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1136
Eciency and the results of microtomy are largely dependent on the proper preparation of
the specimen prior to section cuing, and the use of a sharp knife in good condition. This is
discussed separately under ‘cuing of sections’ (Chapter 37 of this Volume).
Medical Laboratory Technology: Volume 3
Knife sharpener
One of the key points forgeing a good section is to use a sharp knife. The abrasive surface
of a knife sharpener helps to keep the knife sharp. The abrasive surface is provided by glass,
copper or iron plates. The amount of abrasive used for knife sharpening is very important.
Use of excessive amount (common tendency) dees the purpose. Follow manufacturer’s
instructions with necessary modications to get the optimum edge. If more than one sharpener is available, it is a good idea to assign certain knives to a specic sharpener. One critical
step common to most sharpeners is that the knife should be placed in the holder the same
way each time, otherwise sharpening may not duplicate the established cuing facets. Plates
must be kept at, with one plate used for the coarse abrasive and one plate used for the ne
abrasive. Make sure that the plates are not mixed up. Whenever plates lose atness or become
too smooth to knife edge, the knife will not sharpen. The knife will not sharpen if more than
one sharpener is available. It is a good idea to assign certain knives to a specic sharpener.
The knife should be cleaned of any abrasives and carefully dried (always wipe and never
across the edge) aer sharpening. In areas of high humidity, the knife should be lightly oiled
for storage, and then cleaned to remove the oil before using.
Flotation Bath
Flotation baths are used to oat and facilitate separation of paran embedded tissue ribbons
while mounting the section on the slide (Figure 36.5). It is basically a temperature controlled
water bath that is kept at 5–10°C below the melting point of the paran used for embedding.
Ideally, the bath should have a dull black interior to optimize the visibility of the oating
translucent paran ribbons. Aer using the bath, pour o the water while warm and wipe
thoroughly to remove adherent bits of paran while they are still so. Note If it is an
electric bath, pour the water over the rim opposite the electrical element. If water gets into
the electrical element, it may be a source of electrical hazard. One should be careful in using
the otation bath. If the bath is too hot, the ribbons get overstretched. If the ribbon is allowed
to oat for too long period, artefacts may develop and mimic many pathological conditions
like oedema. Ribbons must be stretched gently as they are placed on the otation bath, and
then small wrinkles and folds should be quickly but cautiously teased out of the sections.
Air bubbles may get trapped under the section which can be drawn to one side by blunt
Figure 36.5 Method of guiding section: (a) on to a slide and (b) on a tissue oatation bath
Introduction to Histotechnology and Cytotechnology
dissecting needle that has been bent at an angle to reach under the ribbon. Care should be
exercised so that the ribbon is not pierced or torn. Sections may be removed from the knife
and transferred to the otation bath with ngers, forceps, a brush, or a wooden application
stick. The applicator stick aached to one end of the ribbon will aid in stretching the ribbon
and will oat on the bath intact without geing detached from the ribbon. The preparation of
slides to pick up the paran section is discussed in the tissue processing section.
1137
Dryer Oven and Slide Warmer
In the preparation for staining, paran
has to be removed. In order to remove
paran (deparanization) the slides need
to be warmed which can either be done in
the paran oven or on the slide warmer
(Figure 36.6). The laer is handy and can
be placed adjacent to the staining area.
The slides must be completely dry before
they are deparanized, because water
le on the slides will not mix with xylene
(clearing agent) and will cause incomplete
removal of paran. This is indicated by
white spots that can be seen in tissue as
the slides are removed from xylene. This
artefact is dicult to correct as they stain
more intensely later. These slides should be treated with absolute alcohol to remove any
residual water and then placed back in xylene to remove any remaining paran. The slides
should then be taken through absolute and 95% alcohol to water and stained as desired.
Incomplete drying can also cause sections to wash o during staining. Overheating of the
sections during drying also can create artefacts.
The temperature of the dryers and hot plates are usually kept at 60°C which is just above
the melting point of paran. Forced-air slide dryers take 7–10 min while hot oven requires
1 h, ensuring complete drying.
