This document provides information on preparing blood smears with different staining methods. It discusses preparing thin and thick blood smears using various techniques like the slide method, cover glass method, and spin method for thin smears. For thick smears, a large drop of blood is spread on the slide. Various staining methods are described like Leishman stain, Giemsa stain, Wright stain, and Field stain. Proper preparation, fixation in methanol or ethanol, and staining are required to visualize blood cells and parasites under the microscope. Reticulocyte staining is also briefly mentioned.

1. Preparation of blood smear
with different
staining method
Dr. Ankur Patel
Ankurvety001@gmail.com

2. Blood film ????
• A blood film or peripheral blood smear is a thin layer
of blood smeared on a microscope slide and then stained in such a
way to allow the various blood cells to be examined
microscopically.
Example of properly prepared thick and thin film blood smears

3. Aim of blood smear
• Blood films are usually examined to
investigate hematological problems (disorders of the blood)
and, occasionally, to look for parasites within the blood such
as malaria and filaria.
• Examination of thin blood films is important in the
investigation and management of anaemia, infections, and
other conditions which produce changes in the appearance
of blood cells and differential white cell count.
• A blood film report can provide rapidly and at low cost,
useful information about a patient’s condition.

4. The peripheral blood film (PBF) is of two types:
1. Thin blood film
2. Thick blood film
Three basic steps to make blood film:
1. Preparation of blood smear.
2. Fixation of blood smear.
3. Staining of blood smear.

5. Preparation of blood smear

6. 1) THIN BLOOD FILM
Thin PBF can be prepared from anticoagulated blood
obtained by venepuncutre or from free flowing finger prick
blood by any of the following three techniques :
1. Slide method
2. Cover glass method
3. Spin method

7. Slide Method
Procedure
 Place a drop of blood in the centre of a clean glass
slide 1 to 2 cm from one end.
 Place another slide (spreader) with smooth edge at
an angle of 30-45⁰ near the drop of blood.
 Move the spreader backward so that it makes contact with
drop of blood.
 Then move the spreader forward rapidly over the slide.
 A thin peripheral blood film is thus prepared
 Dry it and stain it.

8. Cover Glass Method
Procedure
 Take a clean cover glass.
 Touch it on to the drop of a blood.
 Place it on another similar cover glass in crosswise direction
with side containing drop of blood facing down.
 Pull the cover glass quickly.
 Dry it and stain it.
 Mount it with a mountant, film side down on a clean glass
slide.

9. Spin Method
This is an automated method.
Procedure
 Place a drop of blood in the centre of a glass slide.
 Spin at a high speed in a special centrifuge, cytospin.
 Blood spreads uniformly.
 Dry it and stain it.

10. THICK BLOOD FILM
This is prepared for detecting blood parasites such as
malaria and microfilaria.
Procedure:
 Place a large drop of blood in the centre of a clean glass slide.
 Spread it in a circular area of 1.5 cm with the help of a
stick or end of another glass slide.
 Dry it
 Staing

12. The shape of blood film

13. Qualities of a Good Blood Film
i. It should not cover the entire surface of slide.
ii. It should have smooth and even appearance.
iii. It should be free from waves and holes.
iv.It should not have irregular tail.

14. Thethicknessofthespreadwhenpullingthesmear
isdeterminedby :
1. The angle of the spreader slide. (the greater the angle,
the thicker and shorter the smear).
2. Size of the blood drop.
3. Speed of spreading.
Notes:
1. If the haematocrit increased, the angel of the spreader
slide should be decreased.
2. If the haematocrit decreased, the angel of the
spreader slide should be increased.

15. large angle
low HCT
small angle
high HCT

16. Parts of a Thin Blood Film
A peripheral blood film consists of 3 parts :
1. Head i.e. the portion of blood film near the drop of blood.
2. Body i.e. the main part of the blood film.
3. Tail i.e. the tapering end of the blood film.

17. Commoncauseof a poorblood smear:
1. Drop of blood too large or too small
2. Spreader slide pushed across the slide in a jerky manner
3. Failure in keep the entire edge of the spreader slide against
the slide while making the smear
4. Failure in keep the spreader slide at a 30⁰ angel with the
slide
5. Failure to push the spreader slide completely across the
slide
6. Irregular spread with ridges and long tail: edges of spreader
dirty or chipped ; dusty slide
7. Holes in film – slide contaminated with fat or grease and air
bubbles
8. Cellular degenerative changes: delay in fixing inadequate
fixing time or methanol contaminated with water

18. A. Blood film with jagged tail made from a spreader with
achipped end.
B. Film which is too thick
C. Film which is too long, too wide, uneven thickness and
made on a greasy slide.
D. A well-made blood film.

19. Examples of unacceptablesmears

20. Biologic causes of a poor smear :
1. Cold agglutinin - RBCs will clump together.
Warm the blood at 37° C for 5 minutes, and then
remake the smear.
2. Lipemia - holes will appear in the smear.
There is nothing you can do to correct this.
3. Rouleaux - RBC’s will form into stacks resembling coins.
There is nothing you can do to correct this.

