FAQs

Introduction

The questions and answers are grouped into six categories:

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 send an email to one of these addresses remember to edit the address 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 screenfulls 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 the 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: <J.A.Kiernan>kiernan[AT]uwo.ca

Questions and answers in print

Since January 2001 the journal Biotechnic&Histochemistry has carried an occasional feature called Notes and Queries consisting of anonymous
questions with signed, peer-reviewed answers. For controversial issues there may be more than one answer. The peer review process means that a Note responding to a Query in B&H is a statement made in a journal that has had strict editorial standards for more than75years. (The name of the journal was changed from Stain Technology to B&H in 1991.)

Biotechnic&Histochemistry is the journal of the Biological Stain Commission. Click this link: BiologicalStainCommission to learn about the B.S.C.-what it is and what it does.

Acknowledgements

I thank the many people who have answered my questions about staining and related methodology, and also those who have asked my advice and made me think and investigate. In recent years the HistoNet listserver has been a 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 at kiernan[AT]uwo.ca. I will immediately expunge the offending Answer and try to find or write another to replace it. If you want to revise something attributed to you, let me know and will incorporate the change.


Table of Contents.
FIXATION, FREEZING ETC

PROCESSING, DECALCIFYING, EMBEDDING

SECTIONING, SLIDE ADHESIVES, MOUNTING

STAINING METHODS, HISTOCHEMISTRY

IMMUNOHISTOCHEMISTRY

MISCELLANEOUS STUFF

FIXATION, FREEZING ETC

** Carbodiimides as fixatives

Question.

Does anyone know what carbodiimide is and how it works?

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.

Barry Rittman
brittman[AT]mail.db.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:

10% NBF
70% EtOH
70% MeOH
Carnoy’s
methacarn (Carnoy’s with methanol instead of ethanol)
zinc formalin, unbuffered
buffered zinc formalin

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 1999. Histological and histochemical Methods, 3rd
ed. Oxford: Butterworth-Heinemann.
Kok LP & Boon ME 1992. Microwave Cookbook for Microscopists.
Leiden: Coulomb Press.
Pearse AGE 1980. Histochemistry, 4th ed. Vol 1.

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:

GlyoFix from Shandon Lipshaw uses glyoxal as the active
ingredient; produces aldehyde-type fixation patterns.

Histochoice from Amresco; active ingredients essentially
undisclosed (aldehydic addition compounds); mode of action
unknown.

HistoFix, formerly from Trend Scientific, perhaps still
available from Baxter, contains pyrrolid-2-one, a polyol,
a urea and a zinc salt; mode of action unknown.

Mirsky’s Fixative from National Diagnostics is an aqueous
solution of a complex di-aldehyde (possible di-aldehyde
starch); mode of action may be aldehyde-like, but very slow.

NoToX from EarthSafe Industries, uses a complex aldehyde
(possibly di-aldehyde glucose) in about 70% alcohol with
antiseptic and antifungal agents; produces a combination of
aldehyde- and alcohol-type fixation patterns.

OmniFix II and OmniFix 2000 from AnCon Genetics is an
alcohol-based solution containing glycol and salts;
produces alcohol-type fixation patterns.

Prefer from Anatech Ltd., uses glyoxal as the active
ingredient; produces aldehyde-type fixation patterns.

SafeFix II from CMS uses glyoxal as the active ingredient;
produces aldehyde-type fixation patterns.

STF (Streck Tissue Fixative) from Streck Laboratories contains
diazolidinyl urea, 2-bromo-2-nitropropane-1,3 diol, zinc
sulfate and a small amount of formaldehyde as active
ingredients; mode of action unknown.

There are two fixatives intended for microwave use:

Preserve from Energy Beam Sciences uses glyoxal as the active
ingredient; produces aldehyde-type fixation patterns.

MicroFix from Energy Beam Sciences is an alcohol/polyethylene
glycol solution. It replaces Merck’s KryoFix, which is no
longer available; produces alcohol-type fixation patterns.

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, Ph.D.
Anatech Ltd.
Battle Creek, MI 49015
(anatech[AT]net-link.net)

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
(brittman[AT]mail.db.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.):

Lidocaine 50 mg per mL, dissolved 95% EtOH (warm to dissolve).
Add slowly to perfusion solution with stirring to make final
lidocaine solution concentration wanted (e.g. 1mg/mL = 0.1%)

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 for the manual … (and I would like to
read about it just for my own knowledge).

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. (Cost about $50/25 g in 1994.)

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. 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 (1972).

