    **  Fatty specimens: Processing into paraffin.
    **  Polymethyl methacrylate embedding for bone
    **  Mold release spray
    **  Paraffin processing of skin
    **  Cryoprotection of specimens
    **  Cutting sections of toe or finger nails
    **  Paraffin wax: crystals, additives and cutting
    **  Xylene substitutes: what are they?
    **  Test for water in used absolute alcohol
    **  Molecular sieves for making anhydrous solvent

 SECTIONING, SLIDE ADHESIVES, MOUNTING

    **  Sections coming off slides. Which adhesive?
    **  Apathy's mounting medium and variants
    **  Silanized (APES or TES or positively charged) slides
    **  Polishing undecalcified bone sections.
    **  Polylysine-coated slides
    **  Wrinkles in plastic sections

    **  EM and semithins after paraffin embedding
    **  Wrinkles in paraffin sections containing cartilage
    **  Separation of paraffin ribbon from sections
    **  Thick paraffin sections
    **  Sectioning plastic-embedded specimens
    **  Iodine for removing mercury deposits
    **  Labeling slides
    **  Sectioning plant material: some hints.

 STAINING METHODS, HISTOCHEMISTRY

    ** Making aldehyde-fuchsine
    ** Phosphatases in decalcified, embedded tissue.
    ** Congo red for amyloid
    ** Cartilage staining with safranine
    ** Stain for Chlamydia (Castaneda's method).
    ** Which staining method for copper is best?
    ** Diastase (amylase) control for glycogen
    ** Evans blue, trypan blue and eosin as tracers.
    ** Gallyas' stain
    ** Gram staining of sections (Brown & Hopps method).
    ** Oxidants for hematoxylin
    ** McFaydean's stain for anthrax bacilli
    ** Microglia with Griffonia lectin.
    ** Picro-sirius red staining
    ** Iron hematoxylin: ripening not needed.
    ** Enzyme histochemistry on cell cultures
    ** Malachite green in stain for Cryptosporidium
    ** Confusing dye names (lissamine fast red as an example)
    ** Mayer's and Gill's hematoxylins
    ** Effects of pH on staining by dyes
    ** Histochemical stain for arsenic
    ** Giemsa staining of blood smears: several hints
    ** Automated H & E staining problems
    ** Verhoeff's stain for myelin and elastin
    ** Acridine orange method for DNA and RNA
    ** Quickly finding something in a newly cut section
    ** Fluorescent lectins: general method
    ** Methyl blue and methylene blue

 IMMUNOHISTOCHEMISTRY

    ** Paraffin or frozen sections for immunohistochemistry
    ** Inhibiting endogenous peroxidase
    ** Using mouse primary antibodies on mouse tissues
    ** Antigen retrieval: A patented or copyright phrase?
    ** p53 protein
    ** Prevention of fluorescence fading
    ** Background in immunostained cartilage
    ** Endogenous biotin in mast cells?

 MISCELLANEOUS QUESTIONS AND ANSWERS

    ** Disposal of used diaminobenzidine (DAB) solutions
    ** Dilution of concentrated acids: formula etc.
    ** Disposal of waste from "special stains"
    ** Magnification of a photomicrograph
    ** Can a method be both published and patented?
    ** Books and articles about artifacts in histology
    ** How dangerous is picric acid?
    ** Which color print film for photomicrography?

 


FIXATION, FREEZING ETC

 

** Carbodiimides as fixatives

Question.

What is a carbodiimide, and how does it work?

Answer.

The name "carbodiimide" is sometimes applied to cyanamide (hydrogen cyanamide, H2NCN. Don't confuse this with calcium cyanamide, CaNCN.), which does not seem to have been used as a fixative.

Carbodiimides are compounds that combine with and cross-link carboxyl groups. They fix proteins by joining together C-termini and/or side chains of glutamic and aspartic acid units. Their general chemical formula is R-N=C=N-R'

In contrast, aldehydes combine mainly with protein nitrogen atoms. Cross-links between the lysine side-chain amino group and the amide nitrogens of peptide linkages are thought to do most of the fixing.

Various carbodiimides have been used as fixatives over the years, but they have never caught on in a big way. They are the sort of things used when more ordinary compounds are unsuitable. See Pearse's Histochemistry, Vol. 1 (3rd ed., Churchill-Livingstone, Edinburgh, 1980) page 107 for a proper account.

If the antigenicity of a protein is critically dependent on free amino groups of an epitope, then one of the carbodiimide fixatives might be a sensible alternative to formaldehyde. If it's for paraffin sections, a chemically unreactive fixative such as Clarke's or Carnoy's might be even more sensible.

John A. Kiernan,
Department of Anatomy & Cell Biology,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Carnoy & alcoholic fixatives

Note: The answer to Question 2 discusses the suitability of alcoholic and other fixatives for immunohistochemistry.

Question 1

Any thoughts on the shelf life/keeping qualities of Carnoy's fixative?

Answer.

I always make Carnoy's fixative fresh just before use. Otherwise you will find that the fixing properties will vary if the solution is kept for any length of time. Making up a fresh solution really only takes a few minutes unless you are talking about Lebrun's modification in which the solution is saturated with mercuric chloride.  Carnoy =  ethanol (60) + chloroform (30) + acetic acid (10).

Barry Rittman
(barry.r.brittman[AT]uth.tmc.edu)

Question 2

Are alcoholic fixatives suitable for immunohistochemistry?

Answer.

Fixatives containing ethanol are generally not all that great for IHC. About 4-5 years ago we experimented with several fixatives in an attempt to find one that would give us the cellular morphology that we were used to and also be optimal for IHC/ICC. We tested out the following fixatives:


The 10% NBF of course gave us the morphology we were used to, and if fixation times were kept to 24-48 hours, epitope retrieval was not required for most antibodies. If tissues needed to be stored longer than 48 hours, they were stored in 70% EtOH until ready to be processed. Of all the fixatives we tested, the worst for IHC was 70% EtOH, then Carnoy's. The best for IHC was 70% MeOH. Cellular morphology for both of these was not all that great. Methacarn gave us both good morphology and good IHC. The zinc formalins gave excellent morphology in many organs, and good IHC staining. It should be noted that the zinc formalins have difficulty penetrating the hematopoietic organs as they react more with the RBCs and therefore penetration is much slower. As those are the organs of interest in our laboratory, we use standard NBF.

We have found that if the tissues are trimmed to a thickness of no more than 3 mm prior to immersion in NBF, fixative solutions are changed at 1 and 12 hours, and after 24 hours in fixative are transferred to 70% EtOH, both cellular morphology and IHC staining are excellent.

One of these days when I have some time I'd like to try some of the other fixatives, as well as some of the commercial ones that are out there, just to see what the total comparisons are going to be like. I would also like to note that Bouin's has seemed to work pretty much all right as I have been doing IHC on some Bouin's fixed testes lately without problems.

Robert Schoonhoven
Laboratory of Molecular Carcinogenesis and Mutagenesis
Dept. of Environmental Sciences and Engineering
University of North Carolina CB#7400
Chapel Hill, NC 27599

Back to Table of Contents

** Perfusion fixative for electron microscopy

Question.

What is a suitable fixative for vascular perfusion of rats, and subsequent electron microscopy of tissues?

Answer.

A neutral, buffered, isotonic formaldehyde-glutaraldehyde mixture should be fine for any kind of electron microscopy. Many workers like to use paraformaldehyde as the source
of formaldehyde.

A classical mixture is M. J. Karnovsky's (J. Cell Biol. 27: 137A-138A, 1965). This is probably the most-cited unrefereed abstract! It contains approximately 4% formaldehyde
and 5% glutaraldehyde in approximately 0.1 M phosphate or cacodylate buffer. Final pH = 7.2. If cacodylate (toxic!) is used, add calcium chloride (0.5 mg/ml) to improve
preservation of membrane phospholipids.

Probably this fixative is frequently misquoted, and the literature is full of references to "half-strength Karnovsky," which probably means half the glutaraldehyde concentration. A glutaraldehyde concentration of 1 to 2% is commonly considered adequate in mixtures of this kind.

John A. Kiernan
Department of Anatomy & Cell Biology
The University of Western Ontario
London, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Fixation of frozen sections

Question.

What is the best fixative for frozen sections?

Answer.

Unfixed tissue, cut with a cryostat (thin sections) or a vibrating microtome (thick sections) should be fixed if this is compatible with the staining technique to be used.

Many enzyme histochemical methods demand unfixed sections, and so do immunohistochemical methods with some (fortunately not most) primary antibodies. Enzyme incubations are often terminated by moving the slide or coverslip bearing the cryostat section from the incubation medium into a neutral, buffered formaldehyde fixative.

Even "minimal" (= inadequate) fixation before staining will greatly improve the structural preservation of tissue. Many enzymes will survive either a minute or two in neutral,
buffered formaldehyde, followed by a wash in buffered saline. Some enzymes and most antigens will survive immersion of the slide or coverslip in cold (about 0 C) acetone for half a minute. The acetone is allowed to evaporate before immersing the section in incubation medium.

Cryostat sections may also be fixed by heating, but this inactivates most enzymes. A drop of an ethanol-poly(ethylene glycol) mixture is placed on the section and the temperature
brought up to 55 C in a microwave oven. (A special laboratory oven is needed to get this amount of control.)

References.

Kiernan JA 2015. Histological and Histochemical Methods. Theory and Practice.  5th ed. Banbury, UK: Scion Publishing.
Kok LP & Boon ME 1992. Microwave Cookbook for Microscopists. Leiden: Coulomb Press.
Pearse AGE 1980. Histochemistry, 4th ed. Vol 1.  Edimburgh: Churchill Livingstone.

John A. Kiernan
Department of Anatomy,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Non-formaldehyde commercial fixatives

Question.

Commercially available fixatives are touted variously as
"non-crosslinking," "less-crosslinking," "formaldehyde-free,"
"better for immunohistochemistry," "less toxic," ,etc., etc.

Is there a recent review, or can someone share a list of
names of commercially available fixatives (supposedly better
for immunohistochemistry) and their vendors?

Answer.

Here are all of the ones that I know about; some of them may be sold under different names by other vendors:

There are two fixatives intended for microwave use:

A rather uncomplimentary comparison of some of these products (Histochoice, KryoFix, Mirsky, NoToX, Omnifix II and STF) has been published (Prento & Lyon, 1997. Commercial formalin substitutes for histopathology. Biotechnic & Histochemistry 72:273-282). Readers should note that none of them were used as directed or intended by the manufacturers (fixation at 4 degrees C), so the results are questionable. Also, none of the glyoxal-based fixatives (GlyoFix, Prefer, SafeFix II, Preserve) were tested; these seem to be the most favored substitutes in the USA at least, because they most nearly mimic the morphological patterns obtained with formalin without formaldehyde's unfavorable effects on immunoreactivity.

Richard W. Dapson
Formerly of Anatech Ltd, Battle Creek, MI 49015
(dick[AT]dapsons.com)

Back to Table of Contents

** Glutaraldehyde and immunohistochemistry

Question.

Does glutaraldehyde fixative (4% paraformaldehyde, 0.5% glutaraldehyde) interfere with fluorescent immunohistochemistry?

Answer 1.

Glutaraldehyde, because of its reactivity and speed, can seriously interfere with antibody binding and lectin binding causing considerable non-specific binding. It is also difficult to remove excess glutaraldehyde from tissue components. I would not recommend it's use for such studies, as in my hands the results have been inconsistent.

Barry Rittman
(barry.r.brittman[AT]uth.tmc.edu)

Answer 2.

Tissues fixed in glutaraldehyde exhibit increased autofluorescence, which is probably due to glutaraldehyde-amino acid compounds that are formed as part of the fixative action. Glutaraldehyde also introduces free aldehyde groups into the tissue, and these will bind any protein reagents that are applied. The nonspecific binding of antibodies can be reduced by pretreatment with a blocking protein (such as bovine albumin, or serum from the species in which a secondary antibody was raised). Before the blocking treatment it is advisable to do a chemical aldehyde blockade (Histochemistry textbooks contain several methods).

John A. Kiernan
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Isopentane: alternative names

Question.

Is isopentane the same as 2-methyl butane?

Answer.

Yes. It is also known as ethyldimethylmethane All are
 (CH3)2CHCH2CH3.

Anita Jennings
(jennings[AT]mayo.edu)

Back to Table of Contents

** Lidocaine in perfusion fixation

Question.

Lidocaine can be added to the fixative during perfusion I would appreciate hearing the Lidocaine concentration again.

Answer.

This is the recipe for lidocaine I used for perfusion-fixing mormyrids (an electric fish):

Lidocaine (= lignocaine = xylocaine) for use in perfusion fixation (Used to relax blood vessels to permit more complete exchange & infiltration of fixative.):

Note: Do not add the lidocaine directly to the perfusion solution, especially if the solution contains salts! The lidocaine will not go into solution.

Philip Oshel
(oshel[AT]shout.net
or poshel[AT]hotmail.com)

Back to Table of Contents

** Michel's fluid for transporting cells or specimens

Question.

Does anyone have any references for Michel's Fixative or Fluid? We use it for an immunofluorescence holding medium, but I don't have a reference on it.

Answer.

Here's my procedure sheet for Michel's transport medium.

MICHEL'S TRANSPORT MEDIUM

Michel's transport medium (pronounced mee-SHELL) is used to transport specimens (such as renal biopsies and lymph nodes) for immunofluorescence studies. It is not a fixative, and is not suitable for any other use (particularly, it is not suitable for transporting living cells for flow cytometry). It should be stored refrigerated (not frozen) until use. Specimens may be kept in it at room temperature until they can be delivered to the reference laboratory. Zeus Medium, a commercial product, is probably similar.

1.0 M potassium citrate buffer pH 7.0:  

Dissolve 21.0 g citric acid monohydrate (or 19.2 g citric acid anhydrous) in 30 mL of hot deionized or distilled water.  Cool.  Adjust pH to 7.0 with 1 M potassium hydroxide (about 35 mL).  Dilute to 100 mL with more water.

Washing solution:

25 mL 1.0 M potassium citrate buffer
50 mL 0.1 M magnesium sulfate heptahydrate (F.W. 246.5)
50 mL 0.1 M N-ethyl maleimide (= 12.5 g in 1 L of water)  (Sigma E3876.)
Water to make 1 L
Adjust to pH 7.0 with 1 M potassium hydroxide
Store in refrigerator. 

Transport medium:

Dissolve 55 grams of ammonium sulfate in 100 mL washing solution. (Add slowly, with mechanical stirring.)
Adjust pH to about 6.9 with 1 M potassium hydroxide (< 2 mL needed)

Specimens can be held at room temperature for five days in transport medium before processing. Specimens received in transport medium should be washed in three changes of washing solution, 10 minutes each wash.

Reference.

Michel B. Milner Y. David K. 1972. Preservation of tissue-fixed immunoglobulins in skin biopsies of patients with lupus erythematosus and bullous diseases. A preliminary report. J. Invest. Dermatol. 59: 449-452.

This procedure received from J. Charles Jennette MD, Immunopathology Laboratory, North Carolina Memorial Hospital, Chapel Hill NC 27514

Bob Richmond
Samurai Pathologist
Knoxville TN
(RSRICHMOND[AT]aol.com)

Back to Table of Contents

** Microwave ovens: Advice for new users

Question.

Can someone experienced with a microwave processor give advice?

Answer 1.

In making your final decision about the purchase of a laboratory microwave oven, you may also find it helpful to use some simple microwave calibration tools to determine objectively if a particular microwave oven will suit your specific needs.

These tools are quick and simple assessments that show you just how evenly your clinical specimens will be heated in a microwave oven.

1. Neon Bulb Array.
Because our eyes can not sense microwaves, they appear invisible to us. A Neon Bulb Array is a tool that indirectly shows the nonuniformity of microwave power in a microwave oven. In principle, microwave irradiation increases the kinetic energy of the neon gas molecules. The neon bulbs glow orange where the microwave power is high enough to ionize the gas molecules (~5 mw/cm2). The neon bulb array is useful for determining the areas of uniform power, cycle
time, and magnetron warm-up time in a microwave oven

2. The Agar-Saline-Giemsa tissue phantom.
Agar-Saline-Giemsa tissue phantoms are used to simulate the size, shape, and absorbance characteristics of biological specimens to verify that the microwave oven will uniformly heat the specimens.

Small agar phantoms (1 cm x 0.5 cm2 blocks or 2 cm diameter by 0.3 cm thickness discs) that contain 0.002% commercial Giemsa stain are added to molten 2% agar in 0.9% sodium chloride. The Giemsa dyes respond to microwave heating by showing different colors at different temperatures. When ASG tissue phantoms are irradiated in an optimized microwave cavity, they show a uniform color change.

These tools have been described and published in peer-reviewed journals since 1990 and have been independently verified by other laboratories. They are commercially available or you can prepare them yourself.

Brief list of references

1. Login, G. R., N. Tanda, and A. M. Dvorak. 1996. Calibrating and standardizing microwave ovens for microwave-accelerated specimen preparation. A review. Cell Vision 3: 172-179.
2. Login, G. R., and A. M. Dvorak.  1884. The Microwave Toolbook. A Practical
Guide for Microscopists. Boston: Beth Israel Hospital.
3. Login, G. R., J. B. Leonard, and A. M. Dvorak. 1998. Calibration and standardization of microwave ovens for fixation of brain and peripheral nerve tissue. Companion to Methods Enzymol. 15.
4. Login, G. R. 1998. The need for clinical laboratory standards for microwave-accelerated procedures. J. Histotechnol. 21: 1-3 (Editorial).

Gary Login, Assistant Professor of Oral Pathology
Beth Israel Deaconess Medical Center

Answer 2.

My experience thus far is purely from a vendors view. The benefits so far:

  1. You can process without xylene
  2. Turnaround can be minutes as opposed to hours.
  3. Cost savings about 1/5 of a traditional processor (Not counting the reagent savings).
  4. Loads of up to 90 cassettes can be processed in one run.


Dawn M. Truscott, HT(ASCP)
Product Specialist
Carl Zeiss, Inc.
(DayDawning[AT]aol.com)

Back to Table of Contents

** Paraformaldehyde: why won't it dissolve?
(Answer includes other information about formaldehyde and fixation)

Question.

Why will paraformaldehyde not dissolve in unaltered seawater without added sodium hydroxide?

Answer.

Paraformaldehyde is a white solid formed by combination of large numbers of formaldehyde molecules in an aqueous solution: a polymer. Formaldehyde, HCHO, is a gas and strictly speaking it doesn't exist in aqueous solution because it tacks on a water molecule to form methylene hydrate, which is HO-CH
2-OH. This is
the active ingredient of fixatives. Methylene hydrate molecules just love one another, and join together (eliminating H2O, so I suppose it's really the original formaldehyde carbon atoms that are so affectionate) to make polymers of all sizes. In commercial formalin (37-40% HCHO by weight) the polymer molecules are small enough to stay in solution. In paraformaldehyde they are big enough to be insoluble.

Manufacturers add some methanol to formalin. This retards the formation of large polymer molecules (see Recommended Reading if you want to know why). Probably the methanol doesn't affect fixative properties when diluted, though some people in the late 1950s claimed that it did. If you buy paraformaldehyde, you can depolymerize it yourself and get a solution of "formaldehyde" (actually methylene hydrate) that doesn't contain any methanol.

From what I've said so far, _Please Take Note!_ it follows that there is no such thing as a "2% (or any other %) paraformaldehyde" solution. Paraformaldehyde is a high polymer, and its molecules are too big to dissolve in water, alcohol or anything else.

You have to depolymerize paraformaldehyde to get it to "dissolve" and form a formaldehyde (really methylene hydrate) solution. The depolymerization is a reaction of the polymer with water: a hydrolysis. It needs hydroxide ions (OH
) as a catalyst, and also some heat to get the job done in reasonable time. In the making of ordinary phosphate-buffered formaldehyde from paraformaldehyde, the usual procedure is to heat the PF with the dibasic sodium phosphate component of the buffer. This contains enough OH ions to catalyse the hydrolysis and depolymerization. You add the acidic part of the buffer (sodium or potassium dihydrogen phosphate) when the solution has become transparent. This occurs when the temperature reaches about 60C. It should not be
necessary to go any hotter than that.

In the earliest recommended fixatives that started with paraformaldehyde, a few drops of sodium hydroxide were added to a heated suspension of paraformaldehyde in water or saline. This hardly affected the pH of the final solution.

Additional question.

My supervisor (who has been trained in histology, unlike myself) said that in most of my staining and fixative methods the phosphate buffer component could be replaced by seawater, with no problems because seawater is a buffer, at the right osmolarity for fish tissue. Is this the case?

Answer, continued.

I don't know how good a buffer sea water is, but it's unlikely to be as robust as 0.1M phosphate. In a fixative the osmolarity is more important than the pH, but for a slowly acting agent like formaldehyde or a slowly penetrating one like osmium tetroxide, the solvent should be as similar as possible to the extracellular fluids of whatever you're fixing. If the formaldehyde (takes hours to do its stuff) is mixed with more rapidly acting fixative agents (alcohol, mercuric chloride, picric acid etc., which act as soon as they reach the cells), the osmolarity is less important, and most such mixtures are acidic too. The formaldehyde does its cross-linking after the proteins have been insolubilized by the coagulant components.

Readings.

For formaldehyde chemistry: 

Walker, JF 1964. Formaldehyde. 2nd ed. New York: Reinhold; London: Chapman & Hall.