21. Fixation of blood smear

22. • To preserve the morphology of the cells, films must be
fixed as soon as possible after they have dried.
• It is important to prevent contact with water before
fixation is complete.
• Methyl alcohol (methanol) is the choice, although ethyl
alcohol ("absolute alcohol") can be used.
• Methylated spirit (95% ethanol) must not be used as it
contains water.
• To fix the films, place them in a covered staining jar or
tray containing the alcohol for 2-3 minutes. In humid
climates it might be necessary to replace the methanol 2-
3 times per day; the old portions can be used for storing
clean slides.

23. Different fixatives

24. Staining of blood smear

25. Variousstainsforperipheralbloodfilm:
 Romanowsky stains are universally employed for staining
of blood films. All Romanowsky combinations have two
essential ingredients i.e. methylene blue and eosin or
azure.
• Methylene blue is the basic dye and has affinity for acidic
component of the cell (i.e. nucleus) and eosin/azure is
the acidic dye and has affinity for basic component of cell
(i.e. cytoplasm).
• Most Romanowsky stains are prepared in methyl alcohol
so that they combine fixation and staining.

26. VariousstainsincludedunderRomanowskystainare
asunder:
1. Leishman stain
2. Giemsa stain
3. Wright stain
4. Field stain
5. Jenner stain
6. JSB stain

27. Leishman Stain:
Preparation
• Dissolve 0.2 g of powdered Leishman’s dye in 100 ml of
acetone-free methyl alcohol in a conical flask.
• Warm it to 50°C for half an hour with occasional shaking.
• Cool it and filter it.
Procedure for staining
• Pour Leishman’s stain dropwise (counting the drops) on the
slide and wait for 2 minutes. This allows fixation of the PBF in
methyl alcohol.
• Add double the quantity of buffered water dropwise over
the slide (i.e. double the number of drops).
• Mix by rocking for 8 minutes.
• Wash in water for 1 to 2 minutes.
• Dry in air and examine under oil immersion lens of the
microscope.

29. Metarubricyte (top left) and polychromatophilic erythrocyte (arrow) in
regenerative anemia. A monocyte (top right) also is present (Dog,
blood, Wright-Leishman stain).

30. Giemsa Stain
Preparation
• Stock solution of Giemsa stain is prepared by mixing 0.15 g of
Giemsa powder in 12.5 ml of glycerine and 12.5 ml of methyl
alcohol.
• Before use dissolve one volume of stock solution in nine volumes
of buffered water (dilution 1:9).

31. Procedure:
• Fill staining dish with staining solution
• Place thin film and thick films into the staining dish.
• Stain blood slides for 45 minutes
• Wash in water.
• Dry it and examine under oil immersion lens of the
microscope.

32. Caution:
Thick films need careful
rinsing!
(since they are not fixed
before staining)
rinsed
Mistake:
Thick films are not
properly! Blood is lost!

36. Trypanosoma org. In blood

38. Wright'sstain:
• Wright’s stain is used to differentiate nuclear and/or cytoplasmic
morphology of platelets, RBCs, WBCs, and parasites.
Procedure
• Thin blood films (only) – Dip Method
1. Dip air dried blood film in undiluted stain for 15 to 30 seconds
(double the staining time for bone marrow smears).
2. Decolorize the stained smears by immersion in distilled or deionized
water and air dry
3. Let air dry in a vertical position.
• Thin blood films (only) – Rack Method
1. Lay air dried slides on staining rack and flood with stain; stain for 10 to
15 seconds(double the staining time for bone marrow smears).
2.Add an equal volume of deionized/distilled water and stain for 10
seconds.
3.Rinse the slide by dipping in deionized/distilled water for 30 seconds.The
slide may also be rinsed by swishing or washing with deionized/distilled
water.

39. Thickbloodfilms(only):
1. Allow film to air dry thoroughly for several hours or overnight. Do
not dry films in an incubator or by heat, because this will fix the
blood and interfere with the lysing of the RBCs.
Note: If a rapid diagnosis of malaria is needed, thick films can
be made slightly thinner than usual, allowed to dry for 1 h, and
then stained.
2. Lake the thick film by immersing in distilled or deionized water for
10 min.
3. Allow the film to air dry thoroughly.
4. Fix air-dried film in absolute methanol for 30 seconds in a Coplin
jar containing absolute methanol.
5. Allow the film to air dry.
6. Dip air dried blood film in undiluted stain for 15 to 30 seconds
(double the staining time for bone marrow smears).
7. Decolorize the stained smears by immersion in distilled or
deionized water and air dry
8. Let air dry in a vertical position.

40. VenousbloodfromPekinduck(Wright'sstain)

41. Trypanosoma brucei_thin_wright-giemsa stain

42. Arrow indicatemalarial parasites

43. Fieldstain: (thin film)
Materials:
• Methanol (absolute)
• Field’s stain A und B
• Tube with water
• Staining dishes
• Filter paper
Procedure:
A. Fix thin film with methanol
for 1 min.

