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PROCESSING, DECALCIFYING, EMBEDDING
** Solvent to replace xylene AND alcohols
Question.
Is there a product that replaces xylene AND alcohols in the
staining procedure? Can you use it before and after the actual
staining is done?
Answer 1.
t-butanol, dioxane and tetrahydrofuran are miscible with
wax, water and resinous mounting media. Of these, only
t-butanol (= tertiary butyl alcohol) is suitable for
ordinary use. (The other two have such hazards as fire,
toxicity and explosive peroxide formation.) t-butanol is
often used in botanical microtechnique; it is quite a bit
more expensive than alcohol or xylene. n-butyl alcohol
mixes with wax and mounting media and is also partly
miscible with water. It's good when you use easily
extracted stains (methyl green-pyronine, for example),
but has unpleasant vapour.
2-butoxyethanol (butyl cellosolve) also has the right
miscibilities, and is quite cheap because it's used
on a big scale industrially.
For microwave processing, isopropyl alcohol is
sometimes recommended. However, this does not mix
with wax. It has to leave the specimen by vaporizing
(boiling) under reduced pressure. This can lead to
considerable tissue damage unless the temperature
and pressure are just right (Bosch et al 1996).
Some staining methods work well, though slowly, without
removing the paraffin beforehand (Kiernan 1996), provided
that there has been no melting or softening of the wax
after mounting the sections on their slides.
References.
Bosch,MMC; Walspaap,CH; Boon,ME (1996): Lessons from the
experimental stage of the two-step vacuum-microwave method
for histoprocessing. Eur. J. Morphol. 34(2), 127-130.
Kiernan,JA (1996): Staining paraffin sections without prior
removal of the wax. Biotechnic & Histochemistry 71(6),
304-310.
John A. Kiernan
London, Canada
(kiernan[AT]uwo.ca)
Answer 2.
We use 99% isopropyl alcohol (IPA) instead ethanol AND xylene
AFTER staining. It is especially useful after staining of lymph
nodes with a modified Maximov-Giemsa method. My laboratory has
used this modification more then 5 years and I have never seen
the same excellent result in comparison with atlases of lymph
nodes biopsy. Moreover, we use IPA with addition of a small
amount of detergent for dehydration of samples. Four changes of
99% IPA+detergent is all you need between water and paraffin.
We never have have problems with any tissues, including large
samples of skin. Our HTs adore IPA.
Dr Yuri Krivolapov
Military Medical Academy
St.-Petersburg, Russia
(krivolapov[AT]bfpg.ru)
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** 2-butoxyethanol ("Clereum") dehydrating or clearing agent
Question.
What are the properties of Clereum? (The MSDS for Clereum
indicates the ingredient information as undiluted
2-butoxyethanol.)
Answer.
It's good to learn that this isn't yet another secret clearing
agent! According to the Merck Index, this compound (also called
butyl cellosolve, or ethylene glycol monobutyl ether) is partly
miscible with water. Its properties as a solvent seem to be
similar to n-butanol; no doubt the higher B.P. (171C) is an
advantage - it won't have n-butanol's nasty cough-making vapour.
Merck says the toxicity is similar to methyl cellosolve
(anaemia, "CNS symptoms" etc; can be absorbed through skin).
The price of 2-butoxyethanol varies with the supplier (May 1997):
Fisher Scientific 4 litres $104 ("Laboratory grade")
Sigma 3 kg $42 (no purity details)
Acros Organics (seems to be part of Fisher)
sell three grades:
2.5 litres $24 (99%)
1 kg $23 (GC)
500 ml $36 (scintillation grade)
If the 99% stuff is OK for histology, perhaps the price isn't
too bad. tert-butanol (99.5%; from Acros) is $67 for 2.5 litres,
and n-butanol (99%) is $27 for 2.5 litres. This makes
2-butoxyethanol quite a good buy for a non-niffy
not-quite-universal solvent. The similarity of its miscibilities
to those of n-butanol suggests that this might be useful for
dehydrating (and clearing) sections that have been stained with
methyl green-pyronine, or other dyes that are easily lost with
ordinary alcoholic dehydration.
John A. Kiernan
Department of Anatomy,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)
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** Decalcification: Acid or EDTA?
Questions.
How should I decalcify a bony specimen or a tooth?
What precautions are needed if galactosidase activity must be
preserved (to identify cells carrying the LacZ gene)?
Answer 1.
Decalcification with EDTA is probably the best method with your
LacZ, due to the enzyme staining you are doing. I would be
careful to adjust the pH of the EDTA solution to the working pH
of enzyme staining in PBS or a TRIS buffer, and rinse carefully
in buffers postdecalcification. Formic acid may ruin LacZ
enzyme staining results.
Gayle Callis
(uvsgc[AT]msu.oscs.montana.edu)
Answer 2.
If the bone is crunchy, you have either not removed all the bone
mineral, or you have transferred the bones from EDTA to alcohol
and have precipitated EDTates in your tissue.
When you decalcify, do you determine the end point using an x-
ray/calcium oxalate/prod with a pointed stick?
How long do you decalcify? Even at 20% EDTA these would take at
least a week with vigorous agitation at room temperature. Is
the EDTA buffered to pH 7? If not, you are using the solution
as an acid decalcifier as well as a chelator. In this case,
assuming your stain still works and will not be affected by acid
pH, change to 10% formic acid, which provides much faster
decalcification. Check the endpoint (when all the calcium is
gone) daily.
[ But see Answer 1 for acid-sensitivity of galactosidase. ]
If you have checked the endpoint and all the calcium is gone,
rinse the tissue in water for at least 8 hours to remove all the
excess EDTA before putting it in alcohol.
Simon Smith
(smiths5[AT]pfizer.com)
Answer 3. (A formic acid procedure for teeth, with oxalate testing)
The protocol we use here at Ind. Univ. School of Dentistry is as
follows:
The protocol we use here at Ind. Univ. School of Dentistry is as
follows:
After teeth are fixed in 10% neutral buffered formalin, they are
placed in wide mouth bottles with a 5% formic Acid solution.
They are then checked each day by pipetting 5 ml of the acid
solution into a test tube to which 1 ml of 2.5% ammonium oxalate
is added. If a white precipitate forms there is still calcium
present. The solution is then changed and the process repeated
the next day. Once I get one negative test the specimen is
grossed as needed and placed back into acid until another
negative is obtained. The specimen is then placed in running
water overnight and processed with the next days run. I know
this can take a long time, but the results are worth it. If you
need anything else let me know.
Lee Ann Baldridge
IUSD Oral Path Group
Indianapolis, IN.
(lhadley[AT]iusd.iupui.edu)
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** Testing for completeness of decalcification
Questions.
How should I test for complete decalcification?
Is the same method OK after either formic acid
or EDTA?
Answer 2.
The ammonium oxalate test is simple. Take a 5 ml sample
of used decalcifying fluid. Neutralize it by adding drops
of strong ammonia (ammonium hydroxide); avoid the fumes!
When the solution turns litmus blue (pH above 7),
add 5 ml of saturated aqueous solution of ammonium
oxalate (about 3%; stable stock solution). Wait for
half an hour. If there is no precipitate, the last
change of decalcifying fluid was free of calcium ions.
According to Eggert & Germain (1979) you can use the ammonium
oxalate test on EDTA. Rosen (1981) said the sensitivity was
higher if you lowered the pH to 3.2-3.6 before doing the test
(instead of neutralizing to pH 7 as done with an acid decalcifier).
Eggert FM, Germain JP 1979. Histochemistry 39: 215-224.
Rosen AD 1981. End-point determination in EDTA decalcification
using ammonium oxalate. Stain Technology 56: 48-49.
John A. Kiernan,
Department of Anatomy & Cell Biology,
The University of Western Ontario,
LONDON, Canada N6A 5C1
(kiernan[AT]uwo.ca)
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** Fatty specimens: Processing into paraffin.
Question.
What is the best way to paraffin-embed specimens that
contain a lot of fat?
Answers.
1. Process by hand, allowing more time and bigger volumes
of all solvents than for non-fatty pieces of tissue.
2. Don't put them through an automatic processor because
you'll get grease in all the solvents. (If you don't
believe this, put a bit of skin in about 10 times its
volume of 95% alcohol for an hour, then add some
water to the alcohol. Result: a milky emulsion.)
3. Xylene is better than a "xylene substitute."
[ Unfortunately I mislaid the sources of these pieces of
advice. For what it's worth, I agree strongly with
the first two, but lack the experience to comment on
the third. J. A. Kiernan. ]
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** Polymethyl methacrylate embedding for bone
Question.