For how formaldehyde works: 

Pearse, AGE: Histochemistry, Theoretical and Applied. Any edition of this book should be OK. 
There's also lots of erudite discussion
in Baker, JR (1958) Principles of Biological Microtechnique. London: Methuen, which is a great classic in the field, now  available free from https://archive.org/details/principlesofbiol01bake/.

For some stuff on the slowness of formaldehyde fixation and importance of an isotonic buffer: 

Paljarvi,L, Garcia,JH & Kalimo,H 1979. Histochem. J. 11: 267-269.
Schook, P 1980. Acta morph. Neerl.-Scand. 18: 31-45. 

See also some of MA Hayat's books on techniques for electron microscopy, which discuss the subject thoroughly.

John A. Kiernan,
Department of Anatomy & Cell Biology,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Saccomano's fixative

Question.

Does anyone have a recipe for Saccomanno fixative (a cytology fixative) which gives the molecular weight of the Carbowax (polyethylene glycol) in the solution? 

Answer.

This formula is from Koss. Roughly equal volumes of Saccomanno's fixative can be added to liquid cytologic specimens such as sputum, urine, bronchial washings, and pleural and peritoneal fluids to stabilize them at room temperature until they can be prepared as filter or cytocentrifuge preparations or cell blocks, and it also works fairly well for small biopsy specimens. It is not suitable for ThinPrep preparations, for which a special fixative is required.

Saccomanno's fixative is 50% alcohol which contains approximately 2% of Carbowax 1540 (Union Carbide Corporation, UCAR). Carbowax 1540 is solid at room temperature, with a melting point of 43 to 46 C. To avoid having to melt it whenever the fixative is prepared, a stock solution can be propared by melting of Carbowax (melted in an incubator or hot air oven at 50 to 100 C) and adding it to an equal volume of water or 50% alcohol. The mixture will not solidify.
Saccomanno's fixative can then be prepared with 430 mL of water, 530 mL of 95% ethanol, and 40 mL of the stock Carbowax solution. Some light green SF or fast green FCF can be added to color the fixative. Koss warns that the denaturants in reagent alcohol may cause excessive hardening of mucus.

I suppose that the 1540 is the molecular weight, but basically it's a catalog number for a long series of these UCAR products that range from thin liquids to dense paraffin-like waxes.

Reference.

Leopold G. Koss, Diagnostic Cytology and Its Histologic
Bases, 3rd ed., Lippincott 1979, page 1192. 
(I don't have the
current edition of this venerable tome. I have never tried to make Saccomanno's fixative, but those who have rank it right up there with hanging wallpaper as a good way to wind up screaming.)

Bob Richmond
Samurai Pathologist
Knoxville TN
(RSRICHMOND[AT]aol.com)

Back to Table of Contents

** Zinc-containing fixatives: What has been published?
(Answers include references, opinions and discussion.)

Questions.

What published work is available with evaluations of zinc-formalin and other such newer fixatives? 

Can a zinc salt really replace mercuric chloride?

Answers.

These questions are discussed quite frequently in the HistoNet listserver group. In February 1998. I wrote that there was a shortage of publications in refereed journals, and also suggested that it was unwise to use a commercial product without knowing its complete composition. (There are published formulations, but in most cases these compare a zinc-containing liquid with neutral buffered formaldehyde, for immunohistochemical detection of one or several antigens. The
exact composition of proprietary fixative mixtures is seldom stated in catalogues etc.)

John Kiernan
London, Canada
(kiernan[AT]uwo.ca)

Dick Dapson disagreed with some of my comments, and provided a helpful list of publications:

John Kiernan wrote  that there is a remarkable shortage of literature comparing zinc formalin solutions with conventional fixatives. Actually, the subject has been covered rather well over a time span of more than 10 years. Here is a sample that shows the evolution of these remarkable fixatives; all are from refereed
journals and (except for the 1981 abstract) have "passed the scrutiny of the regular scientific publication process":

1981. Jones, et al. Transition metal salts as adjuncts to formalin for tissue fixation (abstract). Lab Invest 44: 32A [This is the paper that really started it all, although zinc formulations do appear in the early literature].

1983. Mugnaini et al. Zinc-aldehyde fixation for light-microscopic immunocytochemistry of nervous tissues. J Histoch Cytochem 31: 1435-1438.

1985. Banks. Technical aspects of specimen preparation and special studies. In Surgical Pathology of the Lymph Nodes and Related Organs. Jaffe, ed. W B Saunders Co., pp1-21.

1988. Herman, et al. Zinc formalin fixative for automated tissue processing. J Histotechnol 11: 85-89. [The first really comprehensive study comparing NBF and
unbuffered zinc sulfate formalin].

1990. Tome, et al. Preservation of cluster 1 small cell lung cancer antigen in zinc-formalin fixative and its application to immunohistochemical diagnosis. Histopathol 16: 469-474.

1991. Abbondanzo, et al. Enhancement of immunoreactivity among lymphoid malignant neoplasms in paraffin-embedded tissues by refixation in zinc sulfate-formalin. Arch Pathol Lab Med  115:31-33.

1993. Estrogen and progesterone receptor proteins in zinc sulfate, formalin fixed breast carcionoma: advantages of a supersensitive streptavidin technique. J Histotechnol 16: 51-56.

1993. Dapson. Fixation for the 1990's: a review of needs and accomplishments. Biotechnic & Histochem 68: 75-82. [Like Herman's paper, this provides a critical comparison between NBF and zinc formalin; it also details probable mechanisms and reviews the pertinent literature to date].

1995. L'Hoste, et al. Using zinc formalin as a routine fixative in the histology laboratory. Lab Med 26: 210-214. [Compares NBF and a buffered zinc formalin, using side-by-side color photomicrographs].

Richard W. Dapson, Ph.D.
Formerly of ANATECH LTD.
Battle Creek, MI 49015
(dick[AT]dapsons.com)

My response:

The interested reader should study these publications. Most do not include critical comparisons with other fixatives (except buffered formaldehyde), especially for preservation of intracellular structures. There is a real need for users to compare several fixatives in properly controlled trials, and publish their results.

Zinc mixtures became popular in the early 1990s, but the earliest (probably) of its kind was introduced soon after the fixative action of formaldehyde was discovered by F. Blum (in Germany, in 1893). This is Fish's fixative:

Water: 2000 ml
Formalin: 50 ml
Zinc chloride: 15 g

Fish, Pierre A. 1895. The use of formalin in neurology. Trans. Am. Microsc. Soc. 17: 319-330.  [Fish recommended immersion of the brain for 7-10 days, with
injection of cavities and blood vessels if possible. It's all been done before if you go back far enough! Fish's paper also reviewed the uses of formaldehyde (31 references, only 2 years after it's introduction as a fixative) and he also described other fixative mixtures.]  

J. A. Kiernan 2009. A system for quantitative evaluation of fixatives for light microscopy using paraffin sections of kidney and brain Biotech, Histochem. 84: 1-10.  [Fish's fixative compared favorably with NBF and three other zinc-formalin mixtures for microanatomical preservation. Fish's fixative was inferior to NBF and zinc sulfate-formalin for cytoplasmic and nuclear fixation.]

J. A. Kiernan
Department of Anatomy & Cell Biology
The University of Western Ontario
London, Canada N6K 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Alternatives to mercury-containing fixatives

Question.

What is the best substitute for B-5 fixative, without mercuric chloride?

[ B-5 is: Water 90 ml, Formalin (40% HCHO) 10 ml, Mercuric chloride 6 g, Sodium acetate (anhydrous) 1.25 g.
The sodium acetate brings the pH into the 5.8-6.0 range.
Fix by immersion, 12-24 hours, then transfer to 70-80%
alcohol. See Lillie RD & Fullmer HM 1976 Histopathologic
Technic and Practical Histochemistry. New York:
McGraw-Hill, pp 52-53.]

Answer.

We recently completed a "blind comparison" of B-5 substitutes. We needed to find something, as our water treatment plant had notified us that as part of a Zero Discharge Program they would be monitoring our mercury output. Of course, we were capturing our mercury ... but we still had measurable amounts in our discharged water. The treatment plant immediately zeroed in on our department, and without delay asked if we used mercury fixatives! We agreed that we would cease, or absolutely contain our mercury by June 1, 1998. I felt it better to cease using mercury, so that any future mercury found in the discharge water from
the hospital could be blamed on another source!! We had all of our sink traps cleaned, and tested ... no mercury coming from us!!!

For our study, we used our standard B-5, Z-5, and Z-fix from Anatech, IBF from Surgipath, our 10% NBF, and B-plus fixative from BBC. We used tonsil and lymph nodes for the study, and placed small pieces of tissue in each of the fixatives, and gave them to the pathologists labeled as fixative 1, fixative 2 etc. The pathologists were given an evaluation sheet with each case, and asked to rank the fixatives from 1-6, with 1 being the best. When we had tested a sufficient number of cases, the evaluations were tallied, and lo and behold ... B-5 won! I wasn't surprised, and neither were the pathologists. We all agreed that we would use the second place winner.

This was B-Plus Fix which is sold by BBC (800-635-4477, or write to PO Box 609, Stanwood, WA 98292). However, all the solutions that we tested were acceptable. One surprising result was that our 10% NBF came in 3rd, very close to our 2nd place winner. We have been using our substitute since March, and are pleased with the results so far... However, the pathologists are missing their B-5, which they still refer to as the gold standard.

Sheila Tapper
St. Mary's / Duluth Clinic Health Systems
Duluth, MN
(STapper [AT]smdc.org)

Back to Table of Contents

 


 

PROCESSING, DECALCIFYING, EMBEDDING

 

** Solvent to replace xylene AND alcohols

Question.

Is there a product that replaces xylene AND alcohols in the staining procedure? Can you use it before and after the actual staining is done?

Answer 1.

t-Butanol, dioxane and tetrahydrofuran are miscible with wax, water and resinous mounting media. Of these, only t-butanol (= tertiary butyl alcohol) is suitable for
ordinary use. (The other two have such hazards as fire, toxicity and explosive peroxide formation.) t-butanol is often used in botanical microtechnique; it is quite a bit more expensive than ethanol or xylene. n-Butyl alcohol mixes with wax and mounting media and is also partly miscible with water. It's good when you use easily extracted stains (methyl green-pyronine, for example), but has unpleasant vapour. 2-Butoxyethanol (butyl cellosolve) also has the right miscibilities, and is quite cheap because it's used on a big scale industrially.  For more information about miscibilities of solvents, waxes and mounting media used in microtechnique, see Gray (1954) or Kiernan (2015).

For microwave processing, isopropyl alcohol is sometimes recommended. However, this mixes with wax only at elevated temperatures. It has to leave the specimen by vaporizing under reduced pressure; this can lead to considerable tissue damage unless the temperature and pressure are just right (Bosch et al 1996). Buesa & Peshkov (2009) describe methods using a mixture of isopropyl alcohol and mineral oil for clearing prior to infiltration with wax, water with detergent at 90C for dewaxing slides, and coverslipping air-dried stained sections with resinous mounting medium. 

Some staining methods work well, though slowly, without removing the paraffin beforehand (Kiernan 1996), provided that there has been no melting or softening of the wax after mounting the sections on their slides.

References.

Bosch,MMC; Walspaap,CH; Boon,ME (1996): Lessons from the experimental stage of the two-step vacuum-microwave method for histoprocessing. Eur. J. Morphol. 34(2), 127-130.
Buesa,RJ; Peshkov,MV (2009) Histology without xylene. Ann. Diagn. Pathol. 13: 246-256.

Gray, P (1954) The Microtomist's Formulary and Guide. New York: Blakiston. (Reprinted 1975 by
Krieger, ISBN 0882752472), pages 622-629.
Kiernan,JA (1996): Staining paraffin sections without prior removal of the wax. Biotechnic & Histochemistry 71(6), 304-310.  
Kiernan JA (2015) Histological and Histochemical Methods. Theory and Practice. Banbury, UK: Scion, pages 54-55.


John A. Kiernan
London, Canada
(kiernan[AT]uwo.ca)

Answer 2.

We use 99% isopropyl alcohol (IPA) instead ethanol AND xylene AFTER staining. It is especially useful after staining of lymph nodes with a modified Maximov-Giemsa method. My laboratory has used this modification more then 5 years and I have never seen the same excellent result in comparison with atlases of lymph nodes biopsy. Moreover, we use IPA with addition of a small amount of detergent for dehydration of samples. Four changes of  99% IPA+detergent is all you need between water and paraffin. We never have problems with any tissues, including large samples of skin. Our HTs adore IPA.

Dr Yuri Krivolapov
Military Medical Academy
St.-Petersburg, Russia
(krivolapov[AT]bfpg.ru)

Back to Table of Contents

** 2-Butoxyethanol ("Clereum") dehydrating or clearing agent

Question.

What are the properties of Clereum? (The MSDS for Clereum indicates the ingredient information as undiluted 2-butoxyethanol.)

Answer.

It's good to learn that this isn't yet another secret clearing agent! According to the Merck Index, this compound (also called butyl cellosolve, or ethylene glycol monobutyl ether) is partly
miscible with water. Its properties as a solvent seem to be similar to n-butanol; no doubt the higher B.P. (171C) is an advantage - it won't have n-butanol's nasty cough-making vapour.
Merck says the toxicity is similar to methyl cellosolve (anaemia, "CNS symptoms" etc; can be absorbed through skin).

The price of 2-butoxyethanol varies with the supplier. The "scintillation grade" costs 6 to 8 times as much as "99%" and "laboratory" grades. 

If the 99% stuff is OK for histology, perhaps the price isn't too bad; somewhat less expesive than tert-butanol or n-butanol (99%) is $27 for 2.5 litres. This makes 2-butoxyethanol quite a good buy for a non-niffy not-quite-universal solvent. The similarity of its miscibilities to those of n-butanol suggests that this might be useful for dehydrating (and clearing) sections that have been stained with methyl green-pyronine, or other dyes that are easily lost with ordinary alcoholic dehydration.

John A. Kiernan
Department of Anatomy,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Decalcification: Acid or EDTA?

Questions.

How should I decalcify a bony specimen or a tooth?
What precautions are needed if galactosidase activity must be preserved (to identify cells carrying the LacZ gene)?

Answer 1.

Decalcification with EDTA is probably the best method with your LacZ, due to the enzyme staining you are doing. I would be careful to adjust the pH of the EDTA solution to the working pH of enzyme staining in PBS or a TRIS buffer, and rinse carefully in buffers postdecalcification. Formic acid may ruin LacZ enzyme staining results.

Gayle Callis
(gayle.callis[AT]bresnan.net)

Answer 2.

If the bone is crunchy, you have either not removed all the bone mineral, or you have transferred the bones from EDTA to alcohol and have precipitated EDTates in your tissue.

When you decalcify, do you determine the end point using an x-ray/calcium oxalate/prod with a pointed stick?

How long do you decalcify? Even at 20% EDTA these would take at least a week with vigorous agitation at room temperature. Is the EDTA buffered to pH 7? If not, you are using the solution as an acid decalcifier as well as a chelator. In this case, assuming your stain still works and will not be affected by acid pH, change to 10% formic acid, which provides much faster decalcification. Check the endpoint (when all the calcium is gone) daily.

[ But see Answer 1 for acid-sensitivity of galactosidase. ]

If you have checked the endpoint and all the calcium is gone, rinse the tissue in water for at least 8 hours to remove all the excess EDTA before putting it in alcohol.

Simon Smith
(smiths5[AT]pfizer.com)

Answer 3. (A formic acid procedure for teeth, with oxalate testing)

The protocol we use here at Indiana Univ. School of Dentistry is as follows:

After teeth are fixed in 10% neutral buffered formalin, they are placed in wide mouth bottles with a 5% formic Acid solution. They are then checked each day by pipetting 5 ml of the acid solution into a test tube to which 1 ml of 2.5% ammonium oxalate is added. If a white precipitate forms there is still calcium present. The solution is then changed and the process repeated the next day. Once I get one negative test the specimen is grossed as needed and placed back into acid until another negative is obtained. The specimen is then placed in running water overnight and processed with the next days run. I know this can take a long time, but the results are worth it. If you need anything else let me know.

Lee Ann Baldridge
IUSD Oral Path Group
Indianapolis, IN.
(lhadley[AT]iusd.iupui.edu)

Back to Table of Contents

** Testing for completeness of decalcification

Questions.

How should I test for complete decalcification?
Is the same method OK after either formic acid or EDTA?

Answer.

The ammonium oxalate test is simple. Take a 5 ml sample of used decalcifying fluid. Neutralize it by adding drops of strong ammonia (ammonium hydroxide); avoid the fumes! When the solution turns litmus blue (pH above 7), add 5 ml of saturated aqueous solution of ammonium oxalate (about 3%; stable stock solution). Wait for half an hour. If there is no precipitate, the last change of decalcifying fluid was free of calcium ions.

According to Eggert & Germain (1979) you can use the ammonium oxalate test on EDTA. Rosen (1981) said the sensitivity was higher if you lowered the pH to 3.2-3.6 before doing the test (instead of neutralizing to pH 7 as done with an acid decalcifier).

Eggert FM, Germain JP 1979. Rapid demineralization in acidic buffers. Histochemistry 39: 215-224.
Rosen AD 1981. End-point determination in EDTA decalcification using ammonium oxalate. Stain Technology 56: 48-49.

John A. Kiernan,
Department of Anatomy & Cell Biology,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Fatty specimens: Processing into paraffin

Question.

What is the best way to paraffin-embed specimens that contain a lot of fat?

Answers.

1. Process by hand, allowing more time and bigger volumes of all solvents than for non-fatty pieces of tissue.

2. Don't put them through an automatic processor because you'll get grease in all the solvents. (If you don't believe this, put a bit of skin in about 10 times its volume of 95% alcohol for an hour, then add some water to the alcohol. Result: a milky emulsion.)

3. Xylene is better than a "xylene substitute."

[ Unfortunately I mislaid the sources of these pieces of
advice. For what it's worth, I agree strongly with
the first two, but lack the experience to comment on
the third. J. A. K. ]

Back to Table of Contents

** Polymethyl methacrylate embedding for bone

Question.

Is it permissible to mix polymerized methyl methacrylate with the monomer, when making an embedding medium for undecalcified bone?

Answer.

Using polymethylmethacrylate powder or beads does not affect the polymerization process, but it does make the preparation of the partly polymerized embedding mixture easier and safer.  You may care to refer to the following paper.


Difford, J. (1974) A simplified method for the preparation of methyl methacrylate embedding medium for undecalcified bone. Medical Laboratory Technology 31: 79-81.

John Difford
Royal Free Hospital
London, England.
(adford[AT]compuserve.com)

Back to Table of Contents

** Mold release spray

Question.

Is there something you can spray into an embedding mold to make it easier to extract the solidified wax block?

Answer.

I faced the problem of mold-release spray several years ago by mixing a solution of 5% green dishwashing soap (such as Palmolive) in 50% Ethanol, then putting it into a pump spray
bottle (available form any housewares department). This worked AT LEAST as well as the outrageously expensive stuff sold as "Mold-Release Spray", and it contained no CFC's or
other "evils".

Joanne Lahey
Battelle Duxbury Operations
Duxbury, MA 02332
(laheyj[AT]battelle.org)

Back to Table of Contents

** Paraffin processing of skin

Question.

Could you suggest a processing schedule suitable for skin?

Answer.

These are my processing schedules for skin dehydration and embedding.

By hand:

The times suit my working day. I'm sure they could be altered for any work pattern.

1.)   80% alcohol. = 2 pm.
2.)   80% alcohol. = 5 pm - overnight.
3.)   Abs.alc./8% phenol. = 9 am.
4.)   Abs. alcohol. = 10 am.
5.)   Abs. alcohol. = 12 am.
6.)   Abs.alc./amyl acetate.= 3 pm.
7.)   Amyl acetate. = 4 pm.
8.)   Amyl acetate. = 5 pm. - overnight.
9.)   Amyl acetate. = 9 am.
10.) Amyl acetate. = 12 am.
11.) Xylene. = 3 pm.
12.) Wax. = 4 pm.
13.) Wax. = 5pm. - overnight.
14.) Wax. = Embed.

Using a tissue processor:

These are the times I use on a Shandon Histokinette. 

1.)   80% alcohol. = 2 hours.
2.)   80% alcohol. = 2 hours.
3.)   Abs. alc./8% phenol. = 1 hour.
4.)   Abs. alcohol. = 3 hours.
5.)   Abs. alcohol. = 3 hours.
6.)   Abs.alc./amyl acetate.= 1 hour.
7.)   Amyl acetate. = 3 hours.
8.)   Amyl acetate. = 3 hours.
9.)   Amyl acetate. = 3 hours.
10.) Amyl acetate. = 5 hours.
11.) Xylene. = 1 hour.
12.) Wax. = 9 hours.
13.) Wax. = 9 hours.
14.) Embed.

Ian Montgomery
University of Glasgow, Scotland
(I.Montgomery[AT]bio.gla.ac.uk)

Back to Table of Contents

** Cryoprotection of specimens

Question.

Please recommend a way to protect formaldehyde-fixed mouse brains to avoid cracks and ice crystal holes that form during rapid freezing. 
25% sucrose has been
recommended. Should it be in water or phosphate-buffered saline?

Answer 1.

For ultracryomicrotomy (or should it be cryoultramicrotomy) Tokuyasu (1989) used 2.3 M (= 78%) sucrose in 0.1M phosphate buffer. He was working with blocks much smaller than mouse brain, so you will no doubt have to increase the time. Inflitration of blocks 1 mm wide usually took 30 minutes. He stated that infusion was complete when the specimen no longer floated on the top of the sucrose solution. The same author reported that 10-30% PVP and 1.6-2M sucrose provided still better postfreezing conditions (compared with freezing alone).

