** Sections coming off slides. Which adhesive?
Here is my problem: tissue sections not adhering to the slides.
Any hints on solving this problem?
[ 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
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
[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
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.
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”
Surgipath Medical Industries, Inc.
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.
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,
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.
** Apathy’s mounting medium and variants
Where can I find the recipe to make von Apathy’s mounting
medium? Is there more than one way to make it?
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
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
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
Ian Montgomery, Glasgow, Scotland
** Silanized (APES or TES or positively charged) slides
How do I prepare charged or silanized slides in the
lab, and is it OK to use metal slide racks?
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
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.
University of Nottingham
(With additions and minor editing by J. A. Kiernan, London, Canada).
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)
** Polishing undecalcified bone sections.
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.
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.
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!
** Polylysine-coated slides
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?
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
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.
Bethesda, MD 20889
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.
** Wrinkles in plastic sections
How can I prevent wrinkles in sections (0.5 to 2 micrometers)
of plastic-embedded tissue stained for light microscopy?
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
John A. Kiernan,
Department of Anatomy & Cell Biology,
The University of Western Ontario,
London, CANADA N6A 5C1
** Wrinkles in paraffin sections containing cartilage
Does anyone have a reliable procedure to consistently avoid
wrinkles with cartilage in paraffin sections of trachea (human,
A few tips and wrinkles follow.
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
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”
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. ]
** Thick paraffin sections
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!
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
** Sectioning plastic-embedded specimens
How do you cut flat sections of materials embedded in
poly(glycol methacrylate) (GMA) and other resins, for
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!
MRC Harwell, Oxfordshire, England
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).
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.
** Iodine for removing mercury deposits
[ It is necessary to remove mercury deposits from specimens
fixed in B5 or other fixatives that contain mercuric chloride.
Textbooks recommend either including a solution of iodine (0.5
to 1%) in 70% alcohol in the series of solvents for
dehydrating before embedding, or treating the sections after
hydration with iodine followed by sodium thiosulfate. ]
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?
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.
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.
** Labeling slides
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?
(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
** Sectioning plant material: some hints.
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
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.
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
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.
Berlyn,GP; Miksche,JP (1976): Botanical Microtechnique and
Cytochemistry. Iowa State University Press, Ames, Iowa.
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
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.