Figure 36.6 Slide warmer
Tissue Processor and Stainer
Manual staining is done in smaller laboratories. Use of staining dishes, like the coplin jars
(that holds 5–10 slides) is common and a timer is used for determining the duration in
dierent reagents. The results are, however, not always reliable. Hence, many laboratories
in urban areas of developing countries are now switching to automated tissue processors
Figure 36.7 (a) Automated tissue processor and (b) Automated slide stainer
1138
Medical Laboratory Technology: Volume 3
(Figure 36.7a). This equipment automatically xes, dehydrates, clears, and inltrates tissues
and makes them ready for embedding. The two major types of tissue processor include the
open type and the closed type. In the open type, tissues are transported from one solution to
the next, and in the closed system the tissues stay stationary while the solutions are changed.
The processor must be kept clean and a routine reagent rotation and/or change cycle
determined by usage must be established and rigidly adhered to. The temperature of the
paran must be carefully adjusted to no more that 2–4°C above the melting point of the
paran in use, or the tissue will be brile and over hardened. The temperature must be
monitored and recorded daily, with adjustments made in the paran bath temperature when
indicated by improper temperature readings. The machine consists of the following:
• time clock
• circular superstructure that contains the basket carrier
• receptacle basket
• receptacles (stainless steel or plastic capsules)
• circular deck which holds the reagent beakers and paran baths
Small blocks of tissues are enclosed in the perforated capsules. These capsules are placed
in the basket, which in turn is aached to one of 12 yokes in the superstructure while it is
in the raised position. The entire superstructure descends, immersing the basket in the rst
solution and sealing the other reagent beakers to prevent evaporation. To move the basket
from one reagent to the next in the processing sequence, the entire superstructure ascends and
descends at scheduled intervals controlled by the time clock. During immersion in the uids,
the basket oscillates up and down in a reciprocal motion to keep, the tissue and reagents in a
state of controlled agitation, which signicantly increases the speed of penetration. Aer the
prescribed amount of time in the last paran bath, the tissue remains there until removed
manually.
It takes about 16 h for routine processing. Hence the machine is set in the aernoon and the
processed material is taken out on the following morning for embedding.
The solutions must be changed at regular intervals (twice a week). If any of the solutions
are cloudy, changing should be done more frequently. If there are several beakers of the
same solution, discard only the rst beaker and move the others up in place by decanting the
reagent from one to the other beaker and then add the fresh one at the end. If the solution has
evaporated, replace the lost uid. Clean the beakers with detergent (do not use acid). If the
paran is sticking to the rim of the beaker containing paran, remove it carefully or it may
come in way of the moving parts of the automatic processor.
Automatic stainers (Figure 36.7b) perform routine staining that saves time and yields
consistent reliable results. Slides are placed in staining troughs with separate baskets that
enable up to 20 slides to be stained at the same time. There are three types of automatic
stainers—linear, revolving, and robotic. In linear stainers, slides are transferred from one
container (vat) to the next with the same time allowed in
each container. The time in each dierent solution can be
changed only by varying the number of containers holding
that particular reagent. Revolving stainers operate on the
same principle as the open tissue processor, and the time
allowed in each solution may be varied. Robotic stainers are
computerized and the timing in each solution is programmed.
In the following chapter (Chapter 37 of this volume), automatic tissue processors will be discussed again along with
their procedure.
Incubators
Some special stains require incubation which is done either
in a water bath or in an incubator (Figure 36.8). The incubator
maintains a temperature of 37°C.
Figure 36.8 Incubator
Introduction to Histotechnology and Cytotechnology
1139
Freezers and Refrigerators
Freezers and refrigerators are used for storing many of the reagents. Freezers maintain a
temperature of –20°C while the refrigerators are kept at 4°C.
laBoratory supplIEs
Microscope Slides and Coverslips
Microscope slides and coverslips are most frequently used in the histology laboratory and
hence a short discussion of their cleaning and storage is in order. Procedures for the labelling
of slides and storage of specimens will also be mentioned in the following discussion.
In recent years, special microscope slides have been introduced into the market, which
bear a permanent positive charge. They electrostatically aract tissue sections and cytology
preparations, binding them to the slide. These slides form a bridge so that covalent bonds
develop between sections and the glass. Tissue sections and cytological preparations adhere
beer to these glass slides without the need for special adhesives or protein coatings.
Cleaning of slides and coverslips
Reagent
• Acid alcohol (1% HCl in 70% alcohol)
• 70% ethyl alcohol 1000 mL
• HCl (concentrated) 10 mL
Procedure
• Clean the used slides and coverslips in acid alcohol.
• Rinse them with water and then place in 95% alcohol.
• Finally, take out from the alcohol, polish, and dry each one with lintless cloth (e.g.,
surgical gauze).
• Cleaned slides and coverslips should be stored in clean covered boxes or in dishes with
lids.
• For routine work, dip new slides and coverslips in 95% alcohol and polish. Slides and
coverslips to be used in uorescent microscopy must be thoroughly cleaned with acid
alcohol.
Labelling of slides
If the slides have frosted ends (Figure 36.9), glassmarking ink (or India ink) can be used. If the
slides are not frosted, the slides are etched with
a diamond marking pencil. Some laboratories
use etching for temporary marking of the slides
and then at the nal stage (aer mounting) use a
special label that is aached to the slide. Do not use
a wax pencil for marking the slides.
Another safe way is to write (or type) the identication number on a piece of paper which is placed
on the slide next to the section. The mounting
reagent is then applied, and the coverslip placed in
position. The number remains permanently sealed
with the section.
Figure 36.9 Microscope slides
1140
Medical Laboratory Technology: Volume 3
Containers for slides
Cardboard trays are the most convenient. Wooden or plastic boxes or metal ling cabinets are
more expensive.
Containers for Specimens
The use of plastic bags to store specimens with preserving solutions is becoming more
popular. Only thick plastic bags should be used. The specimen can be kept initially in a
wide-mouthed bole; aer xation, the xative is poured out and the specimen is kept in a
heat-sealed plastic bag.
Paran blocks with embedded specimens should be stored with embedded identication
tags. These are then stored in wooden or plastic boxes. Cardboard boxes can also be used, but
these must be protected from rodents and insects. It is important to store the paran blocks
in a cool place.
rEagEnts
A great number of histologic procedures in use call for a variety of chemicals and reagents.
These include xatives, decalcifying solutions, embedding materials, stains, solvents, clearing reagents, mounting media and a large number of miscellaneous solutions. In preparing
various reagents, one should rst read the label on the bole for care and storage instructions
and precisely follow the directions given in the method of their preparation. Some of the basic
rules of preparing solutions are given later in this section. The boiling point of solvents is
important if certain limits are stated. Many of them are highly ammable and should not be
inhaled. All instructions from the material safety data sheets (MSDS) of all chemicals should
be read and precautions taken accordingly. All reagents and stains should be tested for proper
reactivity before being put into regular use. Given below are some of the reagents used:
• Embedding material: Paran wax (melting point 50–55°C)
• Fixative: Formaldehyde
• Ethyl alcohol, xylene (or xylol)
• Mounting medium: DPX, Permount
• Egg albumin
• Glacial acetic acid, mercuric oxide, ammonium aluminium sulphate
• Decalciers: Formic acid, nitric acid, hydrochloric acid
• Stains: Haematoxylin, erythrosin B, eosin, fast green FCF, alizarin red S, rose Bengal,
aniline blue, fuchsin, auramine-O, safranin-O, azure dyes, Giemsa stain, light green,
Sudan III, malachite green, indigo carmine, brilliant green, Sudan black B, methyl green,
thionin, carmine, Wright stain, methyl orange, carbol fuchsin, Congo red, gentian violet,
neutral red, iodine solution, Gram crystal violet, methylene blue, orange-G, Papanicolaou stain, phloxine, safranin solution and periodic acid Schi reagent.
Mounting Media
Stained sections need to be mounted on an appropriate mounting media and then covered
with coverslip before the sections can be examined under a microscope. There are two basic
mounting media—aqueous and resinous.
Aqueous mounting media
The aqueous mounting media are used when dehydrating and clearing adversely aect the
stain. Gelatine-glycerine mixture is non-hazardous, water-based, mounting media which is
Introduction to Histotechnology and Cytotechnology
ideal for mounting immunohistochemically stained tissues. These tissues can be damaged
by organic solvents, such as xylene or toluene. Hence, tissue sections stained with organic
chromogens are best preserved using aqueous mounting media.
1141
Resinous mounting media
Some mounting media oxidize rapidly upon exposure to air making the slides unsuitable to
examine aer long storage. This is why use of Canada-balsam has now become obsolete. It
is replaced by resinous mounting media. These include DPX, Permount and Biomount. The
resinous media consists of solid resin dissolved in an appropriate solvent; the viscosity of
the medium should be such that the solution will enter the tissue spaces and ow readily
between slide and cover glass. Air bubbles should be displaced quickly. Most resinous media
are dissolved in toluene or xylene. Many laboratories prefer xylene as the solvent because the
slides are usually mounted aer xylene (clearing) treatment. All of the resinous mounting
media will cause a gradual fading. The two most commonly used resin-based mounting
media are DPX, Permount and Biomount. They will now be discussed here.