We presently use 5% PVA (polyvinyl alcohol) in phosphate buffer to cryoprotect bone samples before freezing for enzyme and immunohistochemistry.

One other point that may be worth considering is the method for freezing. If you are thinking of snap-freezing, I would recommend hexane instead of isopentane. Hexane freezes at a
considerably higher temperature: about 80 C. Many moons ago, when I worked in Neuropathology in Scotland, I found that mouse brains tended to crack when frozen in isopentane, but that we had much better preservation when freezing in precooled hexane (we never cryoprotected them though).

Reference.

Tokuyasu KT. 1989. Use of polyvinylpyrrolidine and polyvinyl alcohol cryoultramicrotomy. Histochem. J. 21: 163.

Ronnie Houston
Dallas, Texas
(RHH1[AT]airmail.net)

Answer 2.

It is a common practice to immerse rodent brains in 20-30% sucrose at 4 C, at least until they sink. If they have been fixed for only a short time (less than 48 hours), it is probably best to dissolve the sucrose in PBS rather than water alone.

Rosene et al (1986) found that 20% glycerol with 2% dimethylsulfoxide (DMSO) was better than sucrose. The sucrose concentration needs to be much higher than is commonly used - at least 60% (see Lepault et al, 1997).

References (with brief notes).

Rosene,DL; Roy,NJ; Davis,BJ (1986): A cryoprotection method that facilitates cutting frozen sections of whole monkey brains for histological and histochemical processing without freezing artifact. J. Histochem. Cytochem. 34: 1301-1315.
    Techniques compared. Optimum cryoprotection with 4 day infiltration (4 C) of 20% glycerol & 2% DMSO in buffer or fixative. Then freeze in isopentane at -75 C (dry ice). Better than other cryoprotectants (sucrose etc) and freezing methods.

Lepault,J; Bigot,D; Studer,D; Erk,I (1997): Freezing of aqueous specimens: an X-ray diffraction study. J. Microsc. (Oxford) 187(Sep), 158-166.
    EM & X-ray diffraction of freezing of sucrose solutions. Immersion in a liquid cryogen or high pressure freezing. Sucrose favours formation of amorphous ice; conc must be 60% or above for freezing in a cryogenic liquid.

John A. Kiernan
London, Canada
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Cutting sections of toe or finger nails

Question.

Does anyone have a few hints for sectoning toenails?

[ Here are 5 of many replies to this frequently asked question. ]

Answer 1.

10% Potassium hydroxide. Soak them for at least 4 hrs, but not more than 8.

Noreen S. Gilman (n4xiu[AT]gate.net)

Answer 2.

I have not cut toenails for years. (I do cut my own personal toenails of course!) However, we used to soak them for a short time in Nair, which i believe is like Neet, and we got an
excellent section. [See also Answers 4 and 5.]

The procedure is to process the nails, and after they are embedded treat the paraffin block by putting it in a petri dish containing the Nair. The Nair is put in first and then the block is put on top. We treat the block for 5-10 minutes depending on the size of the nails. We wipe off the block, try cutting it and put it back for further treatment if needed. It is best to cool
the block on iced water after treatment and before cutting and to take the first sections.

Marjorie Hagerty (mhagerty[AT]emc.org)

Answer 3.

I learned a new technique at one of the outstanding workshops at NSH-Albuquerque. Our hospital switched to this method. After grossing, place a representative piece (or ALL if melanoma is indicated) in a cassette and immerse the nail in 5% Tween 80 (Sigma cat#P-4634) for 1-2 hours at least. Overnight won't hurt it. Then remove and process as usual. I find that if you orient the nail to cut it perpendicular to the knife it cuts more easily. Use a charged or polylysine slide (or Elmers glue if it's really likely that it will float).

Andrea Kelly
Albany Medical College
(andrea_kelly[AT]ccgateway.amc.edu)

Answer 4.

There are several methods in Luna's last book "Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts" for softening keratin in nails, etc. Fixation in 10% buffered formalin is necessary to produce crosslinking and thereby prevent keratin from dissolving completely in softening solutions. After fixation and BEFORE processing -- place
specimen in "Neet" or other depilatory cream or permanent wave solution for one to several hours. The key ingredient in these solutions is thioglycollate. * This is best performed under a hood because these products smell really bad and will guarantee an increase in lab traffic by interested personnel wanting to know "What on earth are you doing?" The specimen should bend easily before continuing with next step. Wash the specimen in running tap water for 10 minutes. Dehydrate, clear, and impregnate with paraffin as desired. Processing times will depend on which hoof you are processing -- elephants take a lot longer than goats :-) Get out your nose clip and have fun!

Linda Jenkins
Clemson, SC
(jlinda[AT]ces.clemson.edu)

Answer 5.

We have routinely used "Neet" overnight and had good results. Recently tried "Neet" at 58C (it liquefies) for several hours during the day on a particularly tough nail; it cut beautifully
the next day!  (Neet is a proprietory depilatory. See https://www.hair-removal-products-reviews.com/neet-hair-removal.html.)

Colin Henderson
St. Joseph's Health Centre
London, Ontario, Canada
(colinh[AT]stj.stjosephs.london.on.ca)

Back to Table of Contents

** Paraffin wax: crystals, additives and cutting

Question.

What are the best polymers or other additives for reducing crystal size and improving the cutting propereties of paraffin wax?

Answer.

Paraffin wax is a mixture of (virtually) straight chain hydrocarbons. Note the word "mixture". Unless you go to enormous lengths (of purifying or searching for a fine chemical supplier), you will ALWAYS have a mixture. There is a relationship between hydrocarbon chain length and melting point, but as the waxes are always mixtures, melting points are never exact, either in the compounding or the measuring, but that is another story!

Perhaps more important than the melting point is the "plastic point," but that is virtually ignored by our suppliers. The plastic point occurs about 10 C below the melting point and its meaning should be fairly obvious - try softening a piece of physiotherapy wax in your hands and that should explain all you need to know. The reason the plastic point is important is
related to the sectioning properties of the wax, but we will come to that later! Crystal size is important in the wax surounding the tissue and in the tissue spaces, but not in the tissue per se. Molten wax infiltrates the specimen; the size and shape of crystals will be influenced by the tissues as the molten wax solidifies - i.e. crystalises. So we cannot have "small crystals" infiltrating although smaller crystals will result from solification in denser tissues.

Some of the theory behind this suggests that wax crystalises first as flat "plates," the higher melting point hydrocarbons crystalising first. As successively lower melting points deposit further plate crystals, they pile up upon one another. Distortion due to these dynamic events forces the edges or corners of the most well developed plates to curl and roll. Eventually, that gives rise to needle shaped crystals, which some "experts" consider most ideal for microtomy. All this will be contingent upon the boundaries imposed upon the process by cell and tissue structures. During microtomy, essentially two types of forces are exerted in the cutting process. Flow shearing and point-to-point shearing. Flow shearing is, as you might expect, the smoother and prcedes ahead of the edge of the blade. Point to point shearing has forces seeking the line of least resistence ahead of the blade and these result in a section of uneven thickness - not that you would notice this microscopically.

Imagine the difference between cutting through a jelly and cutting through a beefburger. Now you can imagine where the importance of the plastic point (as opposed to the melting
point) comes in. Additives to paraffin waxes are intended to minimise the point-to-point shearing and improve the plastic flow. The association between the words "plastic" and
"polymers" should now be awakening. Additives to paraffin wax are usually polymers (of known chain length, for they are synthesised exactly), with a major role in "harmonizing the
consistency," in part at least by filling in beteen the wax crystals.

I use pure paraffin wax with no additives, in the belief that proper processing and a SHARP blade are the central features of good microtomy. (I just wish I could practise as well as I can preach!) I have only ever come accross one wax with crystalline structure significantly different from others, and that is Ralwax, which can be helpful when cutting decalcified
specimens, etc.

Russ Allison
Cardiff, Wales (Deceased)

Back to Table of Contents

** Xylene substitutes: what are they?
 
Question.

What are the various liquids sold as substitutes for xylene, and are they really safer and just as good?

Answer.

There are two classes of xylene substitutes: limonenes and aliphatics.

Limonenes are prepared by steam distillation of orange peels. They are terpenoids rather similar to turpentine. They are becoming more expensive and difficult to obtain. Their great
disadvantage is the persistent citrus smell, which many people find intolerable. They are difficult to distil. On the other hand, they are rather minimally toxic, and are easy to dispose of. Various brands are interchangeable.

Aliphatics are synthetic hydrocarbons with about the molecular weight of naphtha. They are odorless, not very toxic, and easily distilled. They are as difficult to dispose of as xylene.

There are at least six brands of aliphatics, and they are NOT interchangeable with each other. They vary consierably in flash point, and they all have different distillation routines.
Richard Allen's Clear-Rite is perhaps the best known of them. Some of the ones offered by ma-and-pa solvent repackagers are quite unsatisfactory.

Bob Richmond, Samurai Pathologist
Knoxville TN
(rsrichmond[AT]aol.com)

Back to Table of Contents

** Test for water in used absolute alcohol

Question.

How can I determine whether used "absolute" alcohol is still OK for the last stage of dehydrating specimens or slides?

Answer.

Some people add anhydrous copper sulphate to the alcohols used for processing tissues. It changes colour (white to blue) in the presence of water, but this does not tell you if there is only a tiny trace of water or enough to make the alcohol immiscible with xylene.

You may be interested in a simple method I developed for this purpose. My job is evaluating histology equipment for the Medical Devices Agency, (an agency of the Department of Health), and I was interested in trying to establish "carry-over" in processing and staining instruments. I started off by adding known dilutions of alcohol, drop by drop, to different amounts of xylene, my basic thinking being that water turns xylene milky, and if one adds enough of the diluted alcohol, the mixture eventually becomes clear again. From this I developed the following method:

A measured 5 ml of xylene (the 5 ml is important) is placed in a 50 ml glass beaker and placed on a black background. Using a 1 ml plastic pasteur/transfer/dropping pipette, add the alcohol for analysis, drop by drop and keep count of the number of drops, until you can just detect a faint turbidity in the xylene. Carry on adding the alcohol to the xylene until the
turbidity just clears, again taking note of how many drops were needed.

Using known dilutions of alcohol, I was able to set up and standardise the method and obtain reproduceable results consistently. The method was not sensitive enough to detect the
water in 99% or 98% alcohol.

  97% = 5 drops to turn xylene milky, 10 drops to clear the mixture
  96% = 4 drops to turn xylene milky, 14 drops to clear the mixture
  95% = 3 drops to turn xylene milky, 34 drops to clear the mixture
  94% = 3 drops to turn xylene milky, 74 drops to clear the mixture
  93% = 3 drops to turn xylene milky, 83 drops to clear the mixture
  92% = 3 drops to turn xylene milky, 98 drops to clear the mixture
  91% = 3 drops to turn xylene milky, 140 drops to clear the mixture
  90% = 3 drops to turn xylene milky, 204 drops to clear the mixture

You would have to initially set up your own range of standard dilutions with the particular alcohol used in your laboratory for the sake of  accuracy. The 1 ml pasteur/transfer/dropping pipettes, they can even be called pastettes, should be held vertically to standardise the size of the drops, and I tried to use the same brand each time.

This is a simple method, and quick to do, although I should think the method would give the Biochemists the shudders. It could help to prolong the life expectancy of the alcohols used
in processors.

Jim Hall
(rmkdh[AT]ucl.ac.uk)

Back to Table of Contents

** Molecular sieves for making anhydrous solvent

Question.

Which type of molecular sieves are used for making anyydrous acetone or alcohol, and how much should I put in the bottle?

Answer.

The molecular sieve to use for acetone is type 3A, mesh 8-12. EM Science Catalog # MX1583L/1 for 500 g or /3 for 2kg.

Before using a molecular sieve, you first have to determine which one to use. Type 3A if for unsaturated hydrocarbons and polar fluids. These include methanol, ethanol, and acetone.
The 3A refers to the size of the molecule it can absorb. In this case, less than 3 angstrom. Molecular sieve 3A has an absorption capacity of 22% by weight.

To dry a liquid, add a slight excess of drying agent.

Next, a little calculation. If the information isn't on the label, call your vendor and retrieve a C of A (certificate of analysis) for the lot of solvent you're using. There should be a specification for water content. This value is the moisture in the bottle upon release. An opened bottle will have higher moisture, depending on how hygroscopic the reagent is. Let's use methanol, which is very hygroscopic, as an example, with the C of A stating that the water content is 1.0%, which equates to 4 ml in a 4 liter bottle. 4 ml of water is equal to 4 g of water. This is 22% of (4 X 100 / 22) = 18.18 g. For excess use 20g of molecular sieves.

Mix thoroughly and allow the liquid to stand. After a few minutes the drying agent settles to the bottom of the container. Separation can be completed by decanting or filtration (suction filtration would work best and fastest ). How often you would dry a solvent out is dependent on application, use, and humidity.

TIP: Depending on application and specifications required, the use of molecular sieves may eliminate to need to purchase expensive super dry reagents.

Rande Kline & Joe Daniels
Technical Services, EM Science
(rkline[AT]emindustries.com)

Back to Table of Contents

 


 

SECTIONING, SLIDE ADHESIVES, MOUNTING

 

** Sections coming off slides. Which adhesive?

Question.

Here is my problem: tissue sections not adhering to the slides.
Any hints on solving this problem?

Answers.

[ Textbooks of microtechnique contain recipes for various adhesives: chrome alum-gelatin, Mayer's albumen and starch paste are traditional. More recent methods include giving the glass surface a positive charge by coating with polylysine or reaction with 3-aminopropyltriethoxysilane (APES or TES) to make "silanized" slides. See also the FAQ item on how to
prepare silanized slides. There is also an FAQ item about polylysine.

Here are 7 hints from individuals. No. 3 is pertinent to the use of any adhesive or none at all. ]

1. We ran into the problem of tissues falling off the slides after about 5 hours of immunohistochemical processing. We seemed to have solved it with Super Frost Plus slides that have some sort of charge on them. 
Pre-prepared silanized slides are commercially available with a variety of trade names that often include "plus".

P. Emry
(emry[AT]u.washington.edu)


2. We go to the expense of using charged slides for everything we do (Plus slides) and nothing really ever floats. If you don't want to go to that expense, we used to use chrome alum-gelatin with fairly good results and only an occasional problem. I personally don't like having chemicals in the waterbath. An exception would be immunos and some frozens for which I would recommend using "Plus" slides regardless.

Xylene in paraffin as a cause. I had an interesting thing happen to me once. I worked Saturdays for a while, training a girl at another lab in Histotechnique. The first Saturday we cut, most sections floated to varying degrees, even things like tonsil. The tonsil cut very nicely and seemed well processed. Well, I was supposed to be the person in the know and I was
stumped. Took me a while but I finally figured out that they didn't change the processors very often and there was lots of xylene in the paraffin, and I mean lots. Apparently this was the problem, because after changing everything and rotating on a regular basis the problem went away. I just thought I would throw that story in - because this experienced histotech didn't realize that excess xylene in the paraffin could cause problems with adhesion of sections.

Marjorie A. Hagerty
(mhagerty[AT]emc.org)

3. Here is another possible contribution to the section loss. After picking up the ribbon on the waterbath do you purposefully pull out the water from between the section and the slide? ....you know, using a lap cloth (or whatever absorbant material you keep around) to touch the edge of the paraffin ribbon and soak up the water from under the ribbon. If the edges of the ribbon adhere to the slide but water remains between the section and the slide, when drying occurs, it is possible that not ALL of the water has evaporated from that space. Obviously, if a little water still separates the specimen from the slide (no matter what adhesive material is present), then the less than complete specimen attachment may not be strong enough to make it through the (even gentle) turbulence of the staining process.

This negative condition is most often seen when a ribbon is picked up and then the slide is immediately placed flat, horizontally, on th edge of the waterbath. It can also occur, though far less fequently, when the slide is immediately placed vertically against the waterbath or into a slide rack. The vertical positioning, however, does increase the draining of the water as long as the bottom of the ribbon has not fully attached to the slide creating a dam of sorts.

Anyway, that's just one more variable for you to consider before perhaps investing in something which may offer no greater adhesive advantage than what you are currently using.

Nancy Klemme
(nancy.klemme[AT]sakuraus.com)

4. Nancy is absolutely correct! Even with super adhesives or charged slides, you're liable to lose sections if the interface of the section and the microscope slide's glass is not water-free. This water is also a cause for "nuclear bubbling" artifact.

Ken Urban
Surgipath Medical Industries, Inc.
Richmond Illinois
(surgamy[AT]mc.net)

5. I bought 6 slide racks, the ones where slides stand on their ends, each holding 50 slides (Solmedia in the UK). These I keep for coating only. I've also got a couple of deep staining pots, again for coating only. I buy poly-L-lysine from Sigma or make my own gelatin-chrome-alum. Load the racks with slides, clean, I don't trust the manufacturers, wash thoroughly and coat with the coating of choice, dry and box. Couldn't be easier, I make enough in 2 days that will last months. Why be ripped off by the supply houses when for a few £/$ you can do it yourself.

Ian Montgomery
(I.Montgomery[AT]bio.gla.ac.uk)

6. Polylysine has free amine groups that form positively charged ions in water that's less alkaline than about pH 9. Slides are smeared with an aqueous solution of this basic amino acid polymer and then air-dried. This confers a positive charge to the slide's surface when immersed in water. Amino acid anions (which predominate in a section of a typical vertebrate animal tissue) are attracted to the polymer that covers the glass. It whether you use poly-L-lysine or poly-D-lysine or poly-DL-lysine, because the stereochemical form of the amino acid does not affect its protonation. Buy the cheapest.

Positively charged slides can also be made in the reaction of an aminoalkylsilane with glass, in the presence of traces of water. It is easy to produce hundreds of "silanized slides in an hour. Alternatively, you can buy the silanized slides, which amounts to paying a company to do this simple job.

John A. Kiernan,
London, Canada
(kiernan[AT]uwo.ca)

7. I was satisfied with poly-L-lysine until I tried Superfrost Plus slides. I went from occasionally losing tissue to never losing it...so I vote for Superfrost Plus.

Mary Ross
(ross.8[AT]osu.edu)
Patricia Emry
(emry[AT]u.washington.edu)

Back to Table of Contents

** Apathy's mounting medium and variants

Question.

Where can I find the recipe to make von Apathy's mounting
medium? Is there more than one way to make it?

Answer 1.

Von Apathy's medium is simple to make and lasts well so it is very straightforward to make it yourself.

Von Apathy's Gum Syrup medium, RI 1.52:

Dissolve 50 grm gum arabic (gum acacia) and 50 grm cane sugar in 50 ml of distilled water with frequent shaking in a 60 degree water bath. Add 50mg of thymol (or 15mg Merthiolate) as a preservative. If too thick for your application increase the amount of distilled water.

While warm put in a vacuum chamber to remove air bubbles.

To prevent "bleeding" of metachromatic staining of amyloid by methyl or crystal violet, modify Apathy's medium as follows. Add 30 to 50 grm of potassium acetate
or 10 grm of sodium chloride, and enough water to bring the volume from 50 ml to 100 ml when everything is dissolved.

This mounting medium sets hard and there is no need to seal the coverglass.

Richard Powell
Darwin, Australia
(richard.powell[AT]nt.gov.au)

Answer 2.

You will find the recipe, only it is called Apathy's gum syrup, in Histopathologic Technic and Practical Histochemistry, edited by RD Lillie & HM Fullmer (3rd edition, 1976, page 101). The recipe given is Lillie and Ashburn's modification. Ref: Arch Pathol 36:432 1943. It was Highman who modified the medium by adding potassium acetate and sodium chloride. Ref: Arch Pathol 41: 559 (1946).

RAB Drury and EA Wallington also mention Highman's variant in the excellent book, "Carleton's Histological Technique," 4th ed. London: Oxford University Press, 1967.

John Kiernan, London, Canada
(kiernan[AT]uwo.ca)
Ian Montgomery, Glasgow, Scotland
(i.montgomery[AT]bio.gla.ac.uk)

Back to Table of Contents

** Silanized (APES or TES or positively charged) slides

Question.

How do I prepare charged or silanized slides in the lab, and is it OK to use metal slide racks?

Answer 1.

Silanized slides have a permanent positive charge associated with the glass surface. This attracts negative ions in the section (things like sulfate of cartilage and carboxylate of protein). You can buy silanized slides; they have a variety of trade names and are more expensive than ordinary slides.

It is easy to make your own positively charged slides using APES (also abbreviated to TES). You can buy 3-aminopropyltriethoxysilane from Sigma (St Louis, MO) or from Strem Chemicals (Newburyport, MA) or from Gelest (Tullytown, PA). Keep it in the fridge; let it warm to room temperature before opening the bottle. The solution in acetone deteriorates after one day.

  1. Wash slides in detergent for 30 minutes.
  2. Wash slides in running tap water for 30 minutes.
  3. Wash slides in distilled water, 2 X 5 minutes.
  4. Wash slides in 95% alcohol 2 X 5 minutes.
  5. Air dry for 10 minutes.
  6. Immerse slides in a freshly prepared 2% solution of 3-aminopropyltriethoxysilane in acetone for 5 seconds.
  7. Shake off excess liquid and wash briefly in distilled water, twice.
  8. Dry overnight at 42C and store at room temperature.

300 ml of silane solution is sufficient to do 200 slides. Treated slides can be kept indefinitely.