Is it permissible to mix polymerized methyl methacrylate
with the monomer, when making an embedding medium for
undecalcified bone?
Answer.
Using polymethylmethacrylate powder or beads does not affect
the polymerization process, but it does make the preparation
of the partly polymerized embedding mixture easier and safer.
You may care to refer to the following paper.
Difford, J. (1974) "A simplified method for the preparation
of methyl methacrylate embedding medium for undecalcified
bone." Medical Laboratory Technology 31: 79-81.
John Difford
Royal Free Hospital
London, England.
(adford[AT]compuserve.com)
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Question.
Is there something you can spray into an embedding
mold to make it easier to extract the solidified
wax block?
Answer.
I faced the problem of mold-release spray several years ago
by mixing a solution of 5% green dishwashing soap (such as
Palmolive) in 50% Ethanol, then putting it into a pump spray
bottle (available form any housewares department). This
worked AT LEAST as well as the outrageously expensive stuff
sold as "Mold-Release Spray", and it contained no CFC's or
other "evils".
Joanne Lahey
Battelle Duxbury Operations
Duxbury, MA 02332
(laheyj[AT]battelle.org)
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** Paraffin processing of skin
Question.
Could you suggest a processing schedule suitable for skin?
Answer.
This is my processing schedule for skin dehydration and
embedding.
By Hand.
The times suit my working day. I'm sure they could be
altered for any work pattern.
1.) 80% alcohol. = 2 pm.
2.) 80% alcohol. = 5 pm - overnight.
3.) Abs.alc./8% phenol. = 9 am.
4.) Abs. alcohol. = 10 am.
5.) Abs. alcohol. = 12 am.
6.) Abs.alc./amyl acetate.= 3 pm.
7.) Amyl acetate. = 4 pm.
8.) Amyl acetate. = 5 pm. - overnight.
9.) Amyl acetate. = 9 am.
10.) Amyl acetate. = 12 am.
11.) Xylene. = 3 pm.
12.) Wax. = 4 pm.
13.) Wax. = 5pm. - overnight.
14.) Wax. = Embed.
Tissue Processor.
These times I use on a Shandon Histokinette, remember
them ?.
1.) 80% alcohol. = 2 hours.
2.) 80% alcohol. = 2 hours.
3.) Abs. alc./8% phenol. = 1 hour.
4.) Abs. alcohol. = 3 hours.
5.) Abs. alcohol. = 3 hours.
6.) Abs.alc./amyl acetate.= 1 hour.
7.) Amyl acetate. = 3 hours.
8.) amyl acetate. = 3 hours.
9.) Amyl acetate. = 3 hours.
10.) Amyl acetate. = 5 hours.
11.) Xylene. = 1 hour.
12.) Wax. = 9 hours.
13.) Wax. = 9 hours.
14.) Embed.
Ian Montgomery
University of Glasgow, Scotland
(I.Montgomery[AT]bio.gla.ac.uk)
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** Cryoprotection of specimens
Question.
Please recommend a way to protect formaldehyde-fixed
mouse brains to avoid crack and ice crystal holes
that for during rapid freezing. 25% sucrose has been
recommended. Should it be in water or phosphate
buffered saline?
Answer 1.
For ultracryomicrotomy (or should it be cryoultramicrotomy)
Tokuyasu (1989) used 2.3 M (= 78%) sucrose in 0.1M phosphate buffer.
He was working with blocks much smaller than mouse brain, so you
will no doubt have to increase the time. Inflitration of blocks
1 mm wide usually took 30 minutes. He stated that infusion was
complete when the specimen no longer floated on the top of
the sucrose solution. The same author reported that
10-30% PVP and 1.6-2M sucrose provided still better postfreezing
conditions (compared with freezing alone).
We presently use 5% PVA (polyvinyl alcohol) in phosphate buffer
to cryoprotect bone samples before freezing for enzyme and
immunohistochemistry.
One other point that may be worth considering is the method
for freezing. If you are thinking of snap-freezing, I would
recommend hexane instead of isopentane. Hexane freezes at a
considerably higher temperature: about 80 C. Many moons ago,
when I worked in Neuropathology in Scotland, I found that mouse
brains tended to crack when frozen in isopentane, but that we
had much better preservation when freezing in precooled hexane
(we never cryoprotected them though).
Tokuyasu KT. 1989. Use of polyvinylpyrrolidine and polyvinylalchohol
cryoultramicrotomy. Histochem. J. 21:163.