DPX mountant is a mixture of distyrene, a plasticizer and xylene. It is a colourless synthetic
resin mounting media that replaced Xylene-balsam in modern laboratories. It preserves
the stain and dries quickly. Permount (Fisher-Scientic) is a toluene-based synthetic resin
mounting medium. Many laboratories consider its use as the right choice for both rapid
mounting and long-term storage of slides. Its low viscosity allows for a thinner mounting
layer oering beer optical quality and bubble-free preparations. It has a refractive index near
that of a xed protein which helps to keep images free of distortion. It is ideal for mounting
coverslips to slides with thick or thin specimens Permount preserves most biological stains
with lile or no fading when the slides are stored in darkness. It contains an anti-oxidant
to prevent the formation of annular rings. Biomount is a specially formulated mounting
medium that reduces fading of some stains. It is suitable for resins as well as wax embedded
sections of tissue.
Preparation of Reagent Solutions
While working in the histology and cytology laboratories the technician will have to prepare
a number of reagent solutions. It is essential to be familiar with the expressions of the strength
of solutions to be made (Chapter 6 of Vol. 1). Most laboratories keep stock solutions which are
diluted to working solutions when needed. Stock solutions have a beer shelf life and take
up less storage space. The technician must be very careful in preparing reagent solutions.
An error made at this stage may cause considerable aggravation. A solution has two components, the solvent and the solute. The strength of the solution is most oen expressed in the
weight/volume ratio or volume/volume ratio. Water is the most common solvent into which
pre-weighed solutes are dissolved and made up to the desired volume. The use of tap water
is discouraged because it contains a considerable number of impurities. Distilled water is the
ideal solvent, but deionized water is both cheaper and easier to obtain and is adequate for
most purposes. It is important to check the eciency of the deionizer column frequently to
make certain that the electrolytes are totally removed (Chapter 4 of Vol. 1).
The presence of chloride as an impurity in the solvent can create problems with the
solutions made for histological studies. A simple way to test the presence of chloride in water
is by adding a drop of silver nitrate solution (2%, w/v, aqueous) in an aliquot of water. If
the water becomes cloudy, chloride is present. Re-purify the water before use. Occasionally,
boiled, cooled and decanted water may be free of chloride and can be used in places where
distillation or deionization facilities are not available.
Dilutions from stock solutions
If a series of working solutions are needed with varying strengths, the technician should be
able to gure out the mathematical relationship (Chapter 6 in Vol. 1). As varying strengths of
1142
Medical Laboratory Technology: Volume 3
alcohols are needed in processing specimens for histological examination, we will use two
examples in order to show the calculation steps.
Note Only 95% alcohol is commercially available unless one treats 95% alcohol with calcium
chloride or some such dehydrant to remove the water (5%). This is not necessary.
Example 1: Prepare 1000 mL of a 70% alcohol (#1, working solution) from 95% alcohol
(#2, stock solution).
Solution:
Formula: S1V1 = S2V
2
where S1 and S2 are strengths of solutions #1 and #2, respectively.
Note All the strengths are expressed in the same unit (%).
S1 = 95 (initial) and S2 = 70 (nal)
V1 and V2 are the volumes of the solutions 1 and 2, respectively.
Consider V1 = xmL when the nal volume V2 = 1000 mL
Then the above formula will be wrien as
90 × x = 75 × l000
or x =
75 × 1000
90
= 736.8 mL
Thus, if 736.8 mL of 90% alcohol is made to (q.s.) 1000 mL with distilled water, the strength of
the resulting solution will be 70% alcohol.
Example 2: Prepare 500 mL of 5% ferric chloride solution (working solution) from 29% ferric
chloride solution (stock solution).
Solution:
Formula: S1V1 = S2V
2
29 × x = 5 × 500
x = 86.2
Take 86.2 mL of 29% ferric chloride and make it to 500 mL. The nal solution will be of 5%
strength.
Example 3: Prepare a 1000 mL solution of 80% alcohol from 95% and 50% alcohol solutions.