James Lowe
University of Nottingham
(James.Lowe[AT]nottingham.ac.uk)
http://www.ccc.nottingham.ac.uk/~mpzjlowe/protocols/silslid.html

Answer 2.

As far as I know, the notion that you must do TES treatment in glass slide trays is another urban myth! We coat thousands of slides annually in metal racks with nary a problem.

Bryan Hewlett (CMH)
(hewlett[AT]exchange1.cmh.on.ca)

Back to Table of Contents

** Polishing undecalcified bone sections.

Question.

Which kinds of grit should I use to polish away the scratches from the surface of a section of plastic-embedded undecalcified bone? Any other advice would also be appreciated.

Answer.

Try using a series of fine grit grinding papers before going to the polishing cloth with 1 ĉm alumina slurry. Remove scratches progressively, by going to a 320 or 400 grit, then 600 grit.
Grind with a figure 8 motion, and rinse well between grits. Then go to your 1 ĉm alumina polish, figure 8 motion, and use Buehler microcloth (velvet type surface) that comes in sticky back, can stick to a plastic surface, or whatever to prevent slippage, polishing takes only a few (2 or 3) minutes. Examine under a magnifying glass for scratches. The first grits for grinding depend on the grit size of your diamond cutoff blade. There is a way to read the codes for this grit: if you have a 320 grit size of diamond, then go to 400 grit (Norton waterproof paper, Tufback Durite) paper first.

Be sure to flow water across tilted grinding surface, to wash bone "dust" and plastic away. I like grinding paper taped to a thick plexiglass rectangle, with one end slightly elevated with a rubber handled hammer. It's cheap! The 1 ĉm slurry (small amount) should be put on a slightly wet polishing cloth; that way it will polish more easily and quickly. For a mirror-smooth surface, go to 0.1 ĉm alumina slurry after the 1 ĉm.  I have tried progressive alumina slurries, 3 ĉm then to 1 ĉm, but it was a waste of time, 1 ĉm worked just as well. Polishing away scratches after 600 grit paper worked well. Finer grits (800, 1000, 1200) didn't help that much and were expensive.

Equivalents are:
   400 grit = 22 ĉm
   600 grit = 14 ĉm
   800 grit = 10 ĉm
   1000 grit = 5 ĉm

Whatever you do, protect your joints from the stress of grinding and polishing. Use holders. The ergonomics of polishing will eventually take its toll, damaging your finger joints -   want a photo? Buy an automatic grinder and polisher if at all possible. This was the best investment we ever made, but too late!

Gayle Callis
(
gayle.callis[AT]bresnan.net)

Back to Table of Contents

** Polylysine-coated slides

Questions.

For how long can you store a solution of poly-L-lysine used as a section adhesive for slides?
For how long can you store the coated slides?
Do you get autofluorescence?
Do you have to use poly-L-lysine, or will the cheaper poly-DL-lysine work equally well?

Answer 1.

I use a 1:10 dilution in PBS of Sigma's stock poly-L-lysine solution (P-8920). Slides sit in the solution for 4 hours (or more if you choose/it is more convenient) and air dry overnight.
This has worked for us without ever a section lost. The poly-L-lysine solution (undiluted from Sigma) says it expired in 1996, but it still worked in summer '97. I have never noticed
any autofluorescence.

I have switched to Superfrost Plus slides; when counting in time to put slides in racks to dip and the time to rebox them, it is more cost-effective for us to buy the superfrost plus.

Noelle Patterson, M.S.
NNMC/NMRI/ICBP, Bethesda, MD 20889
(pattersonn[AT]nmripo.nmri.nnmc.navy.mil)

Answer 2.

The type of polylysine does not matter, so get the cheapest, which is usually the mixed (DL) enantiomers rather than the pure L- form. The reagent and the slides should keep for ever if they don't get infected with micro-organisms or contaminated with dust.

For a simple way to prepare polylysine-coated slides, see Thibodeau, T. R., Shah, I. A., Mukherjee, R. & Hosking, M. B. 1997. Economical spray-coating of histologic slides with
poly-L-lysine. Journal of Histotechnology 20(4): 369-370.

They stated that it was economical and quick to spray polylysine solution on one side of the slides from a simple plastic spray bottle. Results were no worse than dipping, which was more trouble. They used a 1:10 dilution of PLL solution but did not state the concentration, molecular weight or source.

John Kiernan,
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Wrinkles in plastic sections

Question.

How can I prevent wrinkles in sections (0.5 to 2 micrometers) of plastic-embedded tissue stained for light microscopy?

Answer.

The wrinkles form when mounted plastic sections are stained in a hot aqueous dye solution. Chandler & Schoenwolf (1983) found that the wrinkles did not form if sections were dried down onto acid-washed slides, overnight, at 76C. They thought acid-washing might improve the glass surface in some way. The minimum drying time was 6 hours. The temperature was also important. Variation was not fully investigated, but neither 60C nor 90C was efffective in preventing wrinkles.

Reference: 
Chandler, NB & Schoenwolf, GC (1983) Wrinkle-free
plastic sections for light microscopy. Stain Technology 58: 238-240.

John A. Kiernan,
Department of Anatomy & Cell Biology,
The University of Western Ontario,
London, CANADA N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Wrinkles in paraffin sections containing cartilage

Question.

Does anyone have a reliable procedure to consistently avoid wrinkles with cartilage in paraffin sections of trachea (human, mouse, rat)?

Three tips and wrinkles follow.

Answer 1.

This is what works for me most of the time. I only cut human cartilage/trachea so I don't know if the mouse/rat needs to be treated differently. I keep my water bath hot, 50 degrees C,
which may be too hot for whatever paraffin you are using. I use plain paraplast. It is important that the section be thin and that the disposable knife edge is new. I never take the section from the same knife area that I used to shave into the block. First, I shave into the block to the desired depth. Then I soak the block on an ice tray that has water added. Next, I take a section from the first ribbon off the block. I let it float on the waterbath until it looks very smooth, just a matter of a quarter of a minute or maybe a little longer. This usually results in no wrinkles microscopically.

Marjorie A. Hagerty
(mhagerty[AT]emc.org)

Answer 2.

Try picking up the section on the slide (from the waterbath) and then immediately holding the slide for a few seconds on a hot plate. This has to be monitored, because too much heat on such a wet section may cause the rest of the tissue to "explode"

Louise Taylor
(179LOU[AT]chiron.wits.ac.za)

Answer 3.

I have found 1, 2, or 3 drops of the new thick Joy in the waterbath has helped with wrinkles in some of my tissue cutting experiments. Start with just one drop then add more slowly. If
you get too much in it, you spend your time chasing the section around the waterbath.  [ Joy is a liquid dishwashing detergent sold in N. America. ]

Trisha Emry
(emry[AT]u.washington.edu)

Back to Table of Contents


**  Electron microscopy and semithins after paraffin embedding

Question.

I have some precious formaldehyde-fixed, paraffin embedded (FFPE) specimenss that were embedded in agar (to orient the tissue easily) prior to processing into wax.
There is now a need to take the tissue backwards, out of wax, and process it for semithin plastic (glycol methacrylate, GMA, JB-4) sectioning.

Is that possible? Will the tissue having previously been embedded in wax cause problems for the monomer infiltration or the plastic curing? I know that taking tissue back through xylenes and alcohols is supposed to remove all the wax, but does it really?


Answer.

We regularly re-embed paraffin-embedded tissue in epoxy resin for electron microscopy. We are using very small specimens - kidney, liver, 1mm core biopsies.

We run them through xylene 4 X 30 minutes at 60C, 100% ethanol 3 X 10 min, 95%, 70%, 50% ethanol 5 min each, then distilled water, then buffer. Then we go on to processing for resin embedding. The wax is gone and does not interfere with the plastic.
The agar will not affect or be affected by plastic embedding. We use agar to embed cilia for EM with good results.

You might want to do some tests to optimize the timings for your samples.

Tim Morken
Supervisor, Electron Microscopy/Neuromuscular Special Studies
Department of Pathology
UC San Francisco Medical Center
timothy.morken[AT]ucsf.edu

Back to Table of Contents



** Separation of  paraffin ribbon from sections

Question  

When I place my sections on the waterbath the paraffin pulls away from the edge of the tissue. I have tried at different temperatures . The tissue itself is fine so I do not believe the separation is due to processing. I use the same paraffin (Fisher Histoplast IM)
for both processing and embedding.  I was considering melting a block or two down and embedding them in another kind of paraffin. Suggestions?

Answer
.

One cause of tissue separation is the difference in temperature between tissue and embedding wax. Try taking specimens from molten wax and embedding directly in the wax-filled mold.

Tony Henwood
Pathology Department, The Children's Hospital at Westmead
Westmead NSW 2145, AUSTRALIA

(tony.henwood[AT]health.nsw.gov.au)

Back to Table of Contents


** Thick paraffin sections

Question.

I need a method for cutting near perfect 50 micron sections of paraffin processed tissue. They are not only difficult to cut, but will not stay on the slides! Please advise.

Answer.

We use the plus slides and put about 25 drops of Elmer's school glue in the waterbath. This combination works VERY well for us. [
Elmer's school glue is a white polyvinyl acetate product that can be removed from unwanted places with warm water. ]

Sarah Ann Christo
(schristo[AT]cvm.tamu.edu)

Back to Table of Contents

** Sectioning plastic-embedded specimens

Question.

How do you cut flat sections of materials embedded in poly(glycol methacrylate) (GMA) and other resins, for light microscopy?

Answer 1.

I always use glass knives (standard or Ralph type) but using a tungsten carbide blade should not be a problem.

Cutting speed is (in my experience) critical, and have found that very slow (almost to the point of stop!) will provide a crease free section. This is where patience is a virtue: tedious but worth the wait.

The section may tend to "roll" but this is not a problem, in fact I find this an advantage. Simply remove the section from the blade and place onto a warm surface (the palm of your hand will suffice) and watch in amazement as the section unfolds (a bit like those fortune fish from many years ago). Then drop the section onto warm distilled water to remove any further folding. It's a bit laborious, but usually best to handle only one section at a time. I hope this is of some help. There are also a few "tricks" with the staining!

Terry Hacker
MRC Harwell, Oxfordshire, England
(T.Hacker[AT]har.mrc.ac.uk)

Answer 2.

I have cut a lot of plastic, and here's what I do:


  (1) Cut at about 6-7 microns.
  (2) Soak, soak, soak! (about 2-3 hours depending on what kind of polymer).
  (3) Use positively charged slides, with Elmer's glue in the waterbath.
  (4) Use one of the newer, heavier microtomes (we have the Leica 2035s).

Lori Miller
Flagstaff, AZ
(lmiller[AT]wlgore.com)

Answer 3.

Plastic sections do not ribbon, unless you put a dab of rubber cement on the the top and bottom of the block, but usually we pick them up one section at a time.

Curling is very common. What I do is start the sectioning but do not finish; keep it attached to the block, then you can use a brush or fine forceps and unroll it, pulling at a diagonal.
Leaving it attached lets you pull without completely pulling the section off the block. When you have it fairly open and flat, complete the sectioning stroke thereby releasing the section.
I used to slide the MMA section onto a spatula, keeping it wet with alcohol, and then slide it off the spatula onto a slide onto a hot plate. Keep dropping alcohol onto to the section and
it should flatten out.

GMA is much easier to pick off the block. Do the same thing but keep everything very dry, pick up the section with a fine forceps and drop it onto a water bath and it will flatten out.
Scoop onto a slide from the water.

Patsy Ruegg
(patsy.ruegg[AT]uchsc.edu)

Back to Table of Contents

** Iodine for removing mercury deposits

[ It is necessary to remove mercury deposits from specimens fixed in B5 or other fixatives that contain mercuric chloride. Textbooks recommend either including a solution of iodine (0.5 to 1%) in 70% alcohol in the series of solvents for dehydrating before embedding, or treating the sections, after hydration, with iodine followed by sodium thiosulfate. ]

Question.

I am interested in the possibility adding of iodine to the first xylene, in a staining machine. What is the percentage or recipe for that solution, and will it corrode metal parts?

Answer 1.

We use a 0.5% solution of iodine in xylene for 5 minutes. We have been doing this regularly for about 6 months and have only had a problem with a couple of lymph nodes, in that the mercuric crystals were not completely removed. We had to give additional treatment off the machine.

We have a Leica stainer, and everything inside looks like stainless steel. It seems to be unaffected by the iodine thus far. We do always use the same staining dish and lid for the
iodine/xylene because the plastic is stained.

Marg Hagerty
(mhagerty[AT]emc.org)

Answer 2.

At my previous lab we used 1 percent iodine in the first xylene to clear out the mercury crystals. That was using glass jars and metal racks in a manual method. There were no problems with corrosion of the racks.

Tim Morken
San Francisco, CA
(
timothy.morken[AT]ucsf.edu
)

Back to Table of Contents

** Labeling slides

Question.

Have you any suggestions for labels that could be used during the staining process that would still be legible and won't come off in xylene?

Answer.

(a) For slides with a frosted end: Use an ordinary (graphite) pencil. After coverslipping cover the pencil with a thin layer of clear nail polish or diluted (1:5 in toluene or similar) mounting medium.

(b) For plain slides, use a diamond-tip pencil directly on the glass. This is very permanent, but it's more trouble than frosted slides - something of an art, especially if you need to write quite a lot on each slide.

With both methods there's a risk of getting BITS (of either graphite powder or ground glass) on the sections. Graphite is worse, because it's black. It's therfore a good idea to put a
piece of paper over most of the slide for protection while you're writing on the end.

John A. Kiernan,  Anatomy & Cell Biology,
The University of Western Ontario, LONDON, Canada
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Sectioning plant material: some hints.

Questions.

Does anyone know how to make nice sections of plant material? I tried to make paraffin sections but it seemed that the thick cellulose walls of the cells were preventing the penetration of paraffin into the tissue.

Do I have use longer impregnation times for paraffin?

Is it easier to make cryostat sections of plant material?

Do I have to make the cellulose softer? And how do I do that?

Are there any references that pertain specifically to botanical microtechnique?

Answer 1.

I had a large project of plants several years ago. We processed and cut everything from jalapeno peppers to magnolia leaves. At that time I tried to find a book on plant histotechnique and the only one available was Botanical Microtechnique and Cytochemistry by GP Berlyn and JP Miksche. The Iowa State University Press, Ames, IA, 1976. ISBN #8138-0220-2.
We adapted our animal histology techniques to the plants and found that the most important phase was the paraffin. We used Paraplast, but the important part was three changes of paraffin, 1 hour in each. Vacuum was not used on the first paraffin, but was used on the final two.

You must also know that processing will decolorize the chlorophyll in the tissues.

For staining, I would suggest trying a safranin O - light green - alum hematoxylin sequence. It works the best for plant cells.

Cheryl Crowder
(crowder[AT]vt8200.vetmed.lsu.edu)

Answer 2.

Here are a few general references for plant microtechnique. The methods are similar in principle to those for animal tissues, but allowance must be made for the high water content and
fragility of plant specimens.

My own experience is very limited, but it fully supports the advice of Berlyn & Miksche. Cut your pieces with a VERY sharp razor blade, using a sawing motion, and do not expect decent sections from anywhere near the cut surfaces of the specimen.

Dehydrate as gently as possible, to avoid sudden collapse of the tissue, which distorts all the cells. There are three ways to dehydrate a plant specimen gently:

1. By immersing in 10% aqueous glycerol, which mixes with the water in the tissue, and waiting for the water (90% of the volume) to evaporate. This takes many days. The glycerol is then gradually displaced by alcohol, then xylene, then paraffin. Although slow, this procedure is not unduly labor-intensive.

2. By using a long series of graded water-alcohol mixtures, from about 15% up to 100% alcohol. This keeps someone busy for the best part of a day, and it is easy to forget the plant specimens if you are doing other things.

3. Acid-catalyzed chemical dehydration with 2,2-dimethoxypropane is a single step, usually less than one hour. It is nevertheless "gentle" to the tissue, though perhaps a bit more traumatic than the glycerol evaporation method.

Special methods are needed for wood and other hard plant materials. They are comparable to, but different from, methods used in processing of bone. See references below, especially to books by  Berlyn & Miksche (1973) and Ruzin (1999). 

Nostalgic note. Anyone who studied Biology in Britain or the Commonwealth from the 1940s to the early '70s (maybe even more recently?) will remember the practical component of the A-level (or HSC) public examination that came at the end of the upper-sixth form (like Grade 12 in N. America). This always included sectioning a piece of plant by hand (with a cut-throat razor; no embedding and no microtome). The free-floating sections then had to be stained, mounted, examined, and drawn with a pencil. These thick (?20-40
μm) sections showed the plant anatomy pretty well under a X10 objective. Thinner paraffin sections provide better detail with a X40 objective, but only if the general tissue architecture is intact. The structural preservation seems to depend heavily on the way the specimen is processed into wax.

References.

Berlyn,GP; Miksche,JP (1976): Botanical Microtechnique and Cytochemistry. Iowa State University Press, Ames, Iowa. 336 pages.
   Has chapters on fixation, processing, wax & plastic embedding, staining (methods with hemalum, safranine, light green etc; detailed accounts of 8 methods); Histochemistry.

Clark,G (Ed.) (1973): Staining Procedures used by the Biological Stain Commission. 3rd ed. Williams & Wilkins, Baltimore. 418 pages. 
Clark,G (Ed.) (1981): Staining Procedures.
4th ed. Williams & Wilkins, Baltimore. 512 pages.
   These, the last two editions of the Biological Stain Commission's manual, are different books with the same editor but largely different crews of contributors. Both are valuable sources of well documented practical information, with lengthy bibliographies.  

Jensen, WA (1962): Botanical Histochemistry. Freeman, San Francisco.

Kiernan, JA (2015): Histological and Histochemical Methods. Theory and Practice. 5th ed. Banbury, UK: Scion Publishing. 571 pages.

Ruzin, SE (1999): Plant Microtechnique and Microscopy. New York: Oxford University Press. 322 large pages.

Vaughn,KC (Ed.) (1987): CRC Handbook of Plant Cytochemistry. 2 vols. CRC Press, Boca Raton, Florida. 176 & 184 pages.
   Multi-author, 2 vols. Oxidative & hydrolytic enzymes in Vol 1. Carbohydrates, lectins, immunohistochemistry and methods for, Na, Ca, K in Vol 2.

John Kiernan, London, Canada
(kiernan[AT]uwo.ca)

Back to Table of Contents

 


 

STAINING METHODS,  HISTOCHEMISTRY

 

** Making aldehyde-fuchsine

Question.

Paraldehyde is a controlled substance, not that easy to obtain for laboratory use, and it also has a short shelf life. Is there a way to make the aldehyde-fuchsine stain without using
paraldehyde?

Answer.

When aldehyde fuchsine is made in the traditional way, the paraldehyde decomposes in the presence of acid, yielding acetaldehyde. This reacts with pararosaniline to form a new dye, which is the active component of the stain. It is therefore possible to use acetaldehyde (obtainable from regular chemical suppliers) instead of paraldehyde.

The late Peggy Wenk, bless her heart, commented on this in the Journal of Histotechnology, vol 10, #4 (December 1996): Acetaldehyde as a substitute for paraldehyde.

2.5 ml acetaldehyde is used in place of 1.5 ml paraldehyde. The working solution must be refrigerated. It will stain hepatitis B for 3 - 4 weeks, but is good for elastin for several months.

Acetaldehyde costs about $30 for 100 ml and is stable in a refrigerator for about 2 years. (Paraldehyde is stable for only a few months after opening, and is pricey due to handling/admin
fees.) You need to be aware that acetaldehyde is a flammable liquid that boils at 21C. The bottle must be cold when you open it!

Having struggled trying to get paraldehyde, this substitution has made aldehyde-fuchsine staining feasible in a research laboratory.

Gayle Callis
(
gayle.callis[AT]bresnan.net)

Back to Table of Contents

** Phosphatases in decalcified, embedded tissue.

Question.

Can acid phosphatase activity still be demonstrated in formalin fixed, decalcified, paraffin embedded bone sections?

Answer 1.

Have a go, I used to stain for acid phosphatase in 1-10 
μ
sections of demineralized, glutaraldehyde/osmium-fixed epoxy-embedded specimens with no bother. The method was nothing special, just a standard napthol AS-BI phosphate/diazotised pararosaniline technique.

While we're at it, how about alkaline phosphatase in ethanol-fixed, methacrylate embedded sections? Try: McGadey,J. 1970. Histochemie 23. 180-184. Tetrazolium method for
non-specific alkaline phosphatase. This is an excellent method; it has never let me down in any application.

Ian Montgomery
(I.Montgomery[AT]bio.gla.ac.uk)

Answer 2.

I routinely do acid phosphatase staining on formic acid-decalcified GMA-embedded bones. Alkaline phosphatase can also be demonstrated in the GMA and is retained by the alcohol
fixation. The problem that I have found with trying to both from the same block is that the acid phosphatase stains much better with formalin fixation and the alkaline phosphatase stains
better with alcohol fixation.

I have had good results with acid phosphatase using formic acid decalcification and paraffin embedding of rodent skull. 

An excellent article is C. Liu et al. "Simultaneous demonstration of bone alkaline and acid phosphatase activities in plastic embedded sections and differential inhibition of the activities." Histochemistry 86:559-565, 1987,

Martha Strachan
Skeletech, Inc., Kirkland, WA
(mstrachan[AT]skeletech.com)

Back to Table of Contents

** Congo red for amyloid

Question.

