LAB TOPIC PARAFFIN HISTOLOGY

Mary Lee

AIM:
By the end of the topic participants should be able to discuss the path of a piece of tissue
from fixation through processing, embedding, cutting, staining and proper presentation,
having
1. had a brief introduction to a microtome,
2. collected cut sections from the waterbath
3. stained sections using 2 methods, and
4. compared the results of different staining methods.
The practical work to be completed in our laboratory with supervision.

PROCEDURE:
The laboratory work is to be undertaken in 2 sessions.
Session 1
1. Examine processing procedures.
2. Examine embedding techniques
3. Discuss the care and use of knives and microtomes.
4. Have a brief introduction to a microtome and its use.
5. Collect cut sections from the waterbath.
6. Look at serial sections stained with 8 different stains and prepare a comparison
table, followed by discussion.
Session 2
1. Discuss the chemistry of staining
2. Discuss the practicalities of staining
3. Stain 2 sections with H&E.
4. Stain 1 section with Van Gieson
5. Discuss slide presentation and view examples.

ASSESSMENT
Contributes to the laboratory tasks component of your assessment.
1. Question sheet session 1 10 marks
2. Question sheet session 2 10 marks
These are due 1 week after the practical session.

SAFETY
1. Many of the chemicals encountered in paraffin wax tissue processing and staining are
toxic and known carcinogens (ie toluene). Others may be dangerous to store, prepare or
use. Use fume hoods or extractor fans.
2. Knives are extremely sharp, please treat them with respect. Do not carry a knife around
unless it is in its box. Do not catch a dropped knife.
3. No eating, drinking, applying makeup, or sucking on pencils etc while in the laboratory.
4. Wash your hands when leaving the laboratory; especially before eating, going to the toilet
or applying makeup.
5. Wear appropriate protective clothing and closed in footwear.
6. If in doubt, ask!
 PART 1
PARAFFIN PROCESSING

Fresh tissue follows a number of stages before being cast in a mould to make a block
suitable for cutting. Processing stages for paraffin, LVN and TEM are very similar. The
purpose of each stage is the same, the solutions, times and sample size will differ.
For paraffin processed tissue the most usual fixative is formalin. The tissue is dehydrated in
graded ethanol. Toluene and xylene are the most usual clearing agents and the tissue is
infiltrated with paraffin wax with a melting point about 57oC. The tissue is cast in wax
blocks and when cool is ready for cutting.
1. FIXATION
Good preservation of tissues is an important factor in the production of satisfactory
tissue sections.

Aims of fixation:
1. To prevent autolysis or decomposition due to bacterial, autolytic and osmotic
change.
2. To preserve tissue against subsequent changes during processing and embedding.
3. To preserve tissue as near to its original form as possible.
4. To give the tissue a texture which permits easy sectioning.
5. To render the various constituents of the tissue reactive to the proposed stains or
treatments.

Factors affecting fixation are

fixative used
time
temperature
volume of fixative to volume of sample
pH
subsequent studies of the tissue
osmolality
Formalin is a commonly used fixative. Other fixatives may contain one or more of the
following - mercuric chloride, picric acid, acetic acid, ethanol, chloroform, and potassium
dichromate. These may be used singly, together, or sequentially. Fixation may also be
carried out using microwaves.

ALL of the above substances are toxic, and some are carcinogenic.
Formalin containing fixatives may be prepared using paraformaldehyde powder. Researchers
tend to use paraformaldehyde powder. In a clinical setting where large volumes are required
it is more usual to employ commercial formaldehyde. Formaldehyde is a gas that is
dissolved in water to about 40% concentration and stabilised with methanol. Thus:
100% formalin contains 40% formaldehyde
and
10% formalin contains 4% formaldehyde
On standing formalin tends to become acidic and produce a black pigment in the tissue
called formalin pigment. To overcome this problem formalin is usually prepared in a
phosphate buffer at pH 7.4. The buffering capacity is inefficient by bloody specimens, and
such samples should have the fixative changed to remove the excess blood.