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. Calibrating and
standardizing microwave ovens for microwave-accelerated specimen
preparation. A review. Cell Vision 3: 172-179, 1996.
2. Login, G. R., and A. M. Dvorak. The Microwave Toolbook. A Practical
Guide for Microscopists. Boston: Beth Israel Hospital, 1994.
3. Login, G. R., J. B. Leonard, and A. M. Dvorak. Calibration and
standardization of microwave ovens for fixation of brain and
peripheral nerve tissue. Companion to Methods Enzymol 15: (in
press), 1998.
4. Login, G. R. The need for clinical laboratory standards for
microwave-accelerated procedures. J Histotechnol 21: 1-3, 1998
(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-CH2-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 60 C. 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 can have the phosphate
buffer component replaced by seawater with no problems as 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.
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? Thanks in advance!

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.

From 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 rarely
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 (2/19/98) 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.
ANATECH LTD.
1020 Harts Lake Road
Battle Creek, MI 49015
(anatech[AT]net-link.net)

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, 2 years
after it’s introduction as a fixative) and described other
fixative mixtures.]

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

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** 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)

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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 alcohol 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 microwave processing, isopropyl alcohol is
sometimes recommended. However, this does not mix
with wax. It has to leave the specimen by vaporizing
(boiling) under reduced pressure. This can lead to
considerable tissue damage unless the temperature
and pressure are just right (Bosch et al 1996).
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.
Kiernan,JA (1996): Staining paraffin sections without prior
removal of the wax. Biotechnic & Histochemistry 71(6),
304-310.

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 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)

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** 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 (May 1997):

Fisher Scientific 4 litres $104 (“Laboratory grade”)
Sigma 3 kg $42 (no purity details)

Acros Organics (seems to be part of Fisher)
sell three grades:
2.5 litres $24 (99%)
1 kg $23 (GC)
500 ml $36 (scintillation grade)

If the 99% stuff is OK for histology, perhaps the price isn’t
too bad. tert-butanol (99.5%; from Acros) is $67 for 2.5 litres,
and 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)

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** 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
(uvsgc[AT]msu.oscs.montana.edu)

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 Ind. Univ. School of Dentistry is as
follows:

The protocol we use here at Ind. 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)

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** 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 2.

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. 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)

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** 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. Kiernan. ]

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)

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** 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.

This is my processing schedule 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.

Tissue Processor.

These times I use on a Shandon Histokinette, remember
them ?.

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)

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** Cryoprotection of specimens

Question.

Please recommend a way to protect formaldehyde-fixed
mouse brains to avoid crack and ice crystal holes
that for 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).

Tokuyasu KT. 1989. Use of polyvinylpyrrolidine and polyvinylalchohol
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 leased 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)

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** Cutting sections of toe or finger nails

Question.

Does anyone have a few hints for sectoning toenails?

[ Here is a selection 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 58 C (it liquefies) for several hours
during the day on a particularly tough nail; it cut beautifully
the next day!

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 know 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
(allison[AT]cardiff.ac.uk)

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** 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)

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** 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 plastic
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 spec 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 some 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. Fisher/VWR I think carry
them.

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

[Pre-prepared silanized slides are commercially
available with a variety of trade names.]

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 chrom 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 P.S./$ 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 is a waste of money to buy
poly-L-lysine rather than poly-D-lysine or poly-DL-lysine,
because the stereochemical form of the amino acid does not affect
its ionization. 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 someone else’s employer 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 would
be very straightforward to make 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, add 30 to 50 grm of potassium acetate
or 10 grm of sodium chloride.

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

(With additions and minor editing by J. A. Kiernan, London, Canada).

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 æm 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 æm 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 æm 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 æm
alumina slurry after the 1 æm.

I have tried progressive alumina slurries, 3 æm then to 1 æm, but
it was a waste of time, 1 æm 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 æm
600 grit = 14 æm
800 grit = 10 æm
1000 grit = 5 æm

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
(uvsgc[AT]msu.oscs.montana.edu)

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 76 C. 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
60 C nor 90 C 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)?

A few 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

** 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.

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
tungsten carbide 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

Question.

[ 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. ]

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
Atlanta, GA
(timcdc[AT]hotmail.com)

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

Back to Table of Contents

** Sectioning plant material: some hints.

Question(s).

Does anyone know how to make nice sections of plant material?
I tried to make paraffin sections but it seems that the thick
cellulose walls of the cells are 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 evaporation, after immersing in 10% glycerol. 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.

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. This always included sectioning an
alcohol-fixed 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 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 h’tox, 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.
Jensen, WA (1962): Botanical Histochemistry. Freeman, San
Francisco.
Kiernan, JA (1999): Histological and Histochemical Methods.
Theory and Practice. 3rd ed. Butterworth-Heinemann, Oxford.
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, immunohistochem, Na, Ca,
K in Vol 2.

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

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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.

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 cost 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 21 C. The bottle must be cold when you open
it!

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

Gayle Callis
(uvsgc[AT]msu.oscs.montana.edu)

[ With some editing and additional comments by J. A. Kiernan ]

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** 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 æm
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)

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** 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)

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** 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 Lillie’s “Histopathological Technic
and Practical Histochemistry”)

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

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** 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 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.

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

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

Reference: “Biological stains and staining methods.” BDH
leaflet, 1966.
Several modifications of Castaneda’s original technique are
given in: Langeron, M.:”Precis de Microscopie”, 1934 and 1948.

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

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** 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.

This question to the HistoNet listserver elicited
replies that generally favored the “rhodanine”
reagent over “rubeanic acid.” Nomenclature can be
confusing! Don’t confuse rhodaNine with rhodaMine,
and note that in any chemical catalog, p-dimethyl-
is indexed under the letter D, not P. A few
general references for copper histochemistry are
added at the end of this FAQ item.
J.A. Kiernan.

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:

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-Dimethylaminobenzylinene-rhodanine 0.2 g
Absolute ethanol 100 ml

Rhodanine solution (working) –

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

Diluted Mayer’s hematoxylin

Mayer’s hematoxylin 50 ml
Distilled water 50 ml

0.5% aqueous sodium borate (borax)

Note: The use of chemically clean glassware is necessary.
Shake stock solution before measuring and mixing
solutions and shake the working solution before
pouring it onto the slides.

Technic:

1. Hydrate slides to distilled water.
2. Incubate slides in rhodanine working solution
at 37 degree C 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% alcohol to absolute ethanol,
clear, and coverslip with a synthetic 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.

Irons,RD; Schenk,EA; Lee,CK (1977): Cytochemical methods for
copper. Archives of Pathology and Laboratory Medicine 101,
298-301.
Cytochem methods for copper. Comparison of dithiooxamide,
diaminobenzylidene-rhodanine, diethylthiocarbamate.
Pearse, AGE (1985) Histochemistry, Theoretical and Applied,
4th ed. Vol. 2.
Metal histochemistry is extensively reviewed in Chapter 20.
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 histoch 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)

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** 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)

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** 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 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. Acidic
reagents precipitate the insoluble colour-acid.

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
(allison[AT]cardiff.ac.uk)

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** 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)

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** 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)

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** 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:

* KMnO4: 177 mg
* KClO3: 114 mg
* KIO3: 200 mg
* NaIO3: 197 mg
* KCr2O7: 276 mg

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 æm paraffin sections, fixation: Bouin; manual
staining.)

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

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

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** McFaydean'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:

Methylene blue 0.5 g
1% w/v Aq. KOH 1.0 ml
Ethanol 30.0 ml
D.water 70.0 ml
Warm the water to 50C., stir in methylene blue and
add other ingredients, cool and filter before use.

Polychrome methylene blue (Unna):

Methylene blue 1.0 g
Pot. carbonate 1.0 g
Ethanol 20.0 ml
D.water 100 ml
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.

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
McFadyean, 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 methchromasia, 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)

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** 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
(brittman[AT]mail.db.uth.tmc.edu)

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** 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 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 isn’t one of those certified by the Biological Stain
Commission, and some major American vendors do not have it in
their catalogues. The stuff in my lab was bought from BDH (Gurr)
about 15 years ago. 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.

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

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** 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 parts?

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)

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** 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)

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** 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 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

10 ml 95% ethyl alcohol
0.3 gm Basic fuchsine
6 ml Liquid Phenol
94 ml Distilled water
Combine in the listed order.

10% Sulfuric Acid

10 ml Sulfuric acid
90 ml Distilled water

5% Malachite Green

95 ml Distilled water
5 gm 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)

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** 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 in August 1998. 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.

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** 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
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 most 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 GILL

Hematoxylin 1 g Hematoxylin 2 g
Potassium alum 50 g (0.09M) Al sulfate 17.6 g (0.03M)
Sodium iodate 0.2 g Sodium iodate 0.2 g
Citric acid 1 g Acetic acid 40 ml
Chloral hydrate 50 g Ethylene glycol 250 ml
Water to make 1000 ml Water to make 1000 ml

Molar ratio of Al ions to haematein molecules in
the freshly made solution:
Mayer: 32
Gill: 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 f�r wissenschaftliche Mikroskopie 3: 150.
Kiernan, J.A. (1999). Histological and Histochemical Methods:
Theory and Practice, 3rd ed. Oxford: Butterworth-Heinemann.
Llewellyn, Bryan. Stains File. http://www.netbistro.com/~bryand/
(This Web site has a splendid, possibly comprehensive,
collection of hematoxylin stain formulations.)