44. B. Dry microscopic slide on filter paper
C. Immerse slide in Field’s stain B (Eosin) for 5 seconds
D. Immediately wash with water
E. Immerse slide in Field‘s stain A (Methylene blue) for 10 sec
F.Immediately wash with water
G. Dry thin films

45. Materials:
• Methanol No!
• Field‘s stain A und B
• Tube with water
• Filter paper
Procedure:
A. Immerse thick film in Field‘s stain A (Methylene blue) for 3 sec
Do not forget:
• Thick films need to be
• haemolysed and are
• therefore not fixed with
• methanol
Fieldstain: (thick film)

46. B. Rinse immediately in tap water
C. Immerse thick film in Field’s stain B (Eosin) for 3 seconds
D. Then rinse immediately with tap water
E. Let the slide carefully dry

47. Jenner stain:
• The Jenner stain Solution is a mixture of several thiazin
dyes in a methanol solvent
• Ionic and noionic forces are involved in the binding of
these dyes
• The staining solution has anionic and cationic properties
• The negatively charged phosphoric acid groups of DNA
attract the purple polychromatic cationic dyes to the
nuclei
• The blue basophilic granules are stained by the
polychromatic cationic dyes

48. ImmersionStainingProtocol:
• Thoroughly dry blood or bone marrow smears
• Fix smears in absolute methanol for 15 seconds to 5 minutes
• Stain smears in Jenners Stain Solution for 2 minutes
• Stain in mixture of 50ml of Jenners Stain Solution, 75ml of
PH 6.6 Phosphate Buffer Solution and 175ml deionized
water for 5 minutes
• Rinse in standing deionized water for 1.5 minutes or rinse
briefly in running deionized water
• Air dry smears
• Examine smears under a microscope

49. HorizontalStainingProtocol:
• Place slide with thoroughly dried film in a horizontal
staining rack
• Flood smear with absolute methanol for 15-30 seconds and
then drain
• Flood smear with 1ml Jenners Stain Solution and let stand
for 3 minutes
• Add 1mL of pH 6.6 Phosphate Buffer solution and 1 ml
deionized water to smear and let stand for 45 seconds
• Rinse briefly with running deionized water
• Air dry and examine under a microscope
• Perform immuno chemical staining procedure according to
manufacturer.

50. J.S.B.Stain:
Materials and reagents required:
• Eosin yellow (water soluble)
• Methylene blue
• Potassium dichromate
• Di-sodium hydrogen phosphate (dihydrate)
• 1% sulphuric Acid.
• Round bottom flask (2 lit.)
• Healing mantle
• Distilled water
• Staining jars.

51. Staining technique:
• Prepare thin and thick smears from malaria cases on micro slides
• De-haemoglobinise the thick smear
• Fix the thin smear in methanol for few minutes
• Take 3 staining jars for J.S.B. I, J.S.B.II and tap water
• Dip the smears in J.S.B. II for few seconds and immediatedly wash
in water
• Drain the slides free of excess water
• Dip the smears in J.S.B.I for 30-40 seconds
• Wash well in water and dry
• Examine the smears under oil immersion

52. Staining of Thick Smear:
• It can be stained with any of the Romanowsky stains
• listed above except that before staining, the smear is
dehaemoglobinised by putting it in distilled water for 10
minute
Autostainers
• Currently, automatic staining machines are available
which enable a large batch of slides to be stained with a
uniform quality.

53. Jaswant Singh Battacharya (JSB) Stain for thick and thin films:
• This is the standard method used by the laboratories under
the National Malaria Eradication Programme in India

54. PrecautionsinStainingofperipheralbloodfilm
1) Dark blue blood film:
It can be due to overstaining, inadequate washing or
improper pH of the buffer. In this RBCs are blue, nuclear chromatin
is black,granules of the neutrophils are overstained and granules
of the eosinophils are blue or grey.
2) Light pink blood film:
In this RBCs are bright red, the nuclear chromatin is pale
blue and granules of the eosinophils are dark red. It can be due to
understaining, prolonged washing, mounting the film before
drying and improper pH of the buffer.
3) Precipitate on the blood film:
This could be due to inadequate filtration of the stain, dust
on the slide, drying during staining and inadequate washing.

55. TO ACIDIC SUITABLE TO BASIC

57. RETICULOCYTE COUNT
Prabin Shah BScMLT, MSc(Biochemistry)

58. RETICULOCYTES
🟔Reticulocytes are young , premature, non
nucleated red blood cells, contain reticular material
(RNA) that stain gray blue.
🟔Reticulum is present in newly released blood cells
for 1-2 days before the cell reach its full mature
state.

59. RETICULOCYTE STAINS
🟔Reticulocytes are visualized by supra-vital staining (such
as new methylene blue, Brilliant Cresyl Blue, Pure azure
blue) that precipitate the RNA and organelles, forming a
filamentous network of reticulum
🟔On Wright stain. the Reticulocyte appears
polychromatophilic or as a Macrocytic blue red cell.

60. PRINCIPLE
🟔 Whole blood is incubated with supra-vital
staining (new methylene blue). The vital stain
causes the ribosomal and residual RNA to
coprecipitate with the few remaining mitochondria
and ferritin masses in living young erythrocytes to
form dark-blue clusters and filaments (reticulum).
🟔 Smears of this mixture are then prepared and
examined. The number of reