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1. Module 4
Specimen Collection and Preparation
of Blood films; preparation of stains;
and blood film staining
1

2. Specimen Collection and Preparation
of Blood films
2

3. Learning Objectives
At the end of this module , the learner should be
able to:
– Describe the types of blood films used in routine
malaria microscopy
– Collect capillary blood for the preparation of thick
and thin blood films.
– Make a good thick and thin films of blood taken from
people with suspected malaria in a single slide
3

4. Learning …
– Dry the blood films appropriately
– Correct labeling of blood films
– Identify good quality of thick and thin blood films
– Identify the causes of common mistakes in the
preparation of both thick and thin blood films
4

5. Content Outline
– Materials required
– Types of blood films
– Blood sample collection
– Blood film preparation
– Qualities of good thick and thin films
– Common mistakes in making blood films
5

6. Materials Required
6

7. Types of blood films
1. Thick films
– Consist of a thick layer of lysed RBCs .
– Used to detect and quantify malaria parasite
– More sensitive than thin blood film
• The blood elements are more concentrated (~ 30x)
than in an equal area of a thin film.
– Best for detection of low levels of parasitemia.
– Used for rapid detection of the parasite
Challenge
– Does not permit an optimal review of parasite
morphology.
7

8. 2. Thin films
– Consist of blood spread in a single layer.
– In fixed thin film the parasites are found intact inside the
RBC.
– Assist in the identification of the Plasmodium species
– Provides greater specificity
Challenges
– low-density infections can be missed
– Requires more time to read.
8

9. Preparation of Slide
– Use new slides with frosted end to make patient
identification easy.
– Slides must be free of scratches, grease, dust, and
acid or alkali contamination.
– Store slides in a closed dry container.
9

10. Blood Sample Collection : Capillary Blood
• The ideal sample as the density of trophozoites or schizonts is
greater in blood from capillary-rich area.
• Obtained by pricking a fingertip or big toe
• For adults:
 the lateral side of the 3rd or 4th finger is best.
• For infants :
 the big toe is preferred.
10

11. Procedure for collecting finger prick blood
1.Wear gloves during the procedure
2. Keep complete record of patient
identification in a logbook.
3. Prepare pre-cleaned slides in a
clean surface. Handle slides by the
edges to avoid fingerprints.
11

12. Procedure…
4. Select the 3rd or 4th finger and
clean the area with a alcohol swab
and Allow to dry.
5. Puncture using a sterile lancet.
6. Apply gentle pressure to allow blood
drop to ooze out. Wipe away the first
drop of blood
12

13. Transferring of blood to slide: For thin film
– Applay gentle pressure to the finger and collect a single
small drop(2µl) of blood in the centre of the slide.
– Release the pressure immediately to allow recirculation
of blood.
13

14. For thick film
• Apply gentle pressure again to transfer more
blood and collect one bigger (6µl )drop on the
slide, about 1 cm from the drop intended for the
thin film or 1 cm from the end of the slide.
14

15. Thick and thin blood films are made as follows:
Thin film
• Take clean slide with smooth edges and use it as a
spreader.
• Hold the end of a spreader slide against the surface of
the first slide at an angle of 30-45 degree.
• Make thin film by bringing in contact with edge of the
spreader with the blood drop on the middle of the original
slide.
• Wait until the blood spreads along the entire width of the
spreader
• Push it forward rapidly and smoothly until the film is
formed.
15

16. Thin film…
• Make sure that the spreader is in even contact with the surface
of the slide all the time the blood is being spread.
• If the blood is anemic, hold the spreader at a greater angle to
make the blood film slightly thicker.
• The spreader slide may now be used for the next patient.
16

17. Thick film
• Using the corner of the spreader,
quickly join the drops of blood and
spread in a circular/rectangular
manner with 3 to 6 movements to
make a round
• Do not make the film too thick or it
will fall off the slide.
17

18. Preparation of thin and thick film
18

19. Qualities of Good Thin Blood film
– Uniformly spread over the slide
– Thin enough so that It is tongue shaped
– Consists of a single layer of RBCs with feathered end
19

20. Qualities of Good Thick Blood film
– It should be 10 mm away from the edge of the slide
– Rectangular or round in shape with a diameter of about 10 mm
– Its thickness contains 10 layers of RBCs
– At least 10-12 WBCs should be visible / 100x field.
20

21. Common Mistakes in Making Blood Films
1. Too much blood
• On thick film
– After staining the background will be
too blue.
– Result too many WBCs per thick film
field, and these could obscure malaria
parasites .
• On thin film
– RBCs will be on top of one another
– impossible to examine them properly
after fixation.
21

22. Common mistakes
2.Too little blood
• Not able to examine
enough blood in the
standard examination.
• False negative result is
likely.
22

23. Common mistakes…
3.Blood films spread on a
greasy slide
• The blood films will spread
unevenly .
• Some of the thick film will
probably come off the slide
during the staining
23

24. Common mistakes…
4. Edge of spreader slide
chipped
• the thin film spreads unevenly,
is streaky and has many “tails”.
• The spreading of the thick film
may also be affected.
24

25. Common Mistakes…
5.Badly positioned blood films
• Blood films should be correctly sited
on the slide.
• If not, it may be difficult to examine
the thick film.
25

26. Common mistakes…
6. Thin film too big, thick film in the wrong place
• The thick film will be out of place and may be so near the edge
of the slide that it can’t be seen through the microscope.
• During staining or drying, portions of the thick film will scraped
off by the edges of the staining trough or drying rack.
• Very difficult to position the thick film on the microscope stage.
26

27. Common technical mistakes in collection and
preparation of blood film
27
Mistake Effect
Pricking of not well dried finger
• The parasites and host cells may be
fixed by the alcoholic detergent
solution
Use of dirty slides
• The blood will not be spread evenly.
• Generates artifacts
Delay in making blood film once
you transfer the drops of blood
to the slide
• The blood film will not be spread
evenly due to the beginning of the
coagulation process
Too much blood for thin films
• Erythrocytes are laid on multiple
layers.
• Observation is impossible

28. Common mistakes…
…
28
Mistakes Effects
Too little blood used for thin
films
• Parasites may be virtually
absent if parasitaemia is low
Not labeling slides
immediately
• Confusion may arise leading
to unidentified positive or
negative slides
Slides wrapped together
before thick films properly
dried
• The slides stick to one
another

29. Common mistakes…
Mistake Effects
Inappropriate washing of stain from
the slide
• Stain deposits may render the
observation difficult
Long time elapses before staining of
thick films
• Causes Auto fixation which result in
difficulty of hemolysis during staining
Exposure of thick films to excessive
heat
• Autofixation occurs and haemolysis is
Impossible
29

30. Preparation of Stains and Buffered
water
30

31. Learning Objectives
At the end of this module the trainees should be
able to
– Prepare Giemsa staining solution required for
malaria microscopy.
– Describe the importance of using quality
chemicals for reagent preparation.
– Describe how to safely store Giemsa stain.
31

32. Content Outline
– Principles of Romanowsky stains
– Preparation of Giemsa stain
– Storage of Giemsa stain.
32

33. Romanowsky stains
– Polychrome stain
– Used to stain blood cells and organisms.
– Comprise two staining components:
– Eosin Y and Azure B, obtained by oxidation of
methylene blue.
• Eosin stains chromatin dot and stippling shades of
red or pink.
• Methylene blue, stains parasite cytoplasm blue.
33

34. Romanowsky stains: Examples
– Giemsa stain
– Field’s stain
– Leishman’s stain
– Wright’s stain…
34

35. Giemsa stain
• Gemsa stain is an alcohol-based Romanowsky stain.
• Is a mixture of eosin, which stains parasite
chromatin and stippling shades of red or pink, and
methylene blue, which stains parasite cytoplasm
blue.
• White-cell nuclei stain blue to almost black,
depending on the type of white cell.
• The recommended stain for identification of
malaria parasite.
35

36. Giemsa stain
Reagent and materials required
– Giemsa Powder
– Absolute methanol
– Glycerol
– Glass beads 50 pcs
– Brown bottle
– Analytical balance
36

37. Preparation of Giemsa Stain
• To make about 500 ml:
– Giemsa Powder 3.8g
– Absolute methanol 250ml
– Glycerol 250ml
37

38. Procedure…
1. Weigh the Giemsa and transfer to a dry brown bottle of
500 ml capacity which contains a few glass beads.
2. Using a cylinder measure the methanol and add to the
stain. Mix well.
3. Using the same cylinder measure, the glycerol and add to
the stain. Mix well.
38

39. Procedure…
4. Tightly stopper the bottle.
5. Shake the bottle for 2-3 minutes.
6. Add measured glycerol and repeat the shaking.
7. Continue shaking for 2-3 minutes at 30 minutes intervals
at least 6 times.
8. Keep the bottle for 2-3 days; shaking it 3-4 times each
day.
9. Keep small amount of the stock in a small bottle.
10. Label with the name of the reagent and date of
preparation and mark “inflammable”. Store at room
temperature in the dark.
39

40. What you should do after preparation of
Giemsa stock solution
• Keep the stopper screwed tightly.
• Must be diluted with distilled water (Buffered water)
with pH of 7.2.
• Should be tested for proper staining reaction.
• Must be Protected from Moisture and direct sunlight.
• Stored in a cool dry place in a dark bottle.
• Measure a small quantity of stain into a smaller bottle for
one or two days’ use.
40

41. What you should not do after the reagent
preparation
• Never add water to the stock Giemsa solution.
• Do not shake the bottle of stain before use: you will
re-suspend very small, un dissolved crystals of stain.
• Never return unused stain to the stock bottle.
41

42. Quality controls of Giemsa stain
• Done to ensure the staining quality and performance of
Giemsa stain.
• Use known positive and negative films with each new batch
of working Giemsa stain.
• Blood films can be prepared using EDTA anticoagulated
blood from a patient’s.
Note : An ideal blood sample has at least one parasite in
every 2–3 fields.
42

43. Quality control …
• Allow the blood films to dry quickly.
• Fix the films using absolute methanol.
• Place them, touching back to back, in a box with
separating grooves.
• Label the outside of the box with the species, date
and “Giemsa control slides”.
43

44. Quality control …
• The slides can be stored at room temperature for a
minimum of 1 week but will last longer if stored at -200C
or below –70 °C.
• Just before use, remove the slide from the box and allow
the condensation to evaporate.
• Label the slide with the date and “Positive control”.
• The blood film can then be stained and examined.
44

45. Buffer solutions for malaria staining
• A phosphate buffer solution, balanced to pH 7.2 is
important to prepare Giemsa working solution.
• Check the pH using narrow range pH papers or pH
meter and store at room temperature.
• The buffer is stable for several months.
45

46. Materials, Reagents and Equipment Required
Materials
1. Wooden spatula
2. Beaker 250 ml capacity
3. Conical flask 1000ml
capacity
4. Filter papers
5. Brown bottle
6. Measuring cylinder 100ml
capacity
7. Filter papers
8. Labels
Reagents
1. KH2PO4 0.7g
2. NA2HPO4 1.0g
3. Distilled or de-ionized
water 1000ml
Equipment:
• Analytical Balance
46

47. Preparation of buffered water : Procedure
1.Measure 0.7g KH2PO4 and 1g of Na2HPO4.
2. Add 0.7g KH2PO4 to the beaker.
3. Add 150ml of water.
4. Stir with the spatula until the salt is dissolved.
5. Add 1.0g of Na2HPO4 to the beaker.
6. Stir with the spatula.
47

48. Procedure…
48
7. When dissolved add the fluid from the beaker to the
conical flask.
8. Fill the fluid in the conical flask with water until it is made
up to 1ml.
9. Add small quantities of 2% Na2HPO4 if the pH is below 7.2
(acidic) or 2% KH2PO4 if the pH is above 7.2 (basic).
Quality Control
• Check pH of buffered water, and add appropriate
correcting fluid.

49. Buffer tablet
– Buffer tablets that produce a solution of pH 7.2
when dissolved are ready available from
laboratory suppliers but are rather expensive.
49

50. Preparation of 2% correcting fluids
• Used to maintain the Buffer
at pH 7.2 .
Materials
1. Wooden spatula
2. Beaker 250 ml capacity
3. Conical flask 1000ml capacity
4. Filter papers
5. Brown bottles
6. Measuring cylinder 100ml capacity
7. Filter papers
8. Labels
Reagents
1. KH2PO4, 2g
2. NA2HPO4, 2g
3. Distilled or de-ionized water, 1000ml
Equipment
Analytical balance
50

51. Preparation of 2% correcting fluid: Procedure
– Weigh 2g NA2HPO4 and add it to 100ml of water in the
beaker.
– Stir with the wooden spatula.
– Pour the solution into one of the glass bottles.
– Label the bottle “2% disodium hydrogen phosphate”
– Repeat steps 1 and 2 above weighing out 2g of KH2PO4
– Pour the solution into the second glass bottle and label it
correctly.
51

52. Quality Control of Buffered water
– Prepare a buffer reagent carefully.
– Check the PH regularly.
– Store buffered reagents at 2-80C.
– Avoid leaving the reagents in sun light (encourages the
growth of algae).
– Check for contamination at regular intervals .
52

53. Fixation and Staining of Blood
Film
53

54. Learning Objectives
At the end of the unit, the learner should be able to:
– Demonstrate appropriate blood film fixation and
staining techniques
54

