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 ]

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** McFaydean’s stain for anthrax bacilli

Question.

What is M’Faydean’s stain?

Answer.

[ 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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.

** 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)

** 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)

** 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)

** 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

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

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