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)

** 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)

** 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)

** 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)

** 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. ]

** 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)

** Mold release spray

Question.

Is there something you can spray into an embedding

mold to make it easier to extract the solidified

wax block?

Answer.

I faced the problem of mold-release spray several years ago

by mixing a solution of 5% green dishwashing soap (such as

Palmolive) in 50% Ethanol, then putting it into a pump spray

bottle (available form any housewares department). This

worked AT LEAST as well as the outrageously expensive stuff

sold as “Mold-Release Spray”, and it contained no CFC’s or

other “evils”.

Joanne Lahey

Battelle Duxbury Operations

Duxbury, MA 02332

(laheyj[AT]battelle.org)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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)

** 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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