10% Buffered Formalin

Formaldehyde 40% 10ml
Tap water 90ml
Sodium di hydrogen orthophosphate Na H2PO4 0.35g
Disodium hydrogen orthophosphate Na2HPO4 0.65g
(both salts are anhydrous)

Dissolve the salts in some of the water. In a fume hood, add the formaldehyde and then the
remainder of the water. Mix well.
NOTE: Formalin is toxic and carcinogenic. Avoid inhaling the vapours, exposing the skin,
or splashing the eyes. Wear safety glasses while handling concentrated formalin, especially if
you wear contact lenses.
Tissue fixed in buffered formalin is
*Usually fixed at room temperature,
*For 2 - 24 hrs,
*Using 10 times the volume of fixative to the volume of specimen
*And is suitable for routine staining and immuno histochemistry under light microscopy
2. PREPARATION OF BLOCKS
In order for thin sections <20 µm to be cut, the tissue must be hard and supported with
minimal damage to the tissue. There are 4 fairly common techniques
1. paraffin embedding
2. frozen sections
3. resin embedding
4. low viscosity nitrocellulose embedding (LVN)
Paraffin is most commonly used. Once fixation has occurred, a representative area of
approximately 5mm thickness is processed, along with an accompanying label, to infiltrate
the tissue with wax. This is usually performed on an automated tissue processor.
3. DEHYDRATION
The traditional method of dehydration –removing the water from the tissue - is the use of
ascending concentrations of alcohols. This avoids the sudden change from water to absolute
ethanol. that can lead to excessive shrinkage and hardening with subsequent problems in
section cutting.
Dehydration usually commences using 70% ethanol, progressing to absolute alcohol. eg.
70%, 80%, 90%, 100%.
ALCOHOL IS TOXIC and can be absorbed by inhalation as well as ingestion
4. CLEARING
Clearing agents are important intermediaries, since wax is not miscible with water or
dehydrating agents. Clearing solutions must be miscible with both dehydrating fluids
(alcohols) and paraffin wax. They frequently have a high index of refraction and impart a
translucency to tissues, hence the term clearing. The most commonly used agents, xylene
and toluene, tend to harden tissue if left in contact for extensive periods.
NOTE: TOLUENE AND XYLENE ARE KNOWN CARCINOGENS
They can be absorbed by inhalation, ingestion and skin exposure.
5. WAX INFILTRATION
A suitable paraffin wax MP 54-58°C is generally used. Tissue should be kept in heated wax
for the minimum time consistent with complete infiltration (2 to 4 hours). It is important to
retain the wax in a liquid condition but below 60°C while tissue is infiltrated. Higher
temperatures both harden and distort nucleic acid structure and may destroy the plasticisers
added to the wax. Two wax baths are used:
The first removes the clearing agent that diffuses from the tissues to the wax.
The second completes the infiltration.
Vacuum embedding is useful to remove residual air from tissues.
NB: WAX ADDITIVES MAY BE TOXIC. Avoid breathing in the vapours.
60°C is sufficient temperature to cause burns. Take care!
6. EMBEDDING
Following infiltration, tissues are cast in fresh wax in a mould. The blocks should be labelled
so that they can be identified. To make a complete bond between infiltrating and embedding
wax, both must be liquid. The solid block is hard and allows thin sectioning of the tissue.
Rapid cooling of the mould on a cold block is advocated. If the moulds are left to cool for
too long they are inclined to produce planes of crystallisation within the block that lead to
cracking. This can make ribboning difficult when cutting sections.
7. OPERATION OF AN AUTOMATED TISSUE PROCESSOR
The automated rotary tissue processor consists of 12 available solution stations. The fixed
tissue is placed in baskets, which are then rotated at suitable time intervals through the
dehydration and clearing agents and finally the waxes. Some form of agitation is usually
employed. There may be manual clock-face on the instrument allowing different processing
schedules to be selected, or the machine may be electronically controlled, with a keypad.
Automated sealed chamber processors move the available solutions into and out of the
processing chamber, and allow for the selection of heat, vacuum and pressure. The total
number of solutions available is usually between 12 and 14. Pressure and vacuum replace
the need for agitation used in rotary processors or when manually processing. While these
instruments are much more costly and more difficult to repair, they provide a large range of
options in usage and drastically reduce the vapour exposure suffered by persons in the
vicinity. For further information on suitable processing schedules for tissues please refer to
the following methods, or a textbook.