Solution:
The following formula is the elaboration of the same principle of strength/volume relationship.
Here, however, we are involved with two initial solutions which when mixed, gives a third
solution of dierent strength.
(S1 × V1) + (S2 × V2) = S3 ×V
3
The strengths (all in the same % units) of initial solutions are, respectively, S1 (95) and S2
(50) while the nal strength S3 is 80.
Now consider the volume of V1 (95%) = x, then V2 will be 1000 – x because the nal volume
is 1000 mL. If you put the values in the above formula, it looks like this
(95 × x)+ [50 × (1000 – x)] = 80 × 1000
or 95x + 50000 – 50x = 80000
or 45x = 30000
x = 666.7 mL
Thus, if you take 666.7 of the 95% alcohol and make it to 1000 mL with the 50% alcohol, you
will have 1000 mL of 80% alcohol.
Introduction to Histotechnology and Cytotechnology
1143
routInE and spEcIal staInIng: a rEvIEw
Introduction
In a histopathology laboratory, the term “routine staining” refers to the hematoxylin and
eosin stain (H&E) that is used routinely with all tissue specimens to reveal the underlying
tissue structures and conditions. The term “special stains” has long been used to refer to a
large number of alternative staining techniques that are used when the H&E does not provide
all the information a pathologist or researcher needs.
The use of routine H&E stain and special stains play a critical role in tissue-based diagnosis
through histology. Colouring of the transparent tissue sections, allow the pathologists to
look for the presence or prevalence of cell types, structures or even microorganisms such as
bacteria inside the tissues.
Histology Sample Preparation
Tissue specimens are rst prepared for histological studies. This is accomplished through
four steps listed below. The goal is to allow the so tissues supported in a medium that allows
sectioning:
• Fixation that preserves the tissue,
• Processing that dehydrates, clears and inltrates the tissue with paran wax,
• Embedding that allows orientation of the specimen in a “block” that can be sectioned
and is easy to store and handle, and
• Sectioning using a microtome to produce very thin sections that are placed on a
microscope slide ready for staining.
The routine procedure has limitations. Imagine an emergency during surgery. The surgeon
needs to locate a tumor margin to ensure it has all been removed. The surgeon cannot wait to
process the tissues through routine staining. Hence, an immediate report is called for. Under
this condition, a frozen sectioning is the right alternative. Here the tissue is quickly frozen
in liquid nitrogen, which preserves the tissue and provides sucient hardness so it can be
sectioned immediately using a cryostat.
Fixation
Most oen formalin is used as the xative. Buered formalin, alcoholic formalin and Bouin’s
formalin may be used. Avoid xatives with dialdehyde (glutaraldehyde), as the free aldehyde
groups may bind the Schi s reagent and produce non-specic staining.
Paraffin sections
Following xation, process the section through paran embedding and then cut the sections
4–6 micronswide.
Staining: Method 1
1. Deparanise with xylene or a xylene substitute.
2. Rehydrate through graded alcohols to deionized water; tissues must be neutral.
3. Dip slides in Periodic Acid solution for 5 min.
4. Rinse in deionized water (multiple changes).
5. Dip slides in Schi’s reagent (for 15 min).
6. Give a nal wash with lukewarm tap water (for 10 min) before counterstaining.
7. Counterstain for 3-4 min with Gill II Haematoxylin.
8. Rinse in running tap water for 5 min.
9. Dehydrate through two changes of 95% alcohol and absolute alcohol two minutes each.
1144
10. Pass the slides through two changes of xylene and then mount on a mounting media
that is miscible with xylene.
Medical Laboratory Technology: Volume 3
Method 2
1. Deparanize with xylene or a xylene substitute.
2. Rehydrate through graded alcohols to deionized water.
3. Place sections in a plastic Coplin jar containing Periodic Acid (40–50 mL) and micro-
wave for 10 s.
4. Gently mix the solution by swirling and allow it to stand for 1 min.
5. Rinse in several changes of deionized water.
6. Add the Schi’sreagent (40–50 mL) into the plastic Coplin jar and microwave for 15 s.
7. Gently mix the Schi’s reagent by swirling and allow it to stand for 1 min.
8. Wash in lukewarm tap water for 5 min.
9. Counterstain with Gill II Hematoxylin for 3–4 min.
10. Rinse in running tap water for 5 min.
11. Dehydrate through two changes of 95% alcohol and 100% alcohol, two minutes each.
12. Clear in two changes of xylene (two minutes each) and mount with a mounting media
that is miscible with xylene.
