Histology and histochemical staining: Histology and histochemical staining, an alternative method to the more classical formalin fixation

Histology and histochemical staining: Histology and histochemical staining, an alternative method to the more classical formalin fixation 150 150 Affordable Capstone Projects Written from Scratch

Lab exercise block 3
Histology and histochemical staining
In this lab exercise, the candidate will get familiar with the cryopreservation of
samples for cryosectioning of muscle tissue.
This is an alternative method to the more classical formalin fixation followed by
wax-embedding techniques and sectioning in a microtome.
Fresh salmon from the research station will be used as test material for this lab
1. Note down the following measurments of the fish.
a. Length, weight & sex
2. Standardize the sampling location for the steak.
a. The steak is sampled anterior of the dorsal and adipose fin.
 Two steaks/fish (or more just for the practice).
3. Cut a 5-7mm thin streak using a sharp knife.
4. From the steak prepare blocks of 5x5x5mm size. See fig.1
5. Place the blocks on top of a piece of cork (pre-labelled). Mount with
cryofix (cryomatrix). Take all samples from both steaks the two fish.
When all samples are taken they need to be cryofixed.
6. Pour isopentane in a steal cup and fill the nitrogen bucket with liquid N2
(assisted by Ørjan/Anjana).
7. Gently lower the steel beaker into the liquid N2. Stir continuously until
the isopentane is cooled down to the freezing point. Start to freeze around
the edge of the steal cup. Refill N2 when necessary.
8. Gently grab hold of the cork sontaining the sample using a large tweezer
and lower the sample into the super-cooled isopentane.
9. Hold it for a minimum of 45s is single frozen sample for double or triple
frozen samples, hold it for 60s.
When sample is frozen quickly remove the sample from the isopentane
and wrap it in pre-labelled alu-foil and store in a liquid N2 container.
Repeat for all samples.
When all samples are taken they can be transferred to the – 80 freezer and
stored until they will be cut in the cryostat. Alternatively they can be transferred
directly to the cryostat. After samples are transferred to the cryostat they need to
acclimatized to the process temperature in the cryostat. This is normally between
-18 to -24⁰C.
The cryosectioning will be demonstrated and training given on the
lab exercise. This will be performed group vice according to the
plan for the lab exercise. This part will be under supervision from
Anjana and or Ørjan.
Fig. 1. Illustration of how many samples that is needed for different sized fish.
Preparation of Poly-L-lysine solution.
Slides are treated with Poly-L-Lysine solution to improve
adhesion of section to the microscope slide. This is necessary so
that the slides won’t come off during staining and/or washing.
1 l Poly-L-Lysine working solution consists of:
 900 ml distilled water.
 100 ml Poly-L-Lysine solution.
1 l of Poly-L-Lysine solution is enough to treat 900 microscope
slides. Stock and work solution is stored in the fridge in the
histology lab.
1. The microscope slides are placed in the plastic rack which
is located in the cupboard beside the cryostat.
2. The microscope slides are covered with the Poly-L-Lysine
solution and left there for 5 minutes. It is important that the
whole rack is covered. If not make up more solution.
3. Remove racks after 5 minutes and leave on them on the
bench for drying until next day, cover racks with paper.
4. After drying (next day) the slides are marked, to avoid
mixing of treated and untreated slides.
 It is important that the Poly-L-Lysine working
solution bottle is properly marked on the front label
indication how many slides that are left for Ploy-LLysine
Ørjan Hagen
Staining section with Harry’s Hemotoxylin.
Harry’s hemotoxylin is a general nuclei staining often used in the
staining of histological muscle sections since it gives a good
contrast to the surrounding pericellular connective tissue. It
therefore makes it easier to visualise the muscle cells.
1. Place the microscope slides in the staining jar (back to back).
2. Pour Harry’s Hemotoxylin into the jar and let slides develop
for 5-10 min (normally 7-8 min is sufficient, depends on
3. After development, pour off Harry’s Hemotoxylin (recycling,
into the bottle) and leave staining jar under the tap for 10
min for further development (cold water, gentle jet).
4. Mount slides with glycerol gelatine. Place one drop of
glycerol gelatine on top of section and cover it with a cover
glass. Press gentle on top of cover glass (tweezer/spoon) to
get rid of bobbles that might be present. PS. This has to be
done quickly (within 30 sec) or the glycerol gelatine will
PS! Us gloves at all time and do not let sections be exposed to air
for a too long time after staining/before mounting (sections may
dry and shrink).
Ørjan Hagen
Succinate Dehydrogeniase (SDHase) Staining
The SDHase enzyme is a marker for oxidative metabolism in
mitochondria. Electrons are produced when enzyme reacts with its specific
substrate, succinate. In this technique the electrons are passed on to the
ditetrazolium salt (nitro-blue tetrazolium) which forms a purple complex
that precipitates at the site of the reaction.
Solution preparation (prepared in advance of lab exercise)
50mM Phosphate Buffer/80mM Sodium Succinate Solution
1. Make up a 0.1M solution of di-sodium hydrogen orthophosphate
(Na2HPO4) by dissolving 14.4g (18g if hydrated) in 1 litre of
distilled water.
2. Make up a 0.1M solution of sodium di-hydrogen orthophosphate
(NaH2PO4) by dissolving 12.0g (15.6g if hydrated) in 1 liter of
distilled water.
3. Using a magnetic stirrer and pH meter, mix the two solutions to
achieve a pH of 7.6 (stock solution).
4. Measure volume and add the same quantity again of distilled
water (working solution).
5. Add 21.6g/l of sodium succinate (succinic acid)
6. Re-adjust pH to 7.6
Procedure to follow :
1. Using 7 μm sections on poly-L-lysine slides (prepared earlier in lab)
2. Add 2mg/ml nitroblue tetrazolium (MBT) to an aliquot of phosphate
buffer/sodium succinate solution (prepared solution above) (allow
approximate 750μl per slide)
3. Flood slides and incubate in the dark for 1-2 hour.
4. Wash in distilled water
5. Mount in glycerol gelatin
The incubation period differs according to the type of tissue used, and
probably with the thickness of the section. 1 hour is suitable for 7μm cod or
salmon sections. Too short an incubation leads to inadequate staining; a
prolonged incubation leads to the formation of bubbles in the tissue.
Ørjan Hagen