Ronnie Houston
Dallas, Texas
(RHH1[AT]airmail.net)
Answer 2.
It is a common practice to immerse rodent brains in 20-30% sucrose
at 4 C, at least until they sink. If they have been fixed for
only a short time (less than 48 hours), it is probably best to
dissolve the sucrose in PBS rather than water alone.
Rosene et al (1986) found that 20% glycerol with
2% dimethylsulfoxide (DMSO) was better than sucrose.
The sucrose concentration needs to be much higher than
is commonly used - at leased 60% (see Lepault et al,
1997).
References (with brief notes).
Rosene,DL; Roy,NJ; Davis,BJ (1986): A cryoprotection method
that facilitates cutting frozen sections of whole monkey
brains for histological and histochemical processing
without freezing artifact. J. Histochem. Cytochem. 34,
1301-1315.
Techniques compared. Optimum cryoprotection with 4 day
infiltration (4 C) of 20% glycerol & 2% DMSO in buffer
or fixative. Then freeze in isopentane at -75 C (dry
ice). Better than other cryoprotectants (sucrose etc)
and freezing methods.
Lepault,J; Bigot,D; Studer,D; Erk,I (1997): Freezing of aqueous
specimens: an X-ray diffraction study. J. Microsc.
(Oxford) 187(Sep), 158-166.
EM & X-ray diffraction of freezing of sucrose
solutions. Immersion in a liquid cryogen or high
pressure freezing. Sucrose favours formation of
amorphous ice; conc must be 60% or above for
freezing in a cryogenic liquid.
John A. Kiernan
London, Canada
(kiernan[AT]uwo.ca)
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** Cutting sections of toe or finger nails
Question.
Does anyone have a few hints for sectoning toenails?
[ Here is a selection of many replies to this
frequently asked question. ]
Answer 1.
10% Potassium hydroxide. Soak them for at least
4 hrs, but not more than 8.
Noreen S. Gilman (n4xiu[AT]gate.net)
Answer 2.
I have not cut toenails for years. (I do cut my own personal
toenails of course!) However, we used to soak them for a short
time in Nair, which i believe is like Neet, and we got an
excellent section. [See also Answers 4 and 5.]
The procedure is to process the nails, and after they are
embedded treat the paraffin block by putting it in a petri dish
containing the Nair. The Nair is put in first and then the block
is put on top. We treat the block for 5-10 minutes depending on
the size of the nails. We wipe off the block, try cutting it and
put it back for further treatment if needed. It is best to cool
the block on iced water after treatment and before cutting and
to take the first sections.
Marjorie Hagerty
(mhagerty[AT]emc.org)
Answer 3.
I learned a new technique at one of the outstanding workshops at
NSH-Albuquerque. Our hospital switched to this method. After
grossing, place a representative piece (or ALL if melanoma is
indicated) in a cassette and immerse the nail in 5% Tween 80
(Sigma cat#P-4634) for 1-2 hours at least. Overnight won't hurt
it. Then remove and process as usual. I find that if you orient
the nail to cut it perpendicular to the knife it cuts more
easily. Use a charged or polylysine slide (or Elmers glue if
it's really likely that it will float).
Andrea Kelly
Albany Medical College
(andrea_kelly[AT]ccgateway.amc.edu)
Answer 4.
There are several methods in Luna's last book "Histopathologic
Methods and Color Atlas of Special Stains and Tissue Artifacts"
for softening keratin in nails, etc. Fixation in 10% buffered
formalin is necessary to produce crosslinking and thereby
prevent keratin from dissolving completely in softening
solutions. After fixation and BEFORE processing -- place
specimen in "Neet" or other depilatory cream or permanent wave
solution for one to several hours. The key ingredient in these
solutions is thioglycollate. * This is best performed under a
hood because these products smell really bad and will guarantee
an increase in lab traffic by interested personnel wanting to
know "What on earth are you doing?" The specimen should bend
easily before continuing with next step. Wash the specimen in
running tap water for 10 minutes. Dehydrate, clear, and
impregnate with paraffin as desired. Processing times will
depend on which hoof you are processing -- elephants take a lot
longer than goats :-) Get out your nose clip and have fun!
Linda Jenkins
Clemson, SC
(jlinda[AT]ces.clemson.edu)
Answer 5.
We have routinely used "Neet" overnight and had good results.