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

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** 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:

1. Sulfate (actually half-sulfate) of many carbohydrate
components (glycoproteins in some mucus, heparin in mast
cell granules, chondroitin sulfates in cartilage matrix,
etc.) These are strong acids: ionized even at low pH.
Sulphate groups therefore bind cationic (basic) dyes
at any pH. They are the only things stained at pH 1.
2. Phosphate groups, associated with DNA and RNA. These are
weak acids, so they become protonated (not ionized) if
the concentration of protons (hydrogen ions) is high
enough. Typically this occurs below about pH 2.5. The
phosphates of nucleic acids are fully ionized at pH 3.5
to 4. A basic dye at pH 3 to 4 stains nuclei, cytoplasm
that is rich in RNA and. of course, all the sites of
half-sulfate esters.
3. Carboxyl groups. These occur as parts of amino acids
(C-terminal and the side-chains of glutamic and aspartic
acid), sialic acids (mucus and other glycoproteins),
glycosaminoglycans of extracellular matrix carbohydrates
(hyaluronic acid, chondroitin sulfates etc) and free
fatty acids (frozen sections only). Carboxyl groups
ionize over quite a wide range of pH, from 5 up to
about 8. The higher the pH, the stronger and more rapid
the staining by a basic dye. At or above pH 8 it stains
everything.

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. (1999). Histological and Histochemical
Methods: Theory and Practice, 3rd ed. Oxford:
Butterworth-Heinemann.
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)

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** 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.

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

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** 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 RT) 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 buffer will vary form 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

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** 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)

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** 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)

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** 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.

1. Bring paraffin sections to water in usual manner.
2. Stain sections in acridine orange stain for 30 minutes.
3. Rinse sections briefly in 0.5% acetic acid in 100% alcohol.
4. Rinse sections in two additional changes of 100% alcohol.
5. Rinse sections in two changes of xylene.
6. 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.

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 the acridine orange dye
work better than others.)

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

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** 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
(brittman[AT]mail.db.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
Centers for Disease Control
Atlanta
(timcdc[AT]hotmail.com)

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)

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** 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 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)

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** 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.

Reference: Conn’s Biological Stains. 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 are 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 also, and if there are capillary
gaps involved, this does not produce the crummy bubbles that
drive one crazy.

Gayle Callis
(uvsgc[AT]msu.oscs.montana.edu)

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 methonal 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 poly lysine 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
(uvsgc[AT]msu.oscs.montana.edu)

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(s) 1.

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

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

Answer 2.

[ This answer does not really explain what to do, but the
advertised product might interest users of mouse monoclonals.]

DAKO just 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 DAKO Corporation website (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

For more information please contact DAKO Technical Services at
techserv[AT]dakousa.com or call 800-424-0021.

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

Back to Table of Contents

** Antigen retrieval: A patented or copyright phrase?

Question:

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 case 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?)
were people 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.”

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 were not retrieved (= brought back), because
they were already there. The hot water and other chemicals
made them accessible to the primary antibody by removing
physical and chemical barriers (“masks”) to the diffusion
of large molecules.

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
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 about three years ago.

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.

There are, of course, antibodies to each type of p53 now.
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. A real minefield!

Russ Allison, Wales
(Allison[AT]cardiff.ac.uk)

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.

Two references he gives:

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
(timcdc[AT]hotmail.com)

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
(uvsgc[AT]msu.oscs.montana.edu)

Answer 3.

I think that 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 extra slides to test their
effectiveness.

Barry Rittman
(brittman[AT]mail.db.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 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, Senior Scientist
Anatomical Pathology
Royal Prince Alfred Hospital
Sydney, AUSTRALIA
http://www2.one.net.au/~henwood
http://www.pathsearch.com/homepages/TonyHenwood/default.html
(henwood[AT]mail.one.net.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
MSDS’s.

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.
ANATECH LTD.
Battle Creek, MI 49015
(anatech[AT]net-link.net)

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:
100 X M X N
V = ————
B X P X D
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)

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** 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 the
opinions stated in replies to the HistoNet listserver
in the Summer of 1998 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
ANATECH LTD
Battle Creek, MI 49015
(anatech[AT]net-link.net)

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,
Centers for Disease Control
Atlanta, GA 30333, USA
(timcdc[AT]hotmail.com)

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** 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 x, 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 x. 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.
ImagePath Systems,
(al.floyd[AT]juno.com)

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.

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

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** 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 August 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. ]

Linda Margraf relayed this to HistoNet. It was from Mark
Bobrow. He is an author of some of the published procedures
and also one of patent holders.

[ Beginning of M. Bobrow’s message ]

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 ]

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** 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.
Please don’t ask me to send a photocopy!

Russ Allison, Wales
(Allison[AT]cardiff.ac.uk)

The web site of Roy Ellis has many informative images of
artifacts, with quizzes and explanations. Highly recommended!
http://home.primus.com.au/royellis

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

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** 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 1915 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.

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)

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** 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.

Answer.

Fuji or Kodak, use the slowest speed, lowest ASA you can. ASA 25
is good, 100 will produce good 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.
Middleton, WI 53562
(oshel[AT]terracom.net)

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Last updated: September 2014