! Make records of tissue fixation and processing runs.

 PROCESSING SCHEDULES

SCHEDULE 1 STANDARD 16½ HOUR CYCLE

(14 HOURS IF START FROM 70% ALCOHOL)
Tissue blocks between 5mm and 1cm thick
Samples fit into standard or mega cassettes
Automatic Processor Programme A.

STATION SOLUTION TIME CLOCKTIME

1 Formalin set delay to 4:30pm

2 Normal Saline 21 mins 4:30pm
3 50% Ethanol with 1% detergent 2 hrs 5:00pm
4 70% Ethanol with 1% detergent 2 hrs 7:00pm
5 90% Ethanol with 1% detergent 2 hrs 9:00pm
6 Absolute Ethanol with 1% detergent 2 hrs 11:00pm
7 Absolute Ethanol with 1% detergent 2 hrs 1:00am
8 Absolute Ethanol with 1% detergent 2 hrs 3:00am
9 Toluene 1 hr 5:00am
10 Toluene 1 hr 6:00am
11 Wax 1 hrs 7:00am
12 Wax 1 hrs 8:00am
13 Wax vacuum embedder 25 mmHg ½hr 9:00am
14 block out 9.30am

* To start processing in 70% alcohol, set delay for 7pm

** Use low ionic strength detergent (eg Truce) to assist with dehydrating, particularly in fatty samples

SCHEDULE 2 STANDARD 10 HOUR CYCLE

(blocks 1cm2 x 5mm thick)

STATION SOLUTION TIME CLOCK TIME

1 Tissue in formalin SET DELAY TO

10:00pm
2 Normal Saline: 21 mins 10pm
3 50% Ethanol 1 HR 10:30pm
4 70% Ethanol 1 HR 11:30pm
5 90% Ethanol 1 HR 12:30am
6 Absolute Ethanol 1 HR 1:30am
7 Absolute Ethanol 1 HR 2:30am
8 Absolute Ethanol 1 HR 3:30am
9 Toluene 1 HR 4:30am
10 Toluene 1 HR 5:30am
11 Wax 1 HR 6:30am
12 Wax 1 hr 7:30am
13 Wax, vacuum embedder 25 ½HR 8:30am
mmHg block out 9:00am

* 1% low ionic strength detergent is added to all the solutions containing alcohol, stations 3 to 7
** To start processing from 70% alcohol, set delay for 12:30
 CUTTING SECTIONS
1. MICROTOME KNIVES

Microtome knives are sharp! Disposable blades are the most common knife used today for
paraffin sectioning. They fit a holder shaped similarly to a solid knife. The wedge, profile C,
knife is less commonly used these days and requires to be sharpened. In this lab wedge
knives are sharpened on the Shandon Elliott Autosharp. The cutting edge of the knife is
angled onto a metallic plate covered with abrasive paste containing diamonds. The knife is
sharpened using a circular motion. There are other techniques available but the increased
use of disposable blades means knife sharpeners are becoming rare.

2. CUTTING SECTIONS

Paraffin wax sections are cut on a rotary microtome.

Factors involved in producing ribboned sections are:
Block temperature
Air temperature
Humidity
Nature/size of tissue
Angle/sharpness of knife
Thickness of sections being cut

Sections are usually cut at 4-5 µm but can vary from 2 to 30 µm.

3. FLOATING OUT SECTIONS

Once ribbons are cut, they are floated out on a water bath heated to about 45°C and picked
up onto slides. To assist the tissue section to adhere to the glass slide the slides are
sometimes coated with wood glue (Araldite), gelatin dichromate or horse serum.

Sections mounted on slides are incubated overnight in ovens set at 37-45°C and are ready
for staining the following day. They may be dried at 60°C for 15 minutes if they are required
the same day. Sections that readily detach such as nervous tissue, decalcified bone and very
dense tissue benefit from longer drying at lower temperatures.
 QUESTIONS PART1
Total 10 marks.
You should not spend more than an hour answering this. All the information has been
provided in labs and lectures.

1. List 3 chemicals that may be found in fixatives and make a comment about the safe
handling of each one. 1 mark

2. List 6 factors that affect fixation. 1 mark

3. What is the purpose of processing tissue through a) dehydration, b) clearing, and c) wax
infiltration? 1½ marks

4. Do microtome knives get blunt? If so, what do you do about it? ½ mark

5. Develop a short answer exam question worth 4 points, relevant to this topic, and supply
a list of the possible points expected in the answer. 2 marks

6. Using the processing schedules provided as a guide, design a processing schedule for a
large number of punch biopsies of skin. These samples are a cylinder 5 mm long and 2 mm
in diameter, with an epithelial surface at one end. Include both a solution and an immersion
period at each step. Use no more than 12 steps. 2 marks

1. 7.
2. 8.
3. 9.
4. 10.
5. 11.
6. 12.

7. What problems could be encountered in embedding and cutting the samples mentioned
above from Q 6? Suggest some solutions. 2 marks
 PART 2
STAINING

1. STAINING TECHNIQUES
AGITATE SECTIONS 10 TIMES IN EACH SOLUTION. For immersion times 2 minutes
or longer, agitate again before removing the slides.

DRAIN THE SLIDES WELL, BEFORE PLACING INTO THE NEXT SOLUTION.
FAILURE TO DRAIN WILL RESULT IN LOTS OF CONTAMINATION IN
FOLLOWING SOLUTIONS.

2. DEWAXING & HYDRATION

Before sections can be stained using aqueous stains they must be dewaxed. This is achieved
using toluene. The sections are then “taken to water” using graded alcohols and are
subsequently ready for staining.
This involves steps 1 – 6 of the Haematoxylin and Eosin method.

3. WATER WASHING
We have a two tier washing system where fresh water flows into the top container and then
makes its way into the lower container. USE THE LOWER CONTAINER FIRST TO
RINSE THE SLIDES AND THEN THE TOP CONTAINER TO WASH EFFICIENTLY.
Water should flow at a moderate rate and all traces of haematoxylin in the bottom container
should be gone by the time the slides have had their last wash before dehydration. If the
water refuses to obey gravity and flow down, the air bubble can be removed by elevating the
outlet tube a little until the water commences to flow.

4. DEHYDRATION AND CLEARING

Most methods contain instructions ‘dehydrate, clear and mount’. This involves steps 13 and
15 – 21 of the H&E staining method. Slides can be left in the final toluene before mounting
without affecting the staining.
Clearing refers to the action of these solutions to optically clear the sections making them
transparent, due to the solutions high refractive index.

5. MOUNTING
Sections are mounted using the smallest suitable sized coverslip and DPX.
Mounting not only protects the section from damage but is important in resolving the image
under the microscope. DPX is a plastic dissolved in xylene to a specific optical density,
rather than viscosity. To obtain the best resolution the optical density of the mounting
medium should equal, as closely as possible, the optical density of slide and coverslip, which
should also be equal.
Aqueous mounting systems are available but most have lower optical densities than glass,
and thus reduced resolution.
The DPX is dispensed from a small plastic bottle, located in the fumehood, onto a coverslip.

6. STORAGE
Slides should be appropriately labelled and then allowed to dry horizontally for at least a
week before storing upright in trays, files or boxes.
 HAEMATOXYLIN AND EOSIN STAINING
For Paraffin Sections

REAGENTS:
1. Harris’ Haematoxylin 500ml
Oxidant
Add oxidant to haematoxylin then invert 6-8 times or use a magnetic stirrer
for better mixing. Allow to stand for 8 hours prior to use.
2. 1% Acid Alcohol
Alcohol 70 ml
Distilled water 30 ml
Hydrochloric acid 1 ml
3. Alkaline rinse
Tap water 300 ml
Strong ammonia 28% 3 drops
OR
Lithium carbonate approx 0.15 g
Tap water 300 ml
4. Eosin Phloxine
Stock Eosin
Eosin Y 1.0 g
Distilled water 100 ml

Stock Phloxine Stock Phloxine

Phloxine B 1.0 g Phloxine 1.0g
Distilled water 100 ml Distilled Water 100ml

Working solution
Stock Eosin 40 ml
Stock Phloxine 10 ml
Ethanol 330 ml
Distilled water 20 ml
Glacial acetic acid 1.6 ml

REFERENCE:
1. `Manual of Histological Staining Methods of the Armed Forces Institute of Pathology.’ Lee Luna, editor,
3rd edition, McGraw-Hill Book Company 1968. P36

METHOD:

1. Toluene 2 min, drain. DEWAXING

2. Toluene 2 min, drain.

3. 100% Ethanol Agitate 10 times, drain.

4. 100% Ethanol Agitate 10 times, drain. HYDRATING
5. 70% Ethanol Agitate 10 times, drain.
*6. Tap water wash Agitate 10 times, drain.

7. Haematoxylin solution. 1 min, drain.

*8. Rinse well in running tap water.
9. Dip, quickly into acid/alcohol DON’T DRAIN. NUCLEAR
*10. Rinse in running tap water. STAINING
11.Alkaline rinse. agitate 10 times, drain.
*12. Rinse well in running tap water.

13. 70% Ethanol Agitate 10 times, drain. DEHYDRATING

14. Eosin 1 min CYTOPLASMIC STAINING

15. 100% Ethanol Agitate 10 times, drain.

16. 100% Ethanol Agitate 10 times, drain DEHYDRATING
17. 100% Ethanol Agitate 10 times, drain. CONTINUED

18. Toluene Agitate 10 times, drain.

19. Toluene Agitate 10 times, drain. CLEARING
20. Toluene Agitate 10 times.
21. Mount and coverslip in DPX. MOUNTING

Using a two tier washing system use the lower container first with 10 dips, then repeat the dips in the upper
clean water.
If staining is interrupted, sections may be left at any point up to step 6, any following water wash or the final
toluene.
Failure to wash well after the alkaline water at step 12 will result in changes to the pH of following solutions
and less than satisfactory staining for you and others who follow.

RESULTS:
Nuclei, ground substance blue
Cytoplasm, collagen, various shades of pink

VAN GIESON’S STAIN

SOLUTIONS:
1. Weighert’s Haematoxylin
Part A: haematoxylin powder 1.0g
absolute ethanol 100ml
Part B: 30% Ferric Chloride 4.0 ml
hydrochloric acid 1.0 ml
distilled water 95 ml
Mix equal parts of A and B just prior to use.

2. Van Gieson Solution – Modified Unna

Saturated aqueous picric acid 90 ml
Acid fuchsin 0.25 g
Nitric acid 0.5 ml
Glycerin 10 ml
Mix together and add a small amount of picric acid crystals to maintain saturation.

METHOD:
1. Take sections to water:
2. Weigert’s iron haematoxylin sol. 10 min.
3. Rinse in running tap water 2 min.
4. Van Gieson’s stain 3 min.
5. Blot sections carefully using a tissue or filter paper. Place into staining rack if more than 5 slides.
5. Dehydrate quickly through alcohols.
6. Clear and mount.

RESULTS:
Nuclei – brown to black.
Collagen – deep red.
Muscle, cytoplasm – yellow.

REFERENCES:
1. ‘Carleton’s Histological Technique’ Drury and Wallington , 4 th edition, Oxford Medical
Publications 1967 p132 and 166 –167
 STAINING COMPARISON OF DEMONSTRATION SLIDES

H&E

VVG

PAS

Alcian Blue/
VG
LFB/CFV

Reticulin Silver
Picro Mallory

Toluidine Blue
Orcein Elastic
Light Green

Haematoxylin &Eosin (H&E), Verhoff’s Van Gieson (VVG),

Periodic Acid Schiff (PAS), Alcian Blue/Van Gieson (VG),
Luxol Fast Blue (LFB)/Cresyl Fast Violet (CFV), Reticulin stain,
Picro-Mallory, Toluidine Blue
Orcein Elastic/Light Green.
PERIODIC-ACID SCHIFF (PAS)
For Paraffin Sections
SOLUTIONS:
1. Periodic acid
Periodic acid 1gm
Distilled water 100ml
7. Schiff’s Reagent
Pararosaniline or basic fuchsin certified for Schiff’s reagent 1.0g
Distilled water 200 ml
Potassium metabisulphite 2.0g
Hydrochloric acid 2.0ml
Activated charcoal approx 2 g
Bring the water to the boil and take off the heat prior to carefully adding the dye - this will avoid
premature renovation of the laboratory to a deep magenta colour. Mix the solution and cool to 50 0
C. Add the potassium metabisulphite in a fume hood. Mix again, further cool to room temperature,
and then add the acid, again in the fume hood. Mix in the activated charcoal and leave in a dark
cupboard overnight. Filter into a dark bottle and store refrigerated. The solution should be
colourless or pale yellow and should keep for about 6 months. 1.
3. Harris’ haematoxylin see H&E method
4. 1% Acid alcohol see H&E method
5. Alkaline rinse see H&E method

METHOD:
Take sections to water: See H&E steps 1 – 7.
Distilled water rinse well
1% Periodic acid solution 5 min.
Rinse well in running tap water
Schiff’s reagent 10 min.
Rinse in running tap water 5 min.
Haematoxylin 45 seconds, drain.
Rinse well in running tap water
Quickly dip in 1% Acid alcohol solution. DON’T DRAIN
Rinse well in running tap water
Alkaline rinse 10 dips, drain
Rinse well in running tap water.
Dehydrate, clear and mount (see H&E steps 13, and 15 – 21)

RESULTS:
Nuclei – blue/black
PAS positive material – magenta.
These include glycogen, mucin, reticulin, fibrin or thrombi, colloid droplets,
hyaline deposits in arteries, most basement membranes, pituitary and thyroid colloid,
fungi and amyloid infiltration 2.
REFERENCES:
1. ‘Theory and Practice of Histological Techniques’ Bancroft and Stevens 2nd edition, Churchill
Livingstone 1982. P188 – 190.
2. `Manual of Histological Staining Methods of the Armed Forces Institute of Pathology.’ Lee
Luna, editor, 3rd edition, McGraw-Hill Book Company 1968. P159 – 160.
 LUXOL FAST BLUE / CRESYL FAST VIOLET
Paraffin sections of central nervous system

REAGENTS:
1. Luxol fast blue
Luxol Fast Blue MBS and then add the acid. Filter. Solution is stable for 12 months or so. 1
2. Lithium carbonate
Lithium Carbonate 0.05g in 100ml of tap water
3. Cresyl Fast Violet
Cresyl fast violet acetate 0.1 g in 100ml in DDW plus 15drops 10% acetic acid 2
4. Cresyl Fast Violet Differentiator
1ml of glacial acetic acid in 100ml 95% alcohol

METHOD:
1. Dewax and bring to alcohol
2. Place slides in a plastic slide rack in a glass staining dish containing about 400ml of luxol fast blue solution.
3. Prewarm the magnetron of the microwave oven by heating about 500ml of tap water (don’t use distilled) for about
2 minutes. Remove the water. 3
4. Using a temperature probe programme the microwave (650W) to heat to 62 oC using 70% power, and then to
maintain the temperature for one minute. Fit the temperature probe and place into the solution. Cover loosely with a
lid and microwave. Avoid inhaling the vapours. Smaller staining volumes may be used successfully but the solution
tends to escape the container and make a mess!
5. Allow the slides to sit at room temperature for 5 minutes while solutions are prepared.
6. Rinse the slides in tap water
7. Commence the differentiation by dipping the slides into the lithium carbonate 3 times. Drain.
8. Continue differentiation by dipping slides into 70% alcohol 3 times.
9. Stop the differentiation in running tap water.
10. Repeat steps 7-9 until the grey matter (the part containing the neurons) is unstained and the white matter (the
part containing the myelinated fibres) is still blue. If overdifferentiated dehydrate and return slides to dye solution.
Start again at step 2.
11. When differentiation is complete wash well in running tap water.
12. Stain in cresyl fast violet for 2 minutes.
13. Differentiate for 2 dips. This stage is fairly quick.
14. Proceed quickly through dehydrating alcohols, which will complete the differentiation.
15. Clear and mount.

RESULTS:
Myelin blue
Nuclei purple
Nissl granules blue/purple

REFERENCES
1. `Carleton’s Histological Technique’ Drury and Wallington , 4 th edition, Oxford Medical Publications 1967 p267
2. `Manual of Histological Staining Methods of the Armed Forces Institute of Pathology.’ Lee Luna, editor, 3 rd edition, McGraw-Hill Book
Company 1968. P203 – 204.
3. `Microwave Cookbook for Microscopists’ Kok and Boon revised 3rd edition, Coulomb Press, Leydon 1992. P106-107, 188-191

NOTES:
 We use Gurr’s Luxol Fast Blue dye. The bottle we are currently using is probably at least 10 years old! The lot
number is 87562
 Cresyl fast violet acetate does not have a CI number but we are using certified dye from Aldrich. Provided it is
certified by the Biological Stain Commission any manufacturer should do.
 Kok and Boon give 2 methods for LFB but I developed this independently.
 We have an old Phillips model 5004e 650W microwave with temperature probe. Other ovens could be used
satisfactorily if there is temperature control. Take care not to overheat the solution since it is flammable!
 The slides can be left at step 9 if staining is to be interrupted.
 Questions Part 2

Total 10 marks.
You should not spend more than an hour answering this. Most of the information has been
provided in labs and lectures.

1. Why are slides dewaxed prior to staining? ½ mark

2. Which tissue components are dyed by a) haematoxylin and b) eosin? 1 mark

3. Coverslipping is performed in a fume hood. Explain why? ½ mark

4. A Van Gieson stain uses a different haematoxylin solution to the routine H&E stain.
Compare and contrast the two haematoxylin solutions including their usefulness in H&E and
Van Gieson stains. A table might be useful. Compare more than their chemical components.
2 marks

5. What are the bright pink substances in the stomach of the PAS stained section?
1 mark

6. If a VG was staining both muscle and collagen mostly yellow or mostly red, what could
you vary to improve the staining results? 2 marks

7. Develop a short answer exam question worth 4 points, relevant to this topic, and supply a
list of the possible points expected in the answer. 2 marks

8. If you stained a section of skin with a VG stain, what colour would you expect the
following components to dye: Dermis
Epithelium
Arteriole wall
Fat cell 1 mark