Why does alkaline Congo red stain amyloid feebly in sections of some specimens but not others?

Answer.

Here is one possibility. In developing the alkaline Congo red method, Dr. Holde Puchlter noticed decreased staining with prolonged fixation in formalin or NBF. This decrease even
applied to unstained sections stored under conditions where formaldehyde was present in the ambient air.

Susan Meloan
Medical College of Georgia, Augusta
(smeloan[AT]mail.mcg.edu)

Back to Table of Contents

** Cartilage staining with safranine

Question.

How do you stain cartilage with safranine?

Answer.

The Safranin O method for Cartilage goes like this;

1. Dewax section and take to water.
2. Stain nuclei with a suitable iron haematoxylin.
3. Blue in running tapwater.
4. Rinse in distilled water.
5. Stain with 1% light green diluted 1 in 5 with distilled water, for 3 minutes.
6. Rinse in 1% acetic acid.
7. Stain with 0.1% Saffranin O, for 4 - 6 minutes.
8. Rinse in 1% acetic acid and check under microscope. Any overstaining with Safranin can be modified by re-applying the light green solution briefly, and vice versa.
9. Dehydrate with alcohol,clear and mount.

(Modified from the method in R.D. Lillie's "Histopathological Technic and Practical Histochemistry")

John Difford
London, England
(adford[AT]compuserve.com)

Back to Table of Contents

** Stain for Chlamydia (Castaneda's method)

Question.

How do you carry out the Castaneda stain for Chlamydia?

Answer.

Castaneda's stain for elementary bodies and Rikettsiae (1930)

Castaneda's staining solution

   Solution A
      Potassium dihydrogen phosphate, anhydrous 1 g
      Disodium hydrogen phosphate 25 g
      Distilled water 1000 ml
      Formalin (37-40% formaldehyde) 1 ml
Dissolve the potassium dihydrogen phosphate in 100 ml distilled water and the disodium hydrogen phosphate in 900 ml distilled water. Mix the two solutions to give a buffer pH 7.5, and add formaldehyde as a preservative.

   Solution B
      Methylene blue 1 g
      Methanol 100 ml

   Staining solution
      Solution A 20 ml
      Solution B 0.15 ml
      Formalin 1 ml

Safranine-acetic acid

   Safranine (0.2% aqueous solution)    1 part
   Acetic acid (0.1% aqueous solution)  3 parts

Procedure.

  1.  Prepare films from infected tissue and dry in air
  2.  Apply the stain for 3 min.
  3.  Drain, do not wash
  4.  Counterstain for a 1-2 seconds in safranine-acetic acid
  5.  Wash in running water, blot dry.


Rickettsiae, elementary bodies of psittacosis: blue. Cell nuclei and cytoplasm: red.

References: "Biological stains and staining methods." BDH leaflet, 1966.

Several modifications of Castaneda's original technique are given in: Langeron, M. (1916-1949) Précis de Microscopie. Paris: Masson et Cie. [Several editions, including reprints published since 2010. See https://www.abebooks.fr/rechercher-livre/titre/precis-de-microscopie/auteur/langeron/.]

Yvan Lindekens
(yvan.lindekens[AT]rug.ac.be)

Back to Table of Contents

** Which staining method for copper is best?

Question.

Which histochemical staining method is best for copper in human or animal tissues? The choice seems to be between rubeanic acid (not in catalogs) and some impossibly long name that ends in "rhodanine."

Answers.

Notes. This question to the HistoNet listserver elicited many replies. Most favored the "rhodanine" reagent over "rubeanic acid".  Nomenclature can be confusing! 

Rhodanine is a quite different substance from the histochemical reagent used to detect Cu, which is p-dimethylaminobenzylidenerhodanine. In any chemical catalog, p-dimethyl-  is indexed under the letter D, not P.  Don't confuse rhodaNine with rhodaMine!  Neither rhodanine nor any dye with rhodamine as part of its name can be used for histochemical localization of copper in tissues. 

The old name rubeanic acid has unfortunately been perpetuated in histochemistry books, including the comprehensive works of Pearse and Lillie. It is an old name for dithiooxamide, which is the name to seek in a catalog. 

A few general references for copper histochemistry are added at the end of this FAQ item, as Answer 2.

Answer 1.

Rubeanic acid is
H2NCSCSNH2 and is listed in catalogs as Dithiooxamide (by Aldrich, Sigma and other vendors).
I prefer the "rhodanine" method for the demonstration of Copper, which follows.

Fixation: 10% neutral buffered formalin.

Embedding: Paraffin sections cut at 6 microns

Solutions:

   Distilled water, preferably deionized, should be used in all solutions and rinses.

   Rhodanine saturated solution (stock).
      p-dimethylaminobenzylidene-rhodanine 0.2 g.
      Absolute ethanol 100 ml

   Rhodanine solution (working).
      Rhodanine saturated solution (stock) 6 ml
      Distilled water 94 ml

   Note: Chemically clean glassware is necessary. Shake stock solution before measuring and mixing with water, and shake the working solution before pouring it onto the slides in Step 2 of the technique.

   Diluted Mayer's hematoxylin.
      Mayer's hematoxylin (= Mayer's hemalum)  50 ml
      Distilled water  50 ml

   0.5% aqueous sodium borate (borax)

Technique:

 1.  Dewax and hydrate slides (to distilled water).
 2.  Incubate slides in rhodanine working solution at 37C for 18 hours.
 3.  Wash slides well in several changes of distilled water.
 4.  Stain slides in diluted Mayer's hematoxylin for 10 minutes.
 5.  Rinse slides with distilled water.
 6.  Quickly rinse slides in 0.5% sodium borate.
 7.  Rinse slides with distilled water.
 8.  Dehydrate slides through 95% to absolute ethanol, clear, and coverslip with a synthetic resinous mountant.

Results:

   Copper - orange/red.
   Tissue elements - light blue.

Eric C. Kellar
University of Pittsburgh Medical Center
(kellarec[AT]msx.upmc.edu)

Answer 2.

A few references for copper histochemistry, with comments.

Irons,RD; Schenk,EA; Lee,CK (1977): Cytochemical methods for copper. Archives of Pathology and Laboratory Medicine 101, 298-301.
    Cytochem methods for copper. Critical comparison of dithiooxamide, p-diaminobenzylidene-rhodanine, diethylthiocarbamate.  

Nemolato S, Serra S, Saccani S, Faa G (2007) Deparaffination time: a crucial point in histochemical detection of tissue copper. Eur. J. Histochem. 53: 175-178.

Pearse, AGE (1985) Histochemistry, Theoretical and Applied, 4th ed. Vol. 2.   Edinburgh: Churchill Livingstone.
    Metal histochemistry is extensively reviewed in Chapter 20.

Soto M, Cajaraville MP, Angulo E, Marigomez I (1996) Autometallographic localization of protein-bound copper and zinc in the common winkle, Littorina littorea: a light microscopical study. Histochem. J. 28: 689-701.

Szerdahelyi,P; Kasa,P (1986): A highly sensitive method for the
histochemical demonstration of copper in normal rat tissues. Histochemistry 85, 349-352.
    Highly sensitive method for Cu histochemistry.  Magnesium-dithizone, followed by silver intensification.

Szerdahelyi,P; Kasa,P (1986): Histochemical demonstration of copper in normal rat brain and spinal cord. Histochemistry 85, 341-347.
    Histochemical demonstration of Cu in normal brain, spinal cord.

John A. Kiernan
London, Canada
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Diastase (amylase) control for glycogen

Question.

Which is better as a control for glycogen staining: alpha-amylase or human saliva?

Answer.

The bought enzyme (10 mg/ml, in water) takes about 10 minutes to remove the stainable glycogen from a section of liver. The enzyme is not very expensive.

Saliva is free, and it takes about 30 minutes, but some people don't enjoy spitting, or even dribbling, onto their slides. A theoretical disadvantage of spit is that it contains plenty of
digestive enzymes additional to amylase (= diastase), notably ribonuclease and various proteases. However, these are unlikely to remove substances with the same staining properties as
glycogen.

John A. Kiernan
London, Canada
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Evans blue, trypan blue and eosin as tracers.

Question.

Can Evans blue be used as a tissue dye, and will it safely wash out of the tissue during routine paraffin processing? The object is to trace a catheter leakage then have the dye wash out
of the tissue during processing. Would eosin be OK for the same purpose?

Answer 1.

Evans blue is an anionic dye with large molecules, closely related to trypan blue. It was formerly used (? still is in some places) to measure blood volume, because it binds to serum
proteins and stays in the circulation for a few hours. When it leaves the blood, some of it sticks to collagen (the elongated dye molecule favours this) and some is taken into cells,
including macrophages and neurons. The dye-protein complex is fluorescent (red emission) and this was the first fluorescent tracer of neuronal uptake and retrograde axonal transport.

Applied to sections, trypan blue stains everything and can be washed out completely. Slight alkalinity speeds up the procedure. In the presence of another anionic dye with smaller
molecules (like picric acid), trypan blue becomes selective for collagen, but it is no match for acid fuchsine or sirius red F3B. I'm sure Evans blue, which is a VERY similar compound, would have identical properties as a stain.

So: if you want to get rid of the Evans blue, wash the specimens in slightly alkaline water.

Eosin could also be used in the same way. If you're after very small leaks from your catheters, eosin might be more sensitive, because it's quite strongly fluorescent even without binding to anything (green-yellow emission). You could turn off the lab lights and use a Woods light to watch for leaks. Eosin is also removable by slightly alkaline water or by alcohols. 

John Kiernan
London, Canada
(kiernan[AT]uwo.ca)

Answer 2.

Evans blue and trypan blue both can be used to determine cell vitality - live cells exclude the dye(s), dead cells take then up - the trypan (Evans) blue exclusion test.

As far as catheter leakage is concerned, a fluorescent dye would certainly be a good choice. Cavers use them to trace underground rivers, and fluorescent dyes are used for a similar
purpose in opthalmology.

Russ Allison, Wales
(Deceased, 2002)

Back to Table of Contents

** Gallyas' stain

Question.

What is the Gallyas Stain, and what is it for?

Answer.

Ferenc Gallyas, in Hungary, has been studying and inventing silver stains for at least 30 years. They all involve the use of "physical developers" (an ancient and obsolete term from
photography). A physical developer is a mixture containing silver ions and a reducing agent, made stable for several minutes or even a few hours by other additives. Gallyas
introduced silicotungstic acid as a stabilizer. Earlier physical developers used gum arabic, gum mastic, albumen, albumin (no, they aren't the same) and other organic macromolecules.

The name of Gallyas is most often connected with his methods for Alzheimer's neurofibrillary tangles because neuropathologists are, by noble tradition, the biggest users of silver staining. However, there are several other silver staining methods, for a range of tissue components, developed by Gallyas. His work probably forms the rarely acknowledged basis of
immunogold-silver amplification for light microscopy and for some of the silver methods used to detect minute amounts of protein in Western blots.

Physical development was discovered, for photography and histology, by Liesegang (1911), and reintroduced to histological practice in 1955 by Alan Peters, who went on to become a great authority on the ultrastructure of nervous tissue, especially that of the cerebral cortex.

I don't know if this really answers the question, but it's interesting to look at the way someone's name gets attached to a method, even if at first there's doubt about _which_ method.

John Kiernan
London, Canada.
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Gram staining of sections (Brown & Hopps method)

Question.

I just did a B & H gram stain for the first time. All tissue stained various shades of purple against a clear background. There was no yellow or red staining at all. The protocol I used
replaced all acetone differentiation steps with 95% ethanol,  "to avoid over-decolorizing."

What am I doing wrong? Should I:

1. Use acetone instead of 95% ethanol, or a combination of equal amounts?
2. Use saturated aqueous picric acid?
3. Use 0.1% basic fuchsin (instead of 0.01%)?

Answer.

The following modifications of Brown & Hopps give consistent differentiation of Gram negatives with reduced risk of over-differentiation. Cellosolve is used instead of acetone, and
tartrazine instead of picric acid.

The crystal violet staining is as in the original method. Modifications are as follows:-

Substitute Lugol's or Jensen's iodine for Gram's to give a stronger crystal violet-iodine complex.

Use cellosolve (= ethylene glycol monoethyl ether = 2-ethoxyethanol) as decoloriser. The smell can be unpleasant, but it is slower in its action and more easily controlled.

Use 0.5% basic fuchsine, for 5 mins, to counterstain the Gram negative organisms.

After rinsing with water apply Gallego's differentiator (1% acetic acid with 2% formalin, in water) for 5 mins.


Rinse with water and flood sections with 1.5% tartrazine for 1 min.

Rinse the slides with water. Now take one slide at a time: 

Blot with filter paper, flood with cellosolve for 6 - 10 secs, blot again, and then place slide directly in xylene, 2 or 3 changes

Coverslip and mount. Repeat with the remaining slides, one at a time.

The extra step with the cellosolve seems to remove excess fuchsine from cytoplasmic elements in the background, thereby increasing visibility of Gram-negative bacteria.

Mike Rentsch
Lab. Manager, Aust.Biostain.
(ausbio[AT]nex.com.au)

Back to Table of Contents

** Oxidants for hematoxylin

Question.

Can a less toxic oxidizing agent be substituted for mercuric oxide in Harris's alum hematoxylin?

Answer.

Yes. Mercuric oxide for the oxidation of hematoxylin in Harris's hemalum can be replaced with sodium iodate (NaIO3) or other oxidants:

According to Hansen (1895), one of the following is, in general, needed for the oxidation of 1 gm of hematoxylin to hematein:

It is advisable to use only half of these quantities, to delay over-oxidation. Vacca (1985) suggested 75 mg NaIO3 per gm hematoxylin, and P. Bock (1989) suggested 98.5 mg NaIO3 per gm hematoxylin.

References.

Bock, P.: Romeis' Mikroskopische Technik; 1989
Hansen, F.C.C.: Eine schnelle Methode zur Herstellung des
Bohmersen Hematoxylins. Zoolog. Anz. 473; 1895.
Vacca: Laboratory Manual of Histochemistry; 1985.

Almost every hematoxylin can be used regressively, my favorite for general histology is "Mayer's acid hemalum, modified by Lillie":

"Dissolve 5gm hematoxylin by holding overnight in 700 ml distilled water; add 50 gm ammonium alum and 0.25 gm NaIO3. After these have gone into solution, add 300 ml glycerin C.P.
and 20 ml glacial acetic acid. May be used immediately; stain for 5 min."

Procedure. (5-7 ĉm paraffin sections, fixation: Bouin; manual staining.)

  1.  Sections to distilled water.
  2.  Sections to alum-hematoxylin (3 min).
  3.  Sections to acid alcohol (2-3 dips or until differentiated).
  4.  Rinse sections in tap water (about 10 sec, until most of the acid alcohol has dissapeared from the slide).
  5.  Rinse sections in 1% NaHCO3 in distilled water (1 min).
  6.  Rinse sections in distilled water (1 min).
  7.  Sections in 0.5% eosin Y in distilled water (30 sec).
  8.  Rinse sections in distilled water (a few dips, until most of the "free" eosin has dissapeared).
  9.  Dehydrate, clear, mount.

Yvan Lindekens
(yvan.lindekens[AT]rug.ac.be)

Back to Table of Contents

** M'Faydean's stain for anthrax bacilli

Question.

What is M'Faydean's stain?

Answer 1.

[ This has been put together from three replies to a question raised on the HistoNet newsgroup. ]

M'Faydean's stain is a simple stain using any well polychromed methylene blue (e.g. aged Loefflers). It is applied to heat-fixed smears for 10-30 seconds.

Polychroming (demethylation) is traditionally achieved by exposure of Loeffler's soln. to light and air for several months until it acquires a purplish tinge. However the oxidation process can be accelerated by application of heat as in Unna's method. (G. Gurr, 1963 p. 88 & 91); also E. Gurr, 1960, pp. 264-268).

Loeffler's methylene blue:

Warm the water to 50C, stir in methylene blue and add the other ingredients, cool and filter before use.

Polychrome methylene blue (Unna):

Dissolve methylene blue in water, add pot. carb. and alcohol, place in boiling water bath and evaporate to 100 ml.

Any other polychrome methylene blue formulation should work well also. [See also Answer 2 below.]

Results: 
Bacilli appear Navy Blue with Anthax showing a
narrow area (capsule) around and between bacilli that is reddish purple (metachromatic). A strong word of warning: many species of bacillus may also be encapsulated, e.g. Cereus etc. If you produce any positives get them confirmed at a Reference Microbiology Lab. for Infectious Diseases, or try the Armed Forces Institite of Pathology.

Gurr doesn't give any further references in his book as to M'Fadyean, whether the method was published or by personal communication.

References.
"Encyclopedia of microscopic stains," by Edward Gurr. London: Arnold, 1960. (pp 264-268)
"Biological Staining Methods." by George T. Gurr. 7th Edition. 1963. (Published by George T. Gurr Ltd. 136-144, New King's Road, London, S.W.6.)

Mike Rentsch
Australian Biostain P/L
(ausbio[AT]nex.com.au)
Ian Montgomery
(I.Montgomery[AT]bio.gla.ac.uk)
Bryan Hewlett
(hewlett[AT]exchange1.cmh.on.ca)

Answer 2.

In 1903 John M'Fadyean described red coloration of the capsules of Bacillus anthracis organisms in blood taken from dead farm animals and stained with an aged solution of methylene blue. This is now recognized as an example of metachromasia, due to binding of oxidation products of methylene blue (such as azures A, B and C) to the poly(D-glutamic acid) of which the capsule of B. anthracis  is largely composed. In recent years
oxidized (polychromed) methylene blue has been replaced by azure B (CI 52010), a thiazine dye that can be manufactured as a pure substance.

The staining solution is made by dissolving 30 mg azure B (Sigma-Aldrich 22,793-5) in 3 ml of 95% ethanol and adding 10 ml of 0.01% aqueous potassium hydroxide. The final dye concentration is 0.23% (an almost saturated solution of azure B). Air-dried smears are fixed in methanol or ethanol, stained for 1-5 min, rinsed in water and allowed to dry before examining with an oil immersion objective.
Anthrax bacilli are blue with red capsules.

References.
M'Fadyean, J. 1903. A peculiar staining reaction of the blood of animals dead of anthrax. J. Comp. Path. 16: 35-41.
Owen, M.P., Kiernan, J.A. 2004. The M‘Fadyean reaction: a  stain for anthrax bacilli. Biotech. Histochem. 79: 107-108.
Owen, M.P., Schauwers, W., Hugh-Jones, M.E., Kiernan, J.A., Turnbull, P.C.B., Beyer, W. 2013. A simple, reliable M’Fadyean stain for visualizing the Bacillus anthracis capsule. J. Microbiol. Methods 92: 264-269.

John A. Kiernan
Department of Anatomy & Cell Biology
University of Western Ontario
London, Canada
(jkiernan[AT]uwo.ca)

Back to Table of Contents

** Microglia with Griffonia lectin.

Question.

I have been trying to stain for microglia in paraffin sections of rat brain using peroxidase-labeled Griffonia simplicifolia lectin (GSI-B4-HRP) from Sigma. It has been used in various papers for staining of active and resting microglia but I cannot seem to get it to work. Are there any tricks that I might be missing?

Answer.

I have not used this lectin for microglia but have used it for other things. The purity varies considerably because the seeds of Griffonia, when extracted, may yield just one lectin or several isolectins (depending on the seeds), and the B4 lectin is then purified from this mixture. I have found a lot of variation from batch to batch but more so from manufacturer to manufacturer. The best luck I had with this lectin was from Vector Laboratories, Burlingame, California, who specialize in the production of lectins. I have also had problems with some lectin-HRP conjugates. In my experience the conjugates (especially the HRP ones) have only a limited shelf life and this can lead to background staining. Part of your problem may be that lectin binding can be significantly altered by fixation and processing. I would suggest that you first try it on frozen sections to determine whether the conjugate you have is working. 

This lectin usually requires the availability of calcium ions to bind. If you are using OCT freezing compound, this contains sufficient calcium if you don't remove the OCT before staining.

I do not have the latest Vector catalog available at the moment but believe that they have an antibody against GSI B4. This might be a better approach if the problem is one of conjugate
breakdown or excessive background staining.

Another point is that the lectin binding can be easily confirmed with negative (inhibited) controls, inhibitors for GSI B4 include:

o,p-Nitrophenyl-N-acetyl-alpha-galactosamine
Galactose-N-acetyl-alpha-1,3-galactose
Methyl-N-acetyl-alpha-galactosamine
N-acetylgalactosamine


Barry R. J. Rittman
Univ. Texas HSC Dental Branch, Houston, Texas
(barry.r.brittman[AT]uth.tmc.edu)

Back to Table of Contents

** Picro-sirius red staining

Question.

I have been asked to do a "picrosirius" staining procedure. What is it?

Answer.

Picro-sirius red is a solution of sirius red F3B (0.1%) in saturated aqueous picric acid. It is typically used after an iron haematoxylin nuclear stain, much as Van Gieson, but for 60 minutes. Rinse in slightly acidified water and dehydrate in three changes of absolute alcohol. The result is similar to Van Gieson (Collagen red, cytoplasms & red cells yellow) but sirius red F3B shows thinner fibres that are often missed by Van Gieson. The real difference is seen by using a polarizing microscope. With crossed polars the collagen fibres, even very thin ones, appear in brilliant orange, yellow and green colours against a black background. Basement membranes, though stained, do not exhibit this birefringence because their collagen fibres are not aligned.