Recently tried "Neet" at 58 C (it liquefies) for several hours
during the day on a particularly tough nail; it cut beautifully
the next day!
Colin Henderson
St. Joseph's Health Centre
London, Ontario, Canada
(colinh[AT]stj.stjosephs.london.on.ca)
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** Paraffin wax: crystals, additives and cutting
Question.
What are the best polymers or other additives for
reducing crystal size and improving the cutting
propereties of paraffin wax?
Answer.
Paraffin wax is a mixture of (virtually) straight chain
hydrocarbons. Note the word "mixture". Unless you go to
enormous lengths (of purifying or searching for a fine chemical
supplier), you will ALWAYS have a mixture. There is a
relationship between hydrocarbon chain length and melting
point, but as the waxes are always mixtures, melting points are
never exact, either in the compounding or the measuring, but
that is another story!
Perhaps more important than the melting point is the "plastic
point," but that is virtually ignored by our suppliers. The
plastic point occurs about 10 C below the melting point and
its meaning should be fairly obvious - try softening a piece
of physiotherapy wax in your hands and that should explain all
you need to know. The reason the plastic point is important is
related to the sectioning properties of the wax, but we will
come to that later! Crystal size is important in the wax
surounding the tissue and in the tissue spaces, but not in the
tissue per se. Molten wax infiltrates the specimen; the size
and shape of crystals will be influenced by the tissues as the
molten wax solidifies - i.e. crystalises. So we cannot have
"small crystals" infiltrating although smaller crystals will
result from solification in denser tissues.
Some of the theory behind this suggests that wax crystalises
first as flat "plates," the higher melting point hydrocarbons
crystalising first. As successively lower melting points
deposit further plate crystals, they pile up upon one another.
Distortion due to these dynamic events forces the edges or
corners of the most well developed plates to curl and roll.
Eventually, that gives rise to needle shaped crystals, which
some "experts" consider most ideal for microtomy. All this
will be contingent upon the boundaries imposed upon the process
by cell and tissue structures. During microtomy, essentially
two types of forces are exerted in the cutting process. Flow
shearing and point-to-point shearing. Flow shearing is, as you
might expect, the smoother and prcedes ahead of the edge of the
blade. Point to point shearing has forces seeking the line of
least resistence ahead of the blade and these result in a
section of uneven thickness - not that you would notice this
microscopically.
Imagine the difference between cutting through a jelly and
cutting through a beefburger. Now you can imagine where the
importance of the plastic point (as opposed to the melting
point) comes in. Additives to paraffin waxes are intended to
minimise the point-to-point shearing and improve the plastic
flow. The association between the words "plastic" and
"polymers" should now be awakening. Additives to paraffin wax
are usually polymers (of know chain length, for they are
synthesised exactly), with a major role in "harmonizing the
consistency," in part at least by filling in beteen the wax
crystals.
I use pure paraffin wax with no additives, in the belief that
proper processing and a SHARP blade are the central features of
good microtomy. (I just wish I could practise as well as I can
preach!) I have only ever come accross one wax with crystalline
structure significantly different from others, and that is
Ralwax, which can be helpful when cutting decalcified
specimens, etc.
Russ Allison
Cardiff, Wales
(allison[AT]cardiff.ac.uk)
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** Xylene substitutes: what are they?
Question.
What are the various liquids sold as substitutes for
xylene, and are they really safer and just as good?
Answer.
There are two classes of xylene substitutes: limonenes and
aliphatics.
Limonenes are prepared by steam distillation of orange peels.
They are terpenoids rather similar to turpentine. They are
becoming more expensive and difficult to obtain. Their great
disadvantage is the persistent citrus smell, which many people
find intolerable. They are difficult to distil. On the other
hand, they are rather minimally toxic, and are easy to dispose
of. Various brands are interchangeable.
Aliphatics are synthetic hydrocarbons with about the molecular
weight of naphtha. They are odorless, not very toxic, and easily
distilled. They are as difficult to dispose of as xylene.
There are at least six brands of aliphatics, and they are NOT
interchangeable with each other. They vary consierably in flash
point, and they all have different distillation routines.
Richard Allen's Clear-Rite is perhaps the best known of them.
Some of the ones offered by ma-and-pa solvent repackagers are
quite unsatisfactory.
Bob Richmond, Samurai Pathologist
Knoxville TN
(rsrichmond[AT]aol.com)
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** Test for water in used absolute alcohol
Question.
How can I determine whether used "absolute" alcohol is
still OK for the last stage of dehydrating specimens
or slides?
Answer.
Some people add anhydrous copper sulphate to the alcohols used
for processing tissues. It changes colour (white to blue) in the
presence of water, but this does not tell you if there is only a
tiny trace of water or enough to make the alcohol immiscible
with xylene.
You may be interested in a simple method I developed for this
purpose. My job is evaluating histology equipment for the
Medical Devices Agency, (an agency of the Department of Health),
and I was interested in trying to establish "carry-over" in
processing and staining instruments. I started off by adding
known dilutions of alcohol, drop by drop, to different amounts
of xylene, my basic thinking being that water turns xylene
milky, and if one adds enough of the diluted alcohol, the
mixture eventually becomes clear again. From this I developed
the following method:
A measured 5 ml of xylene (the 5 ml is important) is placed in a
50 ml glass beaker and placed on a black background. Using a 1
ml plastic pasteur/transfer/dropping pipette, add the alcohol
for analysis, drop by drop and keep count of the number of
drops, until you can just detect a faint turbidity in the
xylene. Carry on adding the alcohol to the xylene until the
turbidity just clears, again taking note of how many drops were
needed.
Using known dilutions of alcohol, I was able to set up and
standardise the method and obtain reproduceable results
consistently. The method was not sensitive enough to detect the
water in 99% or 98% alcohol.
97% = 5 drops to turn xylene milky, 10 drops to clear the mixture
96% = 4 drops to turn xylene milky, 14 drops to clear the mixture
95% = 3 drops to turn xylene milky, 34 drops to clear the mixture
94% = 3 drops to turn xylene milky, 74 drops to clear the mixture
93% = 3 drops to turn xylene milky, 83 drops to clear the mixture
92% = 3 drops to turn xylene milky, 98 drops to clear the mixture
91% = 3 drops to turn xylene milky, 140 drops to clear the mixture
90% = 3 drops to turn xylene milky, 204 drops to clear the mixture
You would have to initially set up your own range of standard
dilutions with the particular alcohol used in your laboratory
for the sake of accuracy. The 1 ml plastic
pasteur/transfer/dropping pipettes, they can even be called
pastettes, should be held vertically to standardise the size of
the drops, and I tried to use the same brand each time.
This is a simple method, and quick to do, although I should
think the method would give the Biochemists the shudders. It
could help to prolong the life expectancy of the alcohols used
in processors.
Jim Hall
(rmkdh[AT]ucl.ac.uk)
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** Molecular sieves for making anhydrous solvent
Question.
Which type of molecular sieves are used for making anyydrous
acetone or alcohol, and how much should I put in the bottle?
Answer.
The molecular sieve to use for acetone is type 3A, mesh 8-12.
EM Science Catalog # MX1583L/1 for 500 g or /3 for 2kg.
Before using a molecular sieve, you first have to determine
which one to use. Type 3A if for unsaturated hydrocarbons and
polar fluids. These include methanol, ethanol, and acetone.
The 3A refers to the size of the molecule it can absorb.
In this case, less than 3 angstrom. Molecular sieve 3A has
an absorption capacity of 22% by weight.
To dry a liquid, add a slight excess of drying agent.
Next, a little calculation. If the information isn't on the
label, call your vendor and retrieve a C of A (certificate of
analysis) for the lot of solvent you're using. There should be
a spec for water content. This value is the moisture in the
bottle upon release. An opened bottle will have higher moisture,
depending on how hygroscopic the reagent is. Let's use methanol,
which is very hygroscopic, as an example, with the C of A
stating that the water content is 1.0%, which equates to 4 ml
in a 4 liter bottle. 4 ml of water is equal to 4 g of water.
This is 22% of (4 X 100 / 22) = 18.18 g. For excess use
20g of molecular sieves.
Mix thoroughly and allow the liquid to stand. After a few
minutes the drying agent settles to the bottom of the
container. Separation can be completed by decanting or
filtration (suction filtration would work best and fastest ).
How often you would dry a solvent out is dependent on
application, use, and humidity.
TIP: Depending on application and specifications required, the
use of molecular sieves may eliminate to need to purchase
expensive super dry reagents.
Rande Kline & Joe Daniels
Technical Services, EM Science
(rkline[AT]emindustries.com)
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