Microwave and laboratory fumes
Histology laboratory contains volatile chemicals and microwave ovens and both are health
hazards. Microwave ovens must be properly ventilated to prevent accumulation of fumes.
Because of the dierences in microwave manufacturers, seings for power and frequencies
among various models may widely vary. So, it may be necessary to adjust the power levels to
obtain optimum results. In general, microwaves of histology laboratory should be at a power
seing of 800 was unless otherwise noted.
Keep the chemicals always covered. Coplin jars are always provided with cap. The caps
should be loosely applied to prevent spills. Caps with ventilation holes are also convenient
to use.
Ventilation of the histology laboratory should be properly designed. The laboratory should
not be a health hazard for its workers.
Technical notes
Distilled water may be used in place of deionized water.
Expected results
Many tissues and cell types will demonstrate a bright magenta coloration indicative of a
positive PAS reaction. This includes, but is not limited to, glycogen containing hepatocytes,
mucin in the gastrointestinal tract, as well as basement membranes in various sites. Nuclei
should stain purple/blue with haematoxylin.
Recommended controls
Liver or gastrointestinal epithelium (small intestine, appendix, colon)
Preparing Tissue for Staining
Before tissue can be stained and viewed, it must be prepared so that a very thin section, only
one cell thick, can be cut and placed on to a microscope slide. This involves xing the tissue
(so it does not decay) then hardening and supporting it so that it can be cut into very thin
sections needed (typically 2–7 mm). There are two main techniques used for this, referred to
as frozen sections and paran-embedded sections.
Introduction to Histotechnology and Cytotechnology
Frozen sections are used when answers are needed fast, typically during surgery where
the surgeon needs to know the excision margin when removing a tumour. They are quick to
produce, but typically do not create the same section quality as the paran technique. The
process for frozen section preparation is as follows:
• Tissue is quickly frozen to preserve and harden.
• The frozen tissue is sectioned in cryostat (a sectioning microtome in a freezing chamber)
and placed on a microscope slide for staining.
• The section is xed immediately before it begins to decay and is then stained.
When paran sections are to be prepared, the specimen is rst preserved with a xative
and then the tissue structure is supported by inltrating the specimen with paran wax. The
process is more time-consuming than creating frozen sections, but provides beer quality
staining in most cases and the resultant samples (referred to as blocks) can be stored almost
indenitely. The paran section process is as follows:
1. Fixation preserves the tissue (typically using a formaldehyde-based solution).
2. Grossing isolates the particular area of tissue to be sectioned.
3. Tissue processing uses a sequence of reagents to replace an aqueous (water-based)
environment with a hydrophobic one enabling tissue elements to be inltrated with
paran wax.
4. Embedding allows specimen orientation and secures the specimen in a block of wax for
section cuing and storage.
5. Sectioning is done on a microtome that cuts very ne sections which are oated-out on
a water bath then picked up and placed on microscope slides.
6. The slides are then dried in an oven or on a hot plate to remove moisture and help the
tissue adhere to the slide.
7. The tissue on the slide is now ready for staining.
8. The rst staining step is dewaxing which uses a solvent to remove the wax from the
slide prior to staining. This is always done as a part of the staining process. When a
stain is complete the section is covered with a cover glass that makes the preparation
permanent.
1145
Automating Staining Process
Automated strainers are not so common in the laboratories of developing countries. Routine
H&E stain as well as special stains have been used successfully. The best part of the automated
system is to reduce human error but at the same time increased cost and frequent breakdown
(load-shedding) keep the system o limits.
rEvIEw quEstIons
1. What are the functional dierences between histotechnology and cytotechnology?
2. What is autolysis? How can this be prevented?
3. What is the signicance of the following in preparation of histological specimens:
xation, embedding, dehydration and rehydration?
4. What is the use of a microtome? What are the dierent types of microtomes? How
would you maintain a microtome in order to give many years of ecient service?
5. Why is the vacuum embedding oven superior to the ordinary embedding oven?
6. What is the temperature maintained by the paran oven and tissue oating bath?
7. How are the microscope slides and coverslips cleaned and stored?
8. How would you prepare a 500 mL of 70% alcohol from 95% alcohol?
9. How much ferric chloride would you weigh in preparing 1000 mL of 29% solution?