The dye is one for which the Biological Stain Commission offers testing and certification, but some major American vendors do not have it in their catalogues. There are many synonyms. The Colour Index application name is Direct red 80, and the CI number is 35780. Don't use a dye that is not CI 35780 even if it has the words sirius and red in its name.

Some references:

Puchtler H & Sweat F 1964. Histochemie 4, 29-54.
Puchtler H, Sweat FS & Valentine LS 1973. Beitr. Pathol. 150, 174-187.
Junqueira LCU, Bignolas G & Brentain RR 1979. Histochem. J. 11, 447-455.
Lillie RD 1977. Conn's Biological Stains, 9th ed. Baltimore: Williams & Wilkins.
Colour Index CD-ROM (1997) Society of Dyers & Colourists, Bradford, England.
Dapson RW, Fagan C, Kiernan JA, Wickersham TW (2011) Certification procedures for sirius red F3B (CI 35780, Direct red 80). Biotech. Histochem. 86: 133-139.

World Dye Variety.  http://www.worlddyevariety.com/direct-dyes/direct-red-80.html.

John A. Kiernan,
London, Canada
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Iron hematoxylin: ripening not needed.

Question.

Why does Bancroft and Stevens tell me to ripen my alcoholic hematoxylin for a month, when the ferric chloride oxidizes it instantly when you combine the two solutions to make the Weigert's hematoxylin stain?

Answer.

Because B & S is wrong (a very unusual thing in that superb book), and you are right.

For what it's worth, my experiences and occasional experiments fully support the conclusions written in the classical works of Baker, Lillie, Gabe and others. Ferric ions instantly oxidize
hematoxylin to hematein and they also form part of the black complex that is retained in cell nuclei.

John Kiernan
London, Canada
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Enzyme histochemistry on cell cultures

Question.

How do you perform enzyme histochemistry (NADH Dehydrogenase, succinic Dehydrogenase, cytochrome oxidase) on cultured cells grown on slides? Would you use a detergent (or other means) to permeabilize membranes prior to application of the reaction medium?

Answer.

I just take the coverglass from the culture medium, give it a rinse in buffer, incubate for required time, wash gently, then mount. No fixing, no detergent; just incubate and mount. It works, so why complicate matters?

Ian Montgomery
(I.Montgomery[AT]bio.gla.ac.uk)

Back to Table of Contents

** Malachite green in stain for Cryptosporidium

Question.

How do you do a malachite green stain for Cryptosporidium?

Answer.

The Cryptosporidia are stained by carbol fuchsine; malachite green is a counterstain for the background.
This is the procedure that I use. (I also do the parasitology here.) It works fairly well but is not the best diagnostic technique for Cyrptosporidia. There are Meriflour commercial kits that are better than this stain.

A MODIFIED ZIEHL-NEELSEN TECHNIQUE FOR CRYPTOSPORIDIUM

This is used on fecal smears.

Solutions.

Concentrated carbol fuchsine

     Combine in the listed order

10% Sulfuric Acid

5% Malachite Green


Procedure.

1.   Make a thin smear from the fecal sample.
2.   Dry the smear at room temperature.
3.   Fix the smear in absolute methanol for 2-5 minutes.
4.   Dry at room temperature
5.   Fix briefly in a flame.
6.   Stain with concentrated carbol fuchsine for 20-30 minutes without heating.
7.   Rinse in tap water.
8.   Differentiate with 10% sulfuric acid for 20-60 seconds. (Concentrations from 0.25 to 10% can be used; we use 10% sulfuric acid.)
9.   Rinse in tap water.
10. Counterstain with 5% malachite green for 5 minutes.
11. Rinse in tap water.
12. Dry at room temperature.
13. Examine under oil.
14. Cryptosporidia will stain bright red with a blue-green background.

Roberta Horner
Penn State University
(rjr6[AT]psu.edu)

Back to Table of Contents

** Confusing dye names (lissamine fast red as an example)

Question.

Is there another name for Lissamine Fast Red? I can't find it under this name in any dye catalog.

Answers.

Five or six people identified at least three different dyes in the answers to this HistoNet query. This emphasizes the importance of identifying dyes by Colour Index numbers whenever possible. A name like "Lissamine" has no chemical significance and may be attached to widely differing compounds! Some opinions follow (mine is No. 3). Probably all are correct, and there are different uses for the simlarly named dyes.

J. A. Kiernan

1. Another name for Lissamine Fast Red is Acid Red 37. You can try BDH with next Cat no 341772K and it comes in 25 gram containers.

2. I suspect that the dye you're looking for is Sulforhodamine B, also known as Lissamine rhodamine B 200, Acid rhodamine B. The dyers assoc. refer to it as C.I.Acid Red 52. Its C.I.Number is C.I. 45100.

3. The nearest entry in Conn's Biological Stains (9th ed,, 1977) is amidonaphthol red 5B (C.I. 18055, Acid violet 7). Synonyms include lissamine red 6B and many others. The Colour Index number (or application name) is the most reliable identifier of a dye. It should be mentioned in the published instructions for a method. If it isn't, your best bet is to find another,
properly explained staining technique for the job.

4. My assumption has been that the lissamine fast red referred to is the same that Lendrum used in his published method for muscle fibres. The dye name has the synonym Acid red 37,
Colour Index no. 17045. It appears in Floyd Green's excellent reference book "The Sigma Aldrich Handbook of Stains, Dyes and Indicators" with the further synonyms anthranal red G and
fast light red B. The dye synonyms list I refer to most frequently as an easy-to-use first stop was published as a "give away" by Difco in 1974.

5. Lissamine fast red is not mentioned in the 10th (2002) edition of Conn's Biological Stains.

Back to Table of Contents

** Mayer's and Gill's hematoxylins

Question.

I would like to know the differences between two types of hematoxylin: Mayer's and Gill's.

Answer 1.

Haematoxylin dye concentration for Mayer is 1 gm/L compared with 2 gm/L for Gill-I. The preservative for Mayer's is chloral hydrate and for Gill it is ethylene glycol. The acidifying agent
for Mayer's is citric acid, whereas for Gill it is acetic acid.

Both have very good shelf lives of two years or more under correct storage conditions. They both are used mainly as progressive stains, and are well suited to use as counterstains as well. Gill-I has some some strong adherents for progressive cytology staining.

It is possible to make either of these in a non-toxic formulation (NaIO3 as the oxidant) without compromising performance or shelf life.

Mike Rentsch
(ausbio[AT]nex.com.au)

Answer 2.

Both stains are hemalums: they are solutions containing hematein (from oxidized hematoxylin), an aluminium salt (the "mordant," which forms dye-metal complexes with hematein), an organic acid to adjust the pH, and a hydrophilic compound (glycerol, ethylene glycol or chloral hydrate). The last ingredient is variously said to modify the solubilities of other ingredients, retard the oxidation of hematoxylin, "preserve" the solution or do nothing at all. In modern hemalums the hematein is generated by adding enough of an oxidizing agent (most often the iodate ion) to oxidize about half the hematoxylin. The unoxidized hematoxylin provides a reservoir from which more hematein is slowly produced by atmospheric oxidation. This compensates for the atmospheric over-oxidation of hematein to trioxyhematein (which is useless), thereby prolonging the life of the solution.

The compositions of Mayer's and Gill's hematoxylins are set out below. Mayer's recipe was published in 1863, that of Gill, Frost and Miller in 1974. Gill's hematoxylin closely resembles "haematal-16," a mixture published by J. R. Baker in 1962 that contained ethylene glycol but no organic acid.

MAYER'SGILL'S
Hematoxylin 1 gHematoxylin 2 g
Potassium alum 50 g (0.09M)Aluminium sulfate 17.6 g (0.03M)
Sodium iodate 0.2 g Sodium iodate 0.2 g
Citric acid 1 gAcetic acid 40 ml
Chloral hydrate 50 gEthylene glycol 250 ml
Water to make 1000 ml Water to make 1000 ml
Molar ratio of Al3+ ions to haematein molecules in the freshly made solution:  32 Molar ratio of Al3+ ions to haematein molecules in the freshly made solution:  11


A high ratio of aluminium:dye slows down staining and increases the selectivity for nuclei. Both these hemalums are used progressively; in principle, Gill's should stain more quickly than Mayer's. The effect of excess aluminium is seen most strikingly with Ehrlich's hematoxylin, which is saturated with alum and relies on atmospheric oxidation (slow) to provide a low concentration of hematein from an initially large (6 to 7 g/L) reservoir of hematoxylin. Ehrlich's hematoxylin is the slowest of the progressive hemalum stains (up to 30 minutes, compared with 3 to 10 minutes for Mayer's or Gill's). Hemalums for regressive nuclear staining (e.g. Delafield's, Harris's) have lower aluminium:dye ratios than the progressive stains. Acid-alcohol extracts the dye-metal complex more slowly from nuclei than from other components of tissues.

Some references. These are for practical, rather than chemical or theoretical (i.e. speculative) aspects of hemalum staining.

Baker, J.R. (1962). Experiments on the action of mordants. 2. Aluminium-haematein. Quarterly Journal of Microscopical Science 103: 493-517.
Bancroft, J.D. & Cook, H.C. (1984). Manual of Histological Techniques. Edinburgh: Churchill-Livingstone.
Bancroft, J.D. & Stevens, A., eds. (1996). Theory and Practice of Histological Techniques, 4th ed. London: Churchill-Livingstone.
Ehrlich, P. (1886). Die von mir herruhrende Hamatoxylinlosung. Zeitschrift fur wissenschaftliche Mikroskopie 3: 150.
Gill GW, Frost JK, Miller KA (1974) A new formula for a half-oxidized hematoxylin solution that neither overstains nor requires differentiation. Acta Cytol. 18: 300-311.

Gill GW (2010a) Gill hematoxylins: first person account. Biotech. Histochem. 85: 7-18.

Gill GW (2010b) H&E staining: oversight and insights. In Education Guide: Special Stains and H&E (Kumar GL & Kiernan JA, eds) pp.119-130. Free download (whole book) from Agilent Technologies (formerly DAKO).

Kiernan, J.A. (2015). Histological and Histochemical Methods:
Theory and Practice, 5th ed. Banbury, UK: Scion.
Kiernan JA (2018) Does progressive nuclear staining with hemalum (alum hematoxylin) involve DNA, and what is the nature of the dye-chromatin complex? Biotech. Histochem. 93: 133-148.
Llewellyn BD (2009) Nuclear staining with alum-hematoxylin. Biotech. Histochem. 84: 159-177.

Llewellyn, Bryan. Stains File. http://stainsfile.info/ (This Web site has a splendid, possibly comprehensive, collection of hematoxylin stain formulations, also available as a 110-page  PDF file:  http://stainsfile.info/downloads/hxformulas.pdf.)

J. A. Kiernan
Department of Anatomy,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Effects of pH on staining by dyes

Question.

Many stains are acidified, but some are adjusted to a neutral or even an alkaline pH. Why? Are different dyes differently affected by pH changes?

Answer.

For a full answer to your question you will need to refer to a textbook of histological techniques. Here is a simplified answer. It applies to basic (cationic)
and acid (anionic) dyes with fairly small molecules. Attraction of opposite electric charges plays a major part in staining by such dyes. 

The structural macromolecules in a section of a tissue have numerous side-chains that can form either positive or negative ions.

Acid dyes (attracted to positive sites in tissue). The positive ions are associated mainly with proteins.

The side chain of the amino acid arginine (a guanidino group) is a strong base. That means it always carries a positive charge, even at a high pH. It can therefore always attract a negatively charged dye ion. At pH 9 or above, all staining by a simple basic dye (biebrich scarlet is commonly used) is due to arginine.

The other organic group that can form positive ions is the amino group, which occurs at the N-terminus of every chain of amino acids and on the end of the side-chain of lysine. Amino groups are weak acids: at high pH they are not ionized, but at low pH an amino group collects a hydrogen ion (proton) from the solvent and becomes positively charged. The amino group of lysine can collect a proton even when there are not many around, as in a neutral or slightly alkaline medium. Consequently, lysine behaves as a cation and binds acid dyes at pH about 8 or below. N-terminal amino groups are weaker acids: they cannot be protonated much above pH 6, so they are not stained by neutral or alkaline solutions of acid dyes. More and more amino groups become
protonated (ionized) as the pH is lowered. Staining with an acid dye therefore occurs more rapidly and more strongly from the more acid solutions. At a    pH around 2, these dyes stain everything.

The foregoing remarks apply to a "typical" acid dye with sulfonic acid side-chains. Sulfonic acids are strong acids; they exist in solution only as sulfonate anions. (Eosin is not "typical" in this way because it is a salt of a weak acid. Moreover eosin solutions must not be acidified too much or insoluble unionized eosin will be precipitated, leaving a colorless solution.)

Basic dyes (attracted to negative sites in tissue). The three negatively charged chemical groups present in a section are:

Alkaline solutions of basic dyes are used for staining semi-thin plastic sections. With anything thicker the color is too dark to show structural details. For more selective staining, basic dyes are applied as acidic solutions. At pH 1 only the sulfated materials are displayed. As the pH rises from 2.5 to 4.5, nuclei and RNA stain with increasing speed and intensity.

Remember that these simplified arguments do not apply to all dyes, or even to those most commonly used in routine work.

Further reading.

Horobin, R.W. (1982). Histochemistry: An Explanatory Outline of Histochemistry and Biophysical Staining. Stuttgart: Gustav Fischer.
Kiernan, J.A. (2015). Histological and Histochemical Methods: Theory and Practice, 5th ed. Banbury, UK: Scion.
Horobin, R.W. (1988). Understanding Histochemistry: Selection, Evaluation and Design of Biological Stains. Chichester: Ellis Horwood.
Lyon, H. (1991). Theory and Strategy in Histochemistry. A
Guide to the Selection and Understanding of Techniques. Berlin: Springer-Verlag.

John A. Kiernan
Department of Anatomy & Cell Biology
University of Western Ontario
London, Canada.
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Histochemical stain for arsenic

Question.

Is there a staining method for showing the presence of arsenic in tissues?

Answer.

Fix in 10% formalin containing 2.5% copper sulfate for 5 days. Wash for 24 hours in running water. Process and embed in parffin wax. Deparaffinized sections show green granules of Scheele's green (CuHAsO3) which, though insoluble in water, is dissolved by acids and by ammonium hydroxide. By substituting copper acetate for the sulfate, the green granular paris green or cupric acetoarsenite is produced. Its solubilities are similar (Castel's method,
Bull. Histol. Appliq. 13: 106, 1936). A light safranine counterstain gives good contrast.

Source: R. D. Lillie 1965. Histopathologic Technic and Practical Histochemistry, 3rd ed. p. 445 [or 4th ed. (1976), p. 548]. New York: McGraw-Hill.

Roy Ellis
(roy.ellis[AT]imvs.sa.gov.au)

Back to Table of Contents

** Giemsa staining of blood smears: several hints

Question.

My methanol-fixed blood smears are not staining reliably with Giemsa. Some advice is needed, please.

Answer

Fixation of well dried (at room temperature) PB smears can vary from 1-10 minutes; automated systems tend to use about 1-2 minutes and use the methanol only once. For manual staining, most labs would fix for about ten minutes. Precautions must be taken against absorption of water from humid air. The methanol is usually replaced twice daily, but more frequently at those times of the year when humidity is high.

The first sign of unacceptable water content in the fixing methanol will be the appearance of clear refractive spaces on the biconvave surfaces of erythrocytes: perhaps only a few cells per high-power field, but this will increase further as the water content increases, and eventually the films will lose all diagnostic value. Replacement of the methanol when you see more than say 1-2/HPF might not be a bad idea. This artifact may also be seen in some automated systems where the stain pack is not turned over very quickly. Rather than replacing the stain pack, economy of reagent can be maintained by manually fixing the slides before thay go on the machine. This is particularly so for the older Hematek grey models.

Caution. Longer fixation times are required for bone marrow smears: 15-20 minutes, and always use fresh methanol for these.

Most persons using Giemsa prefer to stain the smear first with May Grunwald or Jenner stain, either using it neat or diluting 1:2 with buffer. This pre-step improves the granule definition and clarity, and also changes the traditional reddish purple of nuclei with plain Giemsa to a blue purple as seen with Wright's stain.

The selection of Sorensen's (phosphate) buffer will vary from pH 6.4-7.2, with the lower pH being most popular with Wright's rather than Giemsa. The aim is to select a pH that produces a colour balance that readily allows the user to differentiate between normochromic and polychromic red cells and to distinguish toxic granulation when present, this is usually pH 6.8. If looking for malarial parasites, then a pH of 7.2 is preferable because it allows better contrast to detect chromatin dots, trophozioites etc.

Dilution of the Giemsa solution is best done immediately before use and will vary from 1:8 to 1:12 depending upon your protocol. As a general rule of thumb the higher dilutions require longer staining times of about 20 minutes, and the less dilute stains need between 6 and 12 minutes, depending upon tthe quality of the Giemsa. It was frequently claimed that the longer times gave better definition, but I must admit that I've seen short timed smears that are every bit as good.

For many years good quality Giemsa would be stable after dilution for 6 to 8 hours. For the last 2 or 3 yrs, however, the best you can hope for is 3 to 4 hours. After dilution the solution starts to deteriorate, with the appearance of floccules and a subsequent loss of staining ability or strength. As the time progresses you may need to compensate by increasing the staining time, but after 3 hours you will need to replace it.

Recipes for Giemsa vary, whether it be that of Hayhoe or of Dacie & Lewis, and measurements may be by weight or volume. Stock solutions that have a 50% by volume content of glycerol (Analar or USP) are the most stable. Under no circumstances ever heat your glycerol to more than 45C, even though most texts say 56C. Above these temperatures there is a risk of oxidation, even in the stock solution, I use 45C as a cut-off point to give me a safety margin. Dye content will also vary from 0.45 to 0.8%. Lillie's comments should considered here. After standing for up to 5 days, filtration to remove undissolved material is essential.

Differentiation, by giving the slides two rinses in buffer of two minutes each, is fairly standard, but you can overdo it. A single rinse of three quick dips may in fact suffice. It will depend upon your Giemsa solution and tastes. If overstaining is a problem then consider adding methanol to your buffer rinse,
starting at 5% and adjusting according to results, followed by a water rinse to remove solvent.

Mike Rentsch, "Histomail," Downunder

Back to Table of Contents

** Automated H & E staining problems

Question.

We are having a problem with our H & E being inconsistent (sometimes from day to day, sometimes from batch to batch). We have an automated stainer and use bought solutions of hematoxylin and eosin. We do not change program times or reagents, yet sometimes our stain is light and sometimes it is dark (preferred). We have not changed any processes, vendors, or manufacturers, but our stain is continually changing.

The same hematoxylin, eosin, alcohol, and xylene are on our manual stain line. We stain those following the same times as on the auto stainer and they come out perfect every time.

Answer.

Is your manual stain set-up absolutely identical to your automatic stainer set-up, in time values as well as reagent set-up? If so, the times on the machine may be too short, as explained below.

You commented that when you stain from your manual set-up the staining results are fine. I would recommend that you "manually" stain using your automatic stainer set-up. If you are able to acheive the desired results, then we can identify the mechanical differences between human and machine staining. It would be helpful to compare your stain programs (Manual procedure and automatic times).

Analyzing the stain, is the nuclear stain OK but the counterstain is too light? Is the nuclear stain too light but the counterstain OK? Is the nuclear stain too light and the counterstain too light? Are the stains consistant in their lightness throughout the specimen and throughout all sections on the slide? Do you notice an improvement in the stain after the new reagents have become somewhat diluted?

One of the biggest differences between hand and machine staining is how the surface tension of the reagent currently on the slide is broken and then replaced by the next reagent. When we stain by hand we exert much more and varied force than a machine does when plunging the slides into the reagent. We also knock off more reagent, so less of the reagent clings to the slide with each move. A stainer (machine, not human) simply lowers the slides slowly, in a single plane, into the reagent. Even the agitation of the machine staining is in that single plane (up and down) movement. When we stain by hand we cause the reagent in the dish to bombard the slide from several angles and with greater force that breaks the surface tension in less time than it takes a machine can accomplish. Therefore longer exposure times (of tissues to stain) may be required on a machine to yield the same results as hand staining.

When programming the machines I find it necessary to watch the hand staining carefully in order to make an accurate translation of a "dip" to a time value that the machine could reproduce. A "dip" in acid alcohol in manual staining may not be able to be reproduced by a machine. I may be able to use 1% acid alcohol in hand-staining but have to use 0.5% acid alcohol on the staining machine with a 2-second timing value to get the same results. Ten "dips" in a manual stain may require 30 seconds on a machine. Ten "dips" in a manual alcohol step may require 1 minute on a machine for the same results.

One of the things we need to remember is that the machine will move the slides exactly the same way for the programmed time. We humans (consciously or unconsciously) adjust our handling of the slides based on how the sections or even the reagents look.

Nancy Klemme,
Sakura Finetek USA, Inc.
Torrance, CA 90501
(nancy.klemme[AT]sakuraus.com)

Back to Table of Contents

** Verhoeff's stain for myelin and elastin

Question.

Can Verhoeff's elastic tissue stain (iron hematoxylin with iodine) be used to stain myelin sheaths?

Answer.

H. Puchtler and F. S. Waldrop published "On the Mechanism of Verhoeff's Elastica Stain: A Convenient Stain for Myelin Sheath" in Histochemistry 62:233-247 (1979).