55. Content Outline
– Blood film fixation
– Staining blood film
55

56. Fixation of Blood film: Materials
– Absolute Methanol
– Dropper bottle
– Absorbent cotton wool/Gauze
– Drying rack
56

57. Fixation
• Used to preserve cellular and parasitic morphology.
• Done When the blood films are completely dried.
• Absolute methanol is recommended fixative solution
Note: Auto-fixation may also occur spontaneously with time if thin
films not fixed immediately after dried (7 to 15 days, varying with
humidity and temperature of the atmosphere).
57

58. Fixation methods
Done for 10-20 seconds using one of these three methods:
1. Dropping absolute methanol while holding the slide
with the thin portion down
2. Dipping in a jar containing absolute methanol.
3. Dabbing it with cotton wool dampened with methanol
Note: Avoid methanol, or its fumes encountering the thick film.
58

59. Staining blood films
There are two methods of staining with Giemsa stain:
1. The rapid (10%) method
2. The slow (3%) method.
• The rapid method is used in laboratories where a quick
diagnosis is an essential part of patient care.
• The slow method is used for staining larger numbers of slides,
such as those collected during cross-sectional or
epidemiological surveys and field research.
59

60. Staining Blood Film
1. The rapid (10%)methods
• Filter the stock Giemsa solution
• Prepare fresh a 10% (1:10 ratio) working Giemsa stain solution
– Add 10ml of Giemsa stock solution to 90 ml of buffered
distilled water or 5ml to 45 ml of buffered distilled water
• Pour the diluted solution into a staining jar
• Immerse completely dried slides in a jar.
• Make sure that the stain cover the entire surface of films on the
slide.
60

61. Staining Blood film
• Leave the slides in the stain
for minimum of 10 minutes.
• Wash the stain in a jar
containing water.
• Remove the slides and
clean the back of each
slide with dry gauze.
61

62. Staining …
• Place the slides with the film side down wards in a
drying rack to drain and dry at room temperature.
• Make sure that the thick film does not touch the edge
of the rack.
62

63. Staining of blood films
The slow (3%) method
• This method is less appropriate when a quick result is needed but is
excellent for staining large numbers (20 or more) of slides.
• It is ideal for staining blood films from surveys or research work or
batches of slides for teaching.
• It performs best when slides have dried overnight.
• The method is economical because much less stain is used (3%
rather than 10%).
63

64. Quality Control
• A blood film, thick and/or thin, is run once each
month or with each staining procedure
• The blood film does not necessarily required to be
positive for blood parasites.
• Good color differentiation of red and white cells is an
indication of a good quality stain.
64

65. Field’s stain Preparation and staining
• Useful for rapid detection of malaria parasites
particularly for thick films.
• Schuffener’s dots are not always stained with field’s
stain.
• Made up of Field’s stain A and Field’s stain B.
65

66. Preparation of Field’s A stock solution
 Can be prepared in two different ways:
I. From prepared powder
1. Add 0.5 g of fields satin A powder to 600ml of
hot (~ 600C ) distilled water.
2. Mix until it dissolves.
3. Filter when cool.
66

67. II. From original stains and chemicals
1. Dissolve 10.0 g of anhydrous Na2HPO4 and 12.5 g of
KH2PO4 in 1000 ml of distilled water.
2. Pour half of this solution into 1 liter bottle
containing a glass beads. Add 1.6g of methylene
blue and 1.0 g of azur 1 and mix well.
3. Add the remainder of the phosphate solution
4. Mix well and filter
67

68. Preparation of Field’s stain B- stock solution
 Prepared in two different ways,
From prepared powder
1. Add 4.8 g Field’s stain B powder to 600ml of hot
(~60 0C) distilled water.
2. Mix until dissolved
3. Filter when cool.
68

69. From original stains and chemicals
1. Dissolve 10.0 g of anhydrous Na2HPO4 and 12.5
g of KH2PO4 in 1000 ml of distilled water.
2. Add 2 g of eosin (yellow, water soluble).
3. Mix until dissolved.
4. Filter.
69

70. Field’s staining Procedure for staining thin films
1. Fix film in methanol for 1 minute
2. Wash off methanol with water.
3. Using a pipette, cover the film with diluted Field’s
stain B (1 part by volume of stock solution plus 4
volumes of distilled water buffered at pH 7.2)
70

71. Procedure…
1. Immediately add an equal volume of Field’s
stain A solution and mix well by tilting the slide.
2. Wash off stain with clean water
3. Place slide up right in a draining rack to air dry.
71

72. 72