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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)