They stated: "Verhoeff's elastica stain is definitely not specific for elastin and is inferior to orcein and resorcin-fuchsin because of the required differentiation with its inherent bias to produce patterns which conform to expectations. However, Verhoeff's elastica stain is far superior to other metal-hematein technics for myelin sheaths. The combined Verhoeff-picro-Sirius Red F3BA stain can be performed in 30 min and does not require differentiation. It is therefore suggested to reclassify Verhoeff's elastica stain as a method for myelin sheaths."

Freida Carson
(FreidaC[AT]aol.com)

Back to Table of Contents

** Acridine orange method for DNA and RNA

Question.

Can acridine orange be used to stain DNA and RNA in different fluorescent colors in sections as well as in smears of cells?

Answer.

In the late sixties, early seventies, I used to use the original method (Bertalanffy F.D. A new method for cytological diagnosis of pulmonary cancer. Ann. New York Acad. Sci. 84: 225-238) for screening cytology slides fixed in alcohol for malignant cells, and I thought it worked quite well, as did my
pathologist at the time. The DNA of the nucleus fluoresces brilliant green, and RNA in the cytoplasm of malignant cells is brilliant orange. However, I have never met a cytotechnologist who liked the method, so, when I was forced to hire one because of work load, she quickly relegated this technique to the garbage bin of history.

I don't know of anyone who is currently using the technique. However, as we found it very useful at the time, I worked out a method for using it on paraffin sections, that gives very similar results to the alcohol fixed smears.

Acridine orange stain.

Acridine orange (C.I. 46005) 0.05 gm
Distilled water 500.0 ml
Acetic acid 5.0 ml

   Note. Some batches of acridine orange work better than others. This dye is not one of those tested and certified by the Biological Stain Commission.

  1. Procedure.
  2. Bring paraffin sections to water in usual manner.
  3. Stain sections in acridine orange stain for 30 minutes.
  4. Rinse sections briefly in 0.5% acetic acid in 100% alcohol.
  5. Rinse sections in two additional changes of 100% alcohol.
  6. Rinse sections in two changes of xylene.
  7. Mount sections in a non-fluorescent resinous medium.

Results:  DNA brilliant green. RNA brilliant orange. Most gram positive microorganisms brilliant orange. Most gram negative microorganisms (including Helicobacter) green to pale orange.

Kerry Beebe
Kelowna General Hospital
Kelowna B.C. Canada
(bbracing[AT]silk.net)

Back to Table of Contents

** Quickly finding something in a newly cut section

Question.

Is there any way to quickly stain paraffin sections so that I can evaluate whether or not I need to cut further into the block?

Answer 1.

We used to use a cotton ball moistened with dilute methylene blue to wipe over the surface of the block. This gave us a good idea of the tissue at that level and helped greatly in the orientation. If you prefer you can always place a cut section on a slide and add several drops of dilute aqueous methylene blue (say 0.05-0.1%), this also works well. No need to mount the section.

Barry Rittman
(barry.r.brittman[AT]uth.tmc.edu)

Answer 2.

If the structure is fairly large you can use a pseudo-interference contrast illumination method to see structure in the section. Just move the objective of the
microscope slightly to one side of its normal position and you can see 3D structure without doing any deparaffinizing or staining. You will be surprised how much detail you can make out. This is a great method for finding glomeruli in kidney frozens.

Tim Morken
San Francisco, CA
(
timothy.morken[AT]ucsf.edu
)


Answer 3.

I have used the following technique when searching for glomeruli in kidney biopsies.

Mount the section on the slide as usual. Place the slide on the microscope stage, under a 10x objective. Close the condenser aperture down, and lower the entire condenser away from the microscope stage.

What should result is a slightly out of focus image of the unstained tissue section. You may have to adjust the settings of the aperture and condenser. This works well for large structures such as the glomerulus in the nephron of a kidney.

Patrick M. Haley
HistoTechNologies, inc.
(pmhales[AT]cybergap.net)

Back to Table of Contents

** Fluorescent lectins: general method

Question.

Can anybody give me a working concentration range for staining with lectins conjugated with TRITC?

Answer.

The general rule of thumb when staining with fluorescent protein conjugates is to bracket around 10 micrograms per mL. When using a good fluorescent IgG conjugate, I found that 5 micrograms/mL was a bit dim, whereas 20 micrograms/mL often had a bit too much background. This rule of thumb depends somewhat on the fluorophore (some yield a higher background, etc), but for TRITC conjugates, 10 micrograms/mL usually works well.

Although the molecular weight of your lectin is probably is a bit less than that of IgG, a 2-3 fold difference in molecular weight prabably won't make that much of a difference. I used to use a TRITC conjugate of wheat germ agglutinin at 10 micrograms per mL and it stained beautifully.

Karen Larison, in Oregon
(larisonk[AT]uoneuro.uoregon.edu)

Back to Table of Contents

** Methyl blue and methylene blue

Question.

A method calls for methyl blue, in a mixture with eosin Y. The nearest name I can find on a bottle is methylene blue. Will it be OK to use it instead?

Answer.

No! The only thing these two dyes have in common is a blue color. Otherwise they have opposite staining properties.

Methyl blue, an acid triphenylmethane dye, is one of the components of aniline blue. Aniline blue is a generic name that includes methyl blue (C.I. 42780; Acid blue 93) and water blue or ink blue (C.I. 42755; Acid blue 22). Most dyes that are sold under these names are mixtures of both dyes, but some are mostly methyl blue. A contaminant known as sirofluor is also present in these dyes, and is exploited in fluorescent stains for callose in plants. In staining applications any dyes sold as aniline blue, methyl blue and water blue are interchangeable, provided that the batch meets the Biological Stain Commission's standards in respect of content of reducible blue dye and performance in standardized staining procedures.

Methyl blue (aniline blue) is used in Mann's eosin-methyl blue method and in various trichrome stains such as Mallory's, Gomori's, Cason's and Heidenhain's AZAN. It colors collagen fibers and a few other materials.

Methylene blue (C.I. 52015; Basic blue 9) is a basic thiazine dye. It may have more scientific uses than any other dye. As a simple stain, applied from a mildly acidic solution (pH 3 to 4) it colors nucleic acids and acidic carbohydrates. At neutral or alkaline pH is colors everything. Methylene blue is used in
conjunction with eosin and other dyes in stains for blood cells and parasites, and it is also extensively used in bacteriology. Products of degradation (demethylation or "polychroming") of methylene blue are essential components of the commonly used Romanowsky-Giemsa stains for blood cells. The purple coloration of leukocyte nuclei and magenta color of malaria parasites seen with Wright's and Giemsa's stains, are due to one of these products, the dye known as azure B (C.I. 52010).

Methylene blue (and some other thiazine dyes) can provide beautiful and selective staining of the living neurons and their cytoplasmic extensions, and has been much used to demonstrate the innervation of peripheral tissues. Methyl (aniline) blue cannot be used in this way.

References. 

Conn's Biological Stains. Entries under the various named dyes.
Sigma-Aldrich Handbook of Stains, Dyes and Indicators.
Entries under the various named dyes.

John Kiernan
London, Canada
(kiernan[AT]uwo.ca)

Back to Table of Contents

 


 

IMMUNOHISTOCHEMISTRY

 

** Paraffin or frozen sections for immunohistochemistry

Question.

Are paraffin or frozen (fixed) sections better for IHC? I've had great success in the past with frozen or vibrating microtome sections, and have been trying paraffin lately, but haven't got any good results.

Answer 1.

Generally frozen sections are better for IHC because the antigenic content is well preserved (provided the tissue is snap frozen rapidly, preferably in isopentane, then stored at -70C). A "good" frozen section cut at about 5 microns should provide adequate morphology.

The advantages of paraffin tissue blocks is that larger pieces of tissue can be used, and morphology is a degree better, storage is easier, etc.

The disadvantage of paraffin blocks is the fact that the processing of the tissue (especially when preserved in common fixatives such as formalin or other formaldehyde-based solutions) cross-links certain proteins in and on the cells. Preatreatment to "unmask" cross-linked antigens is often essential. Antigen retrieval techniques include microwaving in citrate buffer and pressure cooker techniques. However, some antigens are destroyed by paraffin processing,
so for these the manufacturer of the antibody should recommend the use of frozen sections only.

Stephen Wayne
Cambridge Antibody Technology
The Science Park, Melbourn,
Royston, Cambridgeshire SG8 6JJ
England.
(stephen.wayne[AT]camb-antibody.co.uk)

Answer 2.

In general, immunoreactivity is often better in cryostat sections than in wax sections, however tissue morphology is usually not as clear. If you are getting satisfactory results with cryostat sections, then I would probably recommend sticking with that technique. However, if need to use wax sections for
whatever reason, there are several ways of tweaking the protocal to try and improve the staining. Any good IHC text book will outline most of these.

Off the top of my head, I would suggest playing around with the fixation conditions or trying some form of antigen unmasking step (particularly if you are currently seeing no specific staining at all).

Ian Jones, PhD
School of Biological Sciences,
Queen Mary and Westfield College,
University of London, England.
(I.W.Jones[AT]qmw.ac.uk)

Back to Table of Contents

** Inhibiting endogenous peroxidase

Questions.

1. What is the best way to inhibit endogenous peroxidase
activity before doing an immunohistochemical method?
2. How long can methanol/H2O2 mixture (for quenching
endogenous peroxidases during IHC) be kept? or should
it be freshly made each time before use?

Different people favour different methods! Here are five
suggestions. All are claimed to work well, so probably
you should start with whatever you think is the easiest
and cheapest.

Answer 1.

We use a homemade version: PBS with 0.03% hydrogen peroxide, and 0.1% sodium azide. Very gentle; doesn't knock sections off slides (frozens); can make up a one-week supply. Use it once, then discard (we use dropper bottles). Our PBS is at pH 7.4. We collect the leftover for chemical disposal of sodium azide.

OR you can purchase DAKO peroxidase blocker with 0.03% H2O2 This block works best with our mouse antibodies as it does not interfere with some of the IHC staining/per recommendation of PharminGen. They use DAKO [now Agilent] also, and if there are capillary gaps involved, this does not produce the crummy bubbles that drive one crazy.

Gayle Callis
(
gayle.callis[AT]bresnan.net)

Answer 2.

We prepare 600ml vats of methanol/H2O2 for use on a DRS601 and replace these weekly. It's left on the machine for 5 working days then dumped. We're handling about 150 ICC slides/day.

Elwyn Rees
(100131.74[AT]compuserve.com)

Answer 3.

Just a personal note on the use of methanol in blocking solutions; I have also found that methanol can be harmful to some antigens, both hemopoetic and some infectious disease antigens. We have found that performing our endogenous peroxidase inactivation prior to any antigen retreival step (either enzyme digestion or heat induced) works best. For antigens sensitive to methanol and frozen sections we use PBS containing 0.1% Na azide and 0.5% H2O2 with excellent results. Just be sure to wash the slides well after this step because the Na azide is a potent peroxidase inhibitor which will eliminate any specific
staining quite well. Using polylysine coated slides will generally keep frozen sections from lifting off.

Brian J. Chelack
(chelack[AT]admin3.usask.ca)

Answer 4.

Quenching with the glucose oxidase method works very well, and is very gentle on sections, particularly frozen sections. The only drawback is a bit more preparation of solutions, but in the long run is a very COMPLETE quenching, better than hydrogen peroxide, according the original publication and method. I highly recommend it.

Gayle Callis
(
gayle.callis[AT]bresnan.net)

Answer 5.

Complete inhibition of endogenous peroxidase (including activity in leukocytes and erythrocytes) can be achieved by treating formaldehyde- or acetone- fixed smears or sections with 0.024 M hydrochloric acid in ethanol for 10 minutes. To make this, add 0.02 ml of concentrated (12 M) hydrochloric acid to 100 ml of ethyl alcohol.

Reference:

Weir EE + 4 others (1974) Destruction of endogenous peroxidase activity in order to locate antigens by peroxidase-labeled antibodies. J. Histochem. Cytochem 22:51-54.

This simple method doesn't seem to be much used. I have tried it, and Yes, it did work.

John Kiernan
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Using mouse primary antibodies on mouse tissues

Question.

Using a mouse monoclonal on sections of mouse tissue often makes a strong background staining because the secondary antiserum binds to mouse immunoglobulin already present in the tissue. Is there a way to get round this difficulty?

Answer 1.

Two published methods seem quite good for this purpose. They are very briefly summarized below. For practical details consult the original papers:

1. Hierck,BP; Iperen,LV; Gittenberger-de Groot,AC; Poelmann,RE (1994): Modified indirect immunodetection allows study of murine tissue with mouse monoclonal antibodies. J. Histochem. Cytochem. 42(11, Nov), 1499-1502.
   Mouse monoclonal reacted with HRP-rabbit anti-(mouse serum); then add excess normal mouse serum & incubate with tissue.
2. Lu,QL; Partridge,TA (1998): A new blocking method for application of murine monoclonal antibody to mouse tissue sections. J. Histochem. Cytochem. 46, 977-983.
   Blocking with mixture of Fab and Fc fragments from rabbit anti-mouse antibody. (Made by papain digestion, then more Fc added). Stops background staining of endogenous mouse IgG by the secondary antiserum.

Corazon D. Bucana, Ph.D.
Houston, Texas
(bucana[AT]audumla.mdacc.tmc.edu)


Answer 2.

[ This answer does not really explain what to do, but the advertised product might interest users of mouse monoclonals. DAKO is now part of Agilent Technologies. ]

DAKO released an immunostaining system for animal tissues. In particular, it excels with mouse antibodies on mouse tissue. We engage a novel technology to ensure clean background and high specificity. Stoichiometric amounts of primary-antibody complex are preformed before it is exposed to the tissue site. This eliminates the unwanted reaction between secondary antibody and mouse tissue.

Please visit the Agilent website (former www.dakousa.com) to request literature on the new DAKO ARK (Animal Research Kit). We presented a poster at the IAP meeting in Boston and this document is available by mail.

A few highlights: 1. One kit for all animal IHC testing utilizing mouse monoclonal primary Abs. 2. Use on tissue from any animal species. 3. Unique process eliminates background staining. 4. Staining results in 45 minutes. 5. Automatable.

Bret Cook
Product Specialist, DAKO Corporation
(general[AT]silcom.com)

Back to Table of Contents

** Antigen retrieval: A patented or copyright phrase?

Two questions:

I was talking to someone the other day concerning immunoperoxidase staining and I mentioned the term "antigen retrieval". I was told that the term is patented and that it was not legal to use the phrase. Has anyone else heard that information. I do know that Biogenex makes and sells "Antigen Retrieval Solution," and we use it in our lab.

Is it really true that we cannot talk or write about antigen retrieval in a general way without the risk of being sued for some infringement of a copyright or a patent?

Answer.

This was the subject of some heated discussion in the HistoNet listserver in 1998. The following remarks are based on the contributions of people too numerous to acknowledge individually, and are colored by my own conclusions.

On the one hand there were the "common sense" viewpoints making the cases that:

(a) A combination of two common words could not possibly amount to an original literary composition (with copyright assignable to an author or publisher), and could never be construed as an invention. (A particular solution could, of course, be invented for the purpose of retrieving antigens, and patented.)
(b) Methods for enhancing the detection of antigens in sections have been published in the scientific literature for several years. All involve treatment with water, which may be cold or hot, and most techniques specify other substances to be dissolved in the water. The solutes include detergents (to damage cell membranes, helping large antibody molecules to enter cytoplasm), urea (disturbs protein conformation and may expose "buried" epitopes), a variety of metal salts, notably zinc sulfate and lead thiocyanate (probably work by changing the conformation of the antigen), and all sorts of buffers, mostly pH 5-6 or pH 8-9. (This probably catalyzes hydrolysis of the cross-links that formaldehyde makes between nearby parts of protein molecules. The optimum pH varies with different antigens. Heat accelerates the reaction, and can be conveniently delivered in a microwave oven.)

On the other hand (Would it be the Left or the Right?): 

People were using these methods daily, in routine procedures, sometimes with a proprietary solution and sometimes varying the technique to suit the antigen?  Feeling their freedom of expression (and perhaps also their livelihoods) threatened, they suggested alternatives to "antigen retrieval", most notably the abbreviation HIER (for "heat induced epitope retrieval").

The word "unmasking", which has a long and honorable history among histochemists, is a conspicuous improvement on "retrieval" because it says what happens. The epitopes of antigens are not retrieved (= brought back), because they were already there. The hot water and other chemicals make them accessible to the primary antibody by removing physical and chemical barriers ("masks") to the diffusion of large molecules.  The barriers usually result from the combination of formaldehyde fixation and paraffin embedding.

BUT people are human and by nature conservative (= change can only make things worse), so it's likely that "retrieve" will win out over "unmask" despite any logical arguments. 

The HistoNet discussions ended when Biogenex said that the firm did not claim exclusive ownership of the "antigen retrieval" word pair, and we could say or write it without being sued.  

John A. Kiernan, MB, ChB, PhD, DSc,
Department of Anatomy & Cell Biology,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** p53 protein

Question.

What is the significance of immunostaining with an antibody to p53?

Answer.

First of all, p53 is the antigen in the tissue, with which the antibody combines (The p is for "protein").  p53 is also sometimes referred to as a TSG - Tumour Suppressor Gene).  p53 was labelled "Molecule of the Year" by either Science or Nature in the 1990s.

The "wild" type p53 is the normal. It suppresses cell transformation and/or mutations. It was traditionally considered to have a very short life and was therefore never present in concentrations large enough to demonstrate immunocytochemically. "Mutant" type p53 has a longer "half-life" and is therefore more easily demonstrated. It used to be that mutant type p53 was the antigen of interest. Then of course, things got more complicated.

Now there are, of course, antibodies to each type of p53.  One thing is for sure: p53 is of fundamental importance in cell transformation. The biggest problem is that many consider that the expression of p53 is quantitatively related to prognosis and can therefore, be used to assess treatment outcomes. Whether quantitation should be by percentage of positive (?tumour) cells or by intensity of staining in the positive (?tumour) cells is still open to debate. Whichever it is, it is obviously important that your results of today can stand statistical comparison with your results of yesterday or tomorrow. Even more importantly, can they be used for comparisons with other labs? The patient may move elswhere for treatment, for example.

One thing I know for certain: it is very easy to make virtually all cells p53-positive - not just tumour cells - if you tweak your immunocytochemical method and any heat induced antigen retrieval you use. This is a real minefield!

Russ Allison, Cardiff, Wales
(Deceased, 2002)

Back to Table of Contents

** Prevention of fluorescence fading

Question.

What is available in the way of chemical additives to aqueous
mounting media, commercial or homemade, to suppress fading of
immunofluorescence preparations?

Answer 1.

Jules Elias has a discussion about this in his book "Immunohistopathology, A practical approach to diagnosis." ASCP Press, 1990. He says 1 percent p-phenylenediamine added to the mounting medium retards fading.

He gives two references:

Johnson, GD, et al, A Simple Method of Reducing the Fading of Immunofluorescece During Microscopy. J Immunol Methods 43:349-380, 1981.
Huff, JC, et.al., Enhancement of Specific Immunofluorescent Findings with use of para-phenylenediamine mounting buffer. J Invest Dermatol 78:49, 1982.

Tim Morken
San Francisco, CA
(
timothy.morken[AT]ucsf.edu
)


Answer 2.

Look into Vectashield, it is supposed to a good mounting media for immunofluorescence. You may not be able to prevent fading entirely, because the exciting light can cause it. Storage of the slides, after coverslipping, should be dark, sometimes in cold, or even in a freezer.

Vectashield is from Vector and it is pricey: $40 for 10 ml.

Gayle Callis
(
gayle.callis[AT]bresnan.net)

Answer 3.

The anti-fade agents that have already been mentioned are all good, I must admit I have never used Vectashield so will not comment on this. However, no mention has been made of the possible variability in results with these materials. Most of the anti-fade agents I have tried vary considerably in their effectiveness. This appears to depend on the specific antibody used, the fluorescent marker, the fluorescence ratio of dye to marker molecule, whether the IHC is direct or indirect and if you remembered to feed your cat before going to work. 

As an example: using lectin labelling of cells with direct or indirect techniques, I found that the FITC label was usually retained for UEA-1 but not for WGA. I would therefore urge anyone who is going to use anti-fade agents to try them first on some unimportant slides to test their effectiveness.

Barry Rittman
(barry.r.brittman[AT]uth.tmc.edu)

Back to Table of Contents

** Background in immunostained cartilage

Question.

I have tried to immunostain sections of whole mouse embryos with several primary antibodies to a nuclear epitope. I am getting nonspecific antibody staining in cytoplasm and in the connective tissue around the cartilage.

I have blocked with embryo powder, normal goat serum, normal horse serum, beat blocking solution from Zymed, and Fab fragments. What could be reacting with secondary alone?

Answer 1.

I do a lot of cartilage and bone IHC markers, mostly on rat, but have done some mouse tissue. Is your primary made in a mouse? Even with rat tissue, anti-mouse secondaries can combine non-specifically with the rat tissue, I put rat serum in my detection and it helps tremendously with the background.

Patsy Ruegg
(rueggp[AT]earthlink.net)

Answer 2.

The different blocking steps you have tested all block hydrophobic areas ("sticky sites") in your specimen. Hydrophobic areas are blocked before the immunoincubation with e.g. normal serum or BSA. Once blocked these sites generally will not give rise to background anymore.

Cartilage and perichondrium are composed of collagen fibers with a positive charge (still present after aldehyde fixation) embedded in proteoglycans which have a negative charge. Most antibodies (primaries and secondaries) are negatively charged at pH 7-8.2. I therfore think that the collagen fibers present in the cartilage tissue are causing your background problem. This charge-determined background can be circumvented by adding negatively charged molecules (e.g. aurion BSA-c) to the wash and incubation buffers. Another possible cause for background (a specific binding to proteoglycans) can be prevented by adding gelatin to your buffers. Do not put both BSA-c and gelatin in the same buffer, because they have charge-determined affinity
for each other as well.

I invite you to visit our web-site for detailed info on the topic above. http://www.aurion.nl

Peter van de Plas
AURION,
Wageningen, Netherlands
(vandeplas[AT]aurion.nl)

Back to Table of Contents

** Endogenous biotin in mast cells?

Question.

Do mast cells (MC) contain any endogenous biotin? They are often falsely positive in immunostaining methods that use avidin.

Answer 1.

Mast cells bind avidin nonspecifically because of ionic attraction between avidin (a basic protein) and heparin (acid polysaccharide in MC granules). This results in false positive staining by ABC. The cure is to use the ABC reagent at pH 9.4. 

For more information, see Bussolati, G & Gugliotta, P 1983. Nonspecific staining of mast cells by avidin-biotin-peroxidase complexes (ABC). J.  Histochem. Cytochem. 31: 1419-1421.

John A. Kiernan, Department of Anatomy & Cell Biology,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Answer 2.

Bussolati and Gugliotta (J. Histochem. Cytochem., 31(12): 1419-1421, 1983) described binding of ABC to mast cells.  They believed this to be due to both the binding of avidin basic residues as well as peroxidase to the sulphate groups of heparin. They showed that binding could be prevented by using the ABC solution at a pH of 9.4. This high pH does not affect either previous binding or localisation of antibody or the affinity of biotin for avidin.

They also showed that the nonspecific binding of avidin could be blocked by a 30 minute pretreatment of sections with a synthetic basic polypeptide such as poly-L-lysine (0.01% in PBS, pH 7.6).

Tony Henwood
Pathology Department, The Children's Hospital at Westmead
Westmead NSW 2145, AUSTRALIA

(tony.henwood[AT]health.nsw.gov.au)
 

Back to Table of Contents

 


 

MISCELLANEOUS STUFF

 

** Disposal of used diaminobenzidine (DAB) solutions

Question.

How should I dispose of used solutions of 3,3'-diaminobenzidine (DAB) that have been used for peroxidase histochemistry.

Answer 1.

While DAB itself has not been the subject of in-depth carcinogenicity studies, it is known to be mutagenic. Further, all members of the benzidine family that have been tested have been proved to be carcinogens. In the United States, at least, all benzidine derivatives are considered carcinogens by the NTP
(National Toxicology Program).

Many people collect the DAB solutions into a bottle containing 5% sodium hypochlorite (which is domestic bleach). After several hours, the DAB is oxidized to an insoluble polymer.

Chlorine bleach is NOT effective in removing the mutagenic properties of DAB. While it possibly may break the molecule down (reaction products are unidentified), introduction of chlorine into the end products simply produces another mutagenic chemical. This has been verified by Lunn & Sansone. Using chlorine bleach is neither chemically sensible nor effective. Fortunately, most if not all suppliers of DAB have eliminated this procedure of detoxification from package inserts and MSDSs.

There are two recommended methods of treatment. The most commonly used one currently involves potassium permanganate and sulfuric acid. End products are known to be non-mutagenic. The second uses horseradish peroxidase to form a solid which is readily isolated. The fluid remaining is non-mutagenic, but the precipitate retains its mutagenicity. The only purpose in performing this method is to reduce the volume of hazardous waste.

With any commercially available device purporting to detoxify hazardous chemicals, it is imperative that the user have documentation from the manufacturer that all reaction products have been properly tested and found to be non-hazardous. It is possible that some devices detoxify the liquid and filter out a hazardous solid. If so, the filter must be handled as a hazardous waste.

For further information, see:

NTP, 1998. National Toxicology Program Update (January 1998), Attachment 2. Available on-line at http://ntp-server.niehs.nih.gov
Lunn & Sansone, 1990. Destruction of hazardous chemicals in the laboratory. Wiley & Sons (pages 35-41)
Lunn & Sansone, 1991. The safe disposal of diaminobenzidine. Appl. Occup. Environ. Hyg. 6:49-53.
Dapson & Dapson, 1995. Hazardous materials in the histopathology laboratory: regulations, risks, handling and disposal. ANATECH LTD., Battle Creek, MI. (pages 25-27, 109-111 and 162-163)

Richard W. Dapson, Ph.D.
Formerly of ANATECH LTD.
Battle Creek, MI 49015
(dick[AT]dapsons.com)

Answer 2.

The procedure for acid permanganate oxidation of spent DAB is as follows. The measurements need not be very accurate. 

An acid permanganate solution is made by dissolving 4 g KMnO4 in 100 ml of dilute sulphuric acid (made by adding 15 ml conc. H2SO4 slowly and carefully to 85 ml of water). This solution is stable. (My experience is that it's very good at cementing in place the glass stoppers or screw caps of bottles containing it.)

Add the solution for disposal to an excess of acidified permanganate and leave overnight (in a fume hood if the solution contained chloride ions, because these will end up as chlorine). Next day, neutralize with sodium hydroxide (carefully; the temperature will rise) and filter. Leave the filter paper to dry in
the funnel, then put it in a plastic bag for disposal.

If you have a large volume of DAB solution, carefully add sulphuric acid (150 ml for each litre) and then dissolve solid potassium permanganate (40 g for each litre).

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

John A. Kiernan,
Department of Anatomy & Cell Biology,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Dilution of concentrated acids: formula etc.

Question.

If I want to make a 1N solution of, for example, hydrochloric acid how do I convert the liquid, concentrated HCl into a gram value. The bottle of concentrated HCl says it is a 35-36% solution.

Answer.

This applies to dilution of all concentrated acids (and also to strong ammonia (ammonium hydroxide) solutions.

The percentage on the label is weight/weight, not weight/volume, so you have to take into account the density of the concentrated acid.

The formula for making one litre of a particular normality, N, is:

where V is the volume of concentrated acid needed, M is is its molecular weight, N is the desired normality, B is the basicity (1 for most common acids; 2 for sulphuric; 3 for phosphoric; 1 for ammonia), P is the percentage by weight in the concentrated acid - the figure on the label, and D is the density of the conc. acid (specific gravity) in grams per ml.

No, I didn't work it out myself; it's from Lange's Handbook of Chemistry.

If the dilution doesn't need to be very precise, you can assume the following normalities for common concentrated acids:

Hydrochloric (36%) 12N
Nitric (71%) 16N
Sulphuric (96%) 36N (= 18M)
Acetic (99%+) 17.4N
Formic (90%) 23.4N

So to make approximately 0.5N hydrochloric acid, you dilute the conc. HCl 24 times. To make a litre, you'd measure 42 ml of the conc. acid (because 1000/24=41.7) and add it to about 800 ml of water. Stir, and make up to a final volume of 1000 ml.

Remember to pour the acid slowly into the water, especially sulphuric acid, which generates a lot of heat when mixed with water.

John A. Kiernan,
Department of Anatomy & Cell Biology,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Disposal of waste from "special stains."

Question.

How should I safely dispose of the waste chemicals generated in a variety of special staining porcedures?

There is no consensus here, especially about the use of "copious running water" for dilution. A sample of collected opinions follows.

Answer 1.

Identify the substances that are dangerous in quite small amounts, such as mercuric chloride or sodium cacodylate, and follow your institution's guidelines for collection and disposal. Most substances used in special stains (dyes, acetic acid etc) can be flushed down the sink with plenty
of running water.

John A. Kiernan
London, Canada.

Answer 2.

There are disposal practices that are forbidden for "Industrial" users that are allowed for "Educational" users.

The last time (some years ago) I took a Hazardous Waste Disposal course, I found out that Industry has strict regulations on e.g. Osmium tetroxide disposal, but it was *recommended* that university labs dump it down the sink. This was allowed, as long as the Os concentration didn't exceed some specified level at the sewage treatment plant. Storing the Os for disposal (even using corn oil and kitty litter) was more likely to result in legal troubles because of laws on how waste must be stored, for how long, and whether at a "local" site (your lab) or a central collection site, etc.

Hazardous waste laws change frequently.

Philip Oshel
(oshel[AT]terracom.net)

Answer 3.

Here is a brief synopsis of advice appropriate for the USA, and to a great extent, Canada. Further details can be found in our book, Hazardous Chemicals in the
Histopathology Laboratory, 3rd ed.

First and foremost, never mix different wastes together unless directed to do so by a licensed waste hauler, or until you have determined that it is safe and proper to do so. Why? You could easily create something far more hazardous. You might be mixing a low-hazard solution that could go down the drain with a high-hazard solution that could only be hauled away; that creates a far larger volume of high-hazard material that you have to pay to get rid of. A good example would be mixing mercury waste from B-5 or de-Zenkerization with a trichrome solution. Remember, too, that alcoholic waste is burnable and thus less expensive to haul away than aqueous waste. Don't dilute alcoholic waste with a lot of aqueous waste, or you will be billed at the aqueous price.

Second, ALWAYS contact your local wastewater authority for advice. In many cases, they can assist in determining disposal procedures, particularly in those communities with proactive outreach programs. Have information ready for them: type of waste (flammable, toxic, etc.), components (don't say Mallory's trichrome, rather list the ingredients), volume and how often. Include MSDS's. Every community has its own unique set of limits for certain chemicals. Chromium, silver and mercury are stringently regulated, so keep those wastes separate from others.

Third, use common sense. Stain waste that does not contain heavy metals, and is of small volume (few hundred ml) is so insignificant that in most sewer districts it can be trickled down the drain. NEVER pour waste down the drain if silver, chromium or mercury is present. This includes rinses following those solutions in the staining program.

Do not pour waste down the drain all at once. Trickle it from a small carboy outfitted with bottom spigot. Never use "copious amounts of water" to flush waste; it is against EPA regulations anywhere in the United States.

Finally, use what others are doing as a guide only. They may or may not have opted for legitimate means of disposal, and even then, their constraints or lack thereof almost certainly will not pertain to you unless you are in the same community.

Richard W. Dapson
Richland, MI
(dick[AT]dapsons.com)

Answer 4.

I have to ask why using copius amounts of water is bad when disposing of waste. I can understand arguments about wasting water, but that would preclude putting solutions down the drain in the first place. So, if you are allowed to put something down the drain, I would think the volume would be beneficial for dilution.

Tim Morken

San Francisco, CA
(
timothy.morken[AT]ucsf.edu
)

Back to Table of Contents

** Magnification of a photomicrograph

Question.

I'm trying to find the calculation used to determine the magnification of a photomicrograph. I know you have to take into consideration several things besides the objective.
Can someone help?

Answer 1.

There are a couple of "gotchas" in figuring magnification. You need the magnification of the objective multiplied by the magnification of the ocular. However, and here is where you need to do some double checking, be sure the ocular in the path to the camera is the magnification you use. On some microscope/camera combinations, a different magnification is used for the camera ocular.

Then there is the matter of whether the microscope has a "tube lens." If the microscope you used is not one of the newest infinity corrected types, then there is most likely a
magnification lens BETWEEN the objective and the ocular. These generally fall into the magnification range of 1.5×, which again would have to be multiplied with the other two magnifications. On some microscopes, the tube lens magnification is marked on a surface betwen the objectives and the oculars, but on others, theres is no external marking. In that case, you will need an original manual for the scope. To complicate matters even further, many camera connect to the microscope trinocular tube with a reduction tube. So the magnification the camera sees is the combination of the various lenses used, divided by the reduction tube. The reduction tubes commonly fall into the range of 0.25× to 0.75×. The reduction factor is generally printed on the outside of the tube that connects the camera to the microscope.

As a general procedure, for any microscope used to take photomicrographs, one should take a picture of a stage micrometer with each objective on the scope, and keep these
pictures in a "calibration" file for that camera/microscope combination. The stage micrometer will be a true "ruler" with divisions of 0.1 and 0.01 mm, so it is easy to check the true magnification of prints or slides. If you don't have a stage micrometer, then use the built in standard: the average diameter of red blood cells after most processing procedures is approximately 7 microns. That is not exact, but is a good way to check that your magnification calculations are in the right ballpark.

Alton D. Floyd, Ph.D.
(Deceased)

Answer 2.

The best way is to photograph a calibrated slide using the same objective and other variable things as for the section. Print the photos at the same enlargement, and measure with a ruler. If a 100 micrometre distance is 32 mm on the print, the magnification is 32000/100 = 320.

Calculations based on the optics commonly lead to ridiculous mistakes. As a rough check, measure something in the photo and see if it's a sensible size. If there are cell nuclei 50 micrometres across, somebody has made an arithmetic error. Erroneous magnifications are often present in the legends of published micrographs.

John A. Kiernan,
Department of Anatomy & Cell Biology,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Can a method be both published and patented?

Questions.

The tyramide amplification system (for showing peroxidase activity at sites of antibody binding or in situ nucleic acid hybridization) is sold commercially in patented kits. The principal reagent (tyramine coupled to biotin or various fluorescent compounds) can be synthesized in the laboratory, following quite simple techniques published in the Journal of Histochemistry and Cytochemistry, and elsewhere. Is there a risk of being sued by the firm that sells the kits, for following a published method to make  a reagent in one's own lab?

Answer.

[ There was some rather heated discussion on the HistoNet listserver in 1998, involving various individuals and one of the patent holders. It centered around the unavailability of individual reagents and a claim that a company might even sue individuals for daring to encourage others to carry out the published syntheses. ]The following message was from Mark Bobrow, an author of some of the published procedures and also one of patent holders. ]

The patent system goes back over five hundred years when, in Britain, one could obtain a patent granted by the King. In the U.S., the first patent commission was headed by George Washington, who personally signed every patent granted during his tenure.

A patent is a right granted by the government. Article I, Section 8 of the United States Constitution states, "The Congress shall have the power to promote the progress of science and useful arts, by securing for limited times to authors and inventors the exclusive right to their respective writings and discoveries."

It is often misunderstood that the purpose of the patent system is, as stated in the Constitution, *to promote the progress of science and useful arts.* The concept is that by disclosing (and not keeping a secret) an invention, technological innovation will continue. In the process of obtaining a patent, the inventor must disclose the invention and the best mode of practising it (in other words, they can't hold anything back, or the patent will not be valid).

In return for disclosing the invention, the government grants the patent holder the right to exclude others from making, using, or selling the invention. Currently, these rights extend for 20 years from the filing date. After the term expires, everyone is free to make, use or sell the product or method which was disclosed in the patent

The right to exclude others from practising the invention applies to everyone, including academic investigators. In terms of being able to use what is in the published literature, U.S. patents are published after they issue; in Europe the applications are published 18 months after filing. So, even though patented products and methods are in the published literature, using them without proper authorization from the patent holder is not legal.

There have been some questions as to the extent of coverage of the tyramide amplification patents. In the spirit of simplification, four basic concepts are claimed. They are the enzyme substrates (e.g., tyramides), the product of the enzyme-substrate reaction, the method of catalyzed reporter deposition (e.g., detecting an analyte with a reporter enzyme using the deposition of a reporter), and assays using the method of catalyzed reporter deposition. If you wish, you may look it up yourselves. One of the patents is U.S. Patent 5,731,158, Catalyzed Reporter Deposition. As an added note, the readers should be reminded that patents are written in a style that is a hybrid of law and science (perhaps a suspension is more descriptive).

Patent information is available on the internet. Here is a list of some sites:

http://www.uspto.gov/. This is the US Patent Office site. You can search for patents here, and get some information about patents in general. Later this year, or early next year, the full text and images of patents will be available.

http://patent.womplex.ibm.com/searchhelp.html. This is an IBM site where one can search for patents and view the entire document (it tends to be slow though).

http://www-sul.stanford.edu/depts/swain/patent/patgeninf.html. General patent information.

[ End of reported communication from M. Bobrow ]

Back to Table of Contents

** Books and articles about artifacts in histology

Question.

Can you recommend any books or articles that illustrate and explain artifacts encountered in sections stained for light microscopy?

Answers.

"An Atlas of Artifacts Encountered in the Preparation of Microscopic Tissue Sections" by Samuel Wesley Thompson and Lee G. Luna. Publisher: Charles C Thomas, Springfield, Illinois, U.S.A. (1978).

There is also a wonderful section on Artifacts (and photographs) in "Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts" by Lee G. Luna, 1992, printed by Johnson Printers, Downers Grove, IL.

Marilyn S. Gamble
(Marilyn.S.Gamble[AT]kp.ORG)

I agree with the value of Lee Luna's book "Histopathological Methods and Color Atlas of special stains and tissue artifacts," especially the value of the colour photomicrographs.

The most comprehensive paper I have seen is: Wallington EA. "Artifacts in tissue sections" Medical Laboratory Science. 1979;36:1-61 (that's right, sixty one pages!) It is the paper which won the Memorial Prize of our institute - Institute of Biomedical Science. In those days, unfortunately, published photos were in B&W only, but there is plenty of text and explanation. Eric was a real gent, a master of histological technique and perhaps the greatest authority on artifacts. 

Russ Allison, Wales
(Deceased)

"Histologic Preparations. Common Problems and Their Solutions" edited by Richard W. Brown. Northfield, IL: American College of Pathologists (2009) (156 pages).

The former web site of Roy Ellis had many informative images of artifacts, with quizzes and explanations. Highly recommended! It is now hosted by IHC World: http://www.ihcworld.com/royellis/gallery/mainpage.htm.

John Kiernan, London, Canada
(kiernan[AT]uwo.ca)

Back to Table of Contents

** How dangerous is picric acid?

Question.

Older colleagues tell of picric acid exploding with great violence, but always in other labs. Is there really a risk of explosion?

Answer.

From the late 19th Century until the First World War, picric acid was used as a high explosive in military shells. Its melting point (122C) is quite well separated from its
exploding temperature (above 300C). Picric acid can be ignited by a nearby spark at temperatures above its flash point of 150C. More sensitive explosives can be formed by chemical reaction of picric acid with other substances. An example is ammonium picrate (which has been used in histology to fix vital stainings with methylene blue).

In December 1917 a French freighter, the Mont Blanc, full of expired explosives, caught fire in the harbour of Halifax, Nova Scotia. The largest man-made, non-nuclear explosion followed, and it's customary to blame it on picric acid, which probably accounted for much of the cargo. (Click here to read more about the Halifax Explosion.)

When you buy a bottle of picric acid for the lab, the yellow powder is mixed with 10% to 40% of its weight of water (varies with the supplier), so it is impossible for the temperature to go above 100C, let alone the 300C required for an explosion. If a jar of picric acid were to dry out, as a result of neglect, it's conceivable that a high temperature might develop from friction when unscrewing a tight bottle cap, but 300C seems highly improbable. Nevertheless, it's usual to loosen a tight cap by standing the jar upside down in water for a few minutes before applying force to it. Percussion can cause a locally high temperature, so you shouldn't hit dry picric acid with a hammer. One of its uses is in matches. Stories of picric acid explosions in labs are like sitings of ghosts: always second-or third-hand.

Various toxic effects are described, especially skin reactions. Oral LD50 values range from 60 to 250 mg/kg depending on the animal. (This puts it in the same league as ferrous sulphate.)

Sources: Various chemistry textbooks; Merck Index; Lange's Handbook of Chemistry; MSDS sheet.

John A. Kiernan,
Department of Anatomy & Cell Biology,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)

Back to Table of Contents

** Which color print film for photomicrography?

Question.

What brand of color 35mm film and ASA (film speed) is best suited for photographing H & E sections? I would like to produce prints, not projection slides.

[Older microscopes often have built-in cameras requiring 35 mm film, so this question still has some relevance in this age of digital photography.] 

Answer.

Fuji is good. Use the slowest speed, (lowest ASA) you can: ASA 25 or 100, for the best results.

If there is much vibration where your camera is, you may need to go to a faster film to shorten your exposure times.

Use professional film, not consumer. The difference is that pro film is refrigerated after it's made, so there is no color shift with aging. Keep used film in your lab refrigerator for this reason.

You don't have to worry much about daylight vs tungsten film because you're shooting negatives and not transparencies. If your photomicroscopy set up controls color temperature, then try to shoot at 5500K (5500 deg), because color film likes sunlight. Use neutral density filters to lower light levels if needed.

Also: who's doing your printing? A film lab or someone used to histo shots? If it's a film lab, then they won't know how to balance the color of your sections, and you're likely to get weird results. If your camera back comes off the scope, take the first one or two shots of a Caucasian person outdoors, sun behind the camera. The automated developing and printing machines are set to correctly balance Caucasian skin tones, and should keep this setting for the rest of the roll. If your camera cannot come off of the scope, then when you send your film to be printed, include an image of an H & E section with correct color balance. This will give the photo lab a reference to use for balancing the colors of your film when printing.

Phil Oshel
Middleton, WI 53562
(oshel[AT]terracom.net)

Back to Table of Contents


     [ End of FAQ document ] 

Biological Stain Commission Home Page

----------------------------
Last updated: October 2019

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

Note on revisions in Version 2.0

The previous version of this collection of questions and answers was numbered 1.6, published in September 2014 at http://publish.uwo.ca/~jkiernan/faqlist.htm. The same information, but differently formatted, appeared on the BSC's WordPress web site (https://biologicalstaincommission.org/faqs/ - same address as this document)  from 2015 until October 2019.  

Revisions in Version 2.0:
I continue to collect and edit new questions and answers, as well as new comments relating to items already in this FAQ.

John   A. Kiernan
Secretary,  Biological Stain Commission
= = =