Parazitologie LP

Slide 1Methods in Parasitology
1. Preparing thin and thick blood films with capillary or venous blood
Swiss Tropical Institute, Basel
Contents
Yvette Endriss, Elisabeth Escher & Birgit Rohr Prof. Hanspeter Rohr (NeoCortex Foundation) Prof. Niklaus Weiss (STI)
1.1 Preparing thin and thick blood films
• •
Prepare the microscope slides
Use microscope slides with or without frosted end
Finger-prick
(capillary blood)
With the patient's left hand, palm upwards, select the third finger. (The big toe can be used with children. The thumb should never be used for adults or children). Use cotton wool lightly soaked in alcohol to clean the finger, using firm strokes to remove grease from the ball of the finger. Let finger air-dry.
Finger-prick
(capillary blood)
With a sterile lancet puncture the ball of the finger using a quick rolling
action.
Clip
Finger-prick
(capillary blood)
By applying gentle pressure to the finger express the first drop of blood and wipe it away with dry cotton wool. Make sure no strands of cotton remain on the finger.
Finger-prick
(capillary blood)
Working quickly with capillary blood and handling clean slides only by the edges, collect the blood as follows: Apply gentle pressure to the finger and collect a single small drop of blood about the size • on the end of the slide. This is for thin film.
Thin films (capillary blood)
Using another clean slide as a "spreader", and with the slide with the blood drops resting on a flat, firm surface, touch the small drop with the spreader (1) and allow the blood to run along its edge. Firmly push the spreader along the slide (2), away from the drops, keeping the spreader at an angle of 45°. Make sure the spreader is in even contact with the surface of the slide. (Look at clip on next slide)
Left hand
Thin films
(capillary blood)
Observe right angle!
• Angle too flat
> film too long
• Angle too steep
> film too short
Mistakes:
> „waves"
Thick films
(Capillary blood)
Apply gentle pressure to the finger and collect two larger drops, about a size • , on the slide as shown in the upper picture.
Handle the „spreader" by the edge, using the corner to spread the blood in a circular form with 3-6 movements.
Thick films
Thick films
You should be able to read the newspaper!
Thick films
Additional mistakes
Labelling
Labelling
Drying
Allow the thin film and the thick film to dry in a flat level position protected from flies, dust and extreme heat.
Drying
Optional:
For example:
Drying
(Collects the dust)
These slides look o.k.!
These slides look more like art work than useful blood slides!
Venous blood can be used instead of capillary blood
Use vacutainers with anticoagulant (EDTA)
For preparing thin and thick films use a glass capillary to drop the ETDA-blood.
Do not use a plastic pipette!
Clip
Spare glass slides
Combination of a thin and a thick film on the same slide.
Spare glass slides
Combination of a thin and a thick film on the same slide.
Methods in Parasitology
stain
Contents
• 2.5 Preparing Giemsa stain
• 2.10 Staining blood films
2.1 Staining blood films with Giemsa
For perfect malaria staining, thin and thick films should be made on separate slides. The pH of the buffer solution should be 7.2 .
Thin films will be fixed with Methanol.
Thick films should not be fixed (to allow haemolysis)!
Therefore avoid exposure of the thick films to methanol vapour when thin and thick films are on the same slide.
Materials:
(Giemsa powder can also be used instead of Giemsa solution)
Fixation for thin films only!
Fixation time for thin films with methanol:
15 - 30 seconds
For perfect malaria staining the pH should be correct! (pH 7.2)
Commercially available buffer tablets can also be used.
Prepare your buffer by using tablets of Soerensen (see next slide)
Combine two stock solutions in a certain proportion
KH2PO4 and
Stock solutions:
A: KH2PO4 (Merck Art. 4873) 9.08 g/l = 1/15 molar
B: Na2HPO4 2 H2O (Merck Art. 6580) 11,87 g/l = 1/15 molar (or Na2HPO4 anhydricum (Merck Art. 6586) 9.47 g/l = 1/15 molar)
Buffer by Soerensen
Prepare staining solution:
Then add
Mix 94 ml buffer solution ph 7,2 with
6 ml Giemsa stock solution >>
6% Giemsa solution
Place thin film and thick films into the staining dish.
Stain blood slides for 45 minutes
(staining a large number of thin films and thick films use two separate staining dishes)
Thin films:
Caution:
(since they are not fixed before staining)
Mistake:
Drying
37°c)
a) Thin films
Contents
Yvette Endriss, Elisabeth Escher & Birgit Rohr Prof. Hanspeter Rohr (Neocortex Foundation) Prof. Niklaus Weiss (STI)
3.1 Field's stain for thin films
A: Thin films
Fixation:
Dry microscopic slide on filter paper
Immerse slide in Field's stain B (Eosin) for 5 seconds
Immediately wash with tap water!
Immerse slide in Field's stain A (Methylene blue) for 10 seconds
Immediately wash with tap water
Dry thin films
B: Thick films
• Tube with water
3.10 Field's stain for thick films
Immerse thick film in Field's stain A (Methylene blue) for 3 seconds
Do not forget: Thick films need to be haemolysed and are therefore not fixed with methanol!
Rinse immediately in tap water
Immerse thick film in Field's stain B (Eosin) for 3 seconds
Then rinse immediately with tap water
Let the slide carefully dry
Solution
4.0 Concentration Method for Microfilariae
Contents
Yvette Endriss, Elisabeth Escher & Birgit Rohr Prof. Hanspeter Rohr (Neocortex Foundation) Prof. Niklaus Weiss (STI)
Materials:
Fill 2 centrifugation tubes with 1 ml of anti- coagulated blood
Add 9 ml of a 2% Formalin solution to each tube
(final dilution: 0.2%)
Wait 5 minutes for the red cells to haemolyse
Mistake:
Centrifugation at 2000 rpm for 3 minutes
After centrifugation:
Decant the supernatant
Add a drop of Methylene-blue to one sediment and mix carefully.
(for better visibility of the microfilariae)
Prepare stained sediment for microscopy. Search and count all microfilariae in the whole sediment under a cover slip (Objective 10x)
4.11 Haematoxylin stain for
If microfilariae are found:
Prepare thick film from the second sediment (containing no Methylene- blue) for identification by Delafield's haematoxylin staining
4.12 Haematoxylin stain for Microfilariae
Carefully dry the thick film
Fix the thick smear with Methanol
Dry the thick smear
Stain the thick film with filtered Delafield's haematoxylin for 30 minutes
Intensify the blue colour by immersing the slide in tap water for 10 minutes and let the thick smear dry afterwards.
5.1 Fresh Stool Examination
• Wooden stick
• Fresh stool
stick
Press cover slip slightly, remove excess liquid with paper towel
Clip
Example:
(*SAF: Sodium acetate-acetic acid-formalin solution)
• 6.8 SAF-Ether concentration
6.1 Sedimentation
Conical glass / plastic bottles Wire mesh
Microscope slides, cover slips Gloves
6.2 Sedimentation
Carefully mix fresh stool with saline in a glass beaker
6.3 Sedimentation
Filter stool suspension through a sieve with two layers of gauze into a conical glass
6.4 Sedimentation
6.5 Sedimentation
Carefully pour off the supernatant fluid until the sediment is visible
6.6 Sedimentation
Take up part of the sediment using a Pasteur pipette. Leave to stand for 2 - 3 minutes (Mini-sedimentation in the pipette)
6.7 Sedimentation
Search for helminth eggs and larvae
6.8 SAF-Ether Concentration
Pour the rest of the sediment into a conical centrifugation tube
(Caution: inflammable!)
ETHER
Remove all layers except the sediment using a pipette on a pump
(Alternatively: loosen the detritus with a wooden stick and decant)
Mix sediment and place a drop on a microscope slide. Add cover slip
Examine the whole sediment
(*SAF: Sodium acetate-acetic acid-formalin solution)
Contents
7.1 SAF method for stool specimen
Materials:
Place stool sample (approx. 1g, size of a hazelnut) into a tube containing 10 ml of SAF
Mix stool thoroughly
Agitate tube vigorously
Filter stool solution using a funnel with gauze
Centrifuge for 1 minute at 2000 rpm
Remove supernatant with a pipette
or
Add 7ml saline
Close tubes with rubber stoppers, shake well
keeping the thumb on the stopper
Remove rubber stopper carefully (pressure!) and centrifuge tubes for 5 minute at 2000 rpm.
After centrifugation: 4 layers are detectable
ETHER
Remove 3 layers (Ether/Detritus/NaCl) with a pipette
or
Sediment should be less than 1 ml (left side)
(if more (right side): repeat the concentration with saline/ether)
Mix the sediment and place a drop on a microscope slide
Add cover slip
Slide ready for microscopy: Search first for helminth eggs (10x objective).
Then, for protozoa, press cover slip slightly, remove excess liquid with paper towel and use 50x objective with oil immersion
Contents
8.1 KATO-Katz technique for helminth eggs
Materials:
Kato-set (Template with hole, screen, nylon or plastic, plastic spatula)
• Newspaper or glazed tile Microscope slides Cellophane as cover slip,
soaked in Glycerol-malachite green solution
• Fresh stool Gloves
Prepare the layer
Glazed tile or newspaper
Place the template with hole in the centre of a microscope slide
Use gloves !
Place a small amount of faecal material on the newspaper or the glazed tile.
Press the screen on top so that some of the faeces filters through and scrape with the flat spatula across the upper surface to collect the filtered faeces.
Add the collected faeces in the hole of the template so that it is completely filled.
Remove the template carefully so that the cylinder of faeces is left on the slide.
Cover the faecal material with the pre-soaked cellophane strip.
Invert the microscope slide and firmly press the faecal sample against the cellophane strip on a smooth hard surface such as a tile. The material will be spread evenly.
Carefully remove the slide by gently sliding it sideways to avoid separating the cellophane strip. Place the slide with the cellophane upwards.
The smear should be examined in a systematic manner and the eggs of each species reported. Later multiply by the appropriate number (see inlet-information of the Kato-set) to give the number of the eggs per gram faeces.
of pinworm eggs
9.1 Adhesive tape method
Stick tape on microscope slide
4-6 negative tapes are needed to rule out a pinworm infection!
Clip
Place a drop of Xylene on the edge of the tape to remove air bubbles

Documents

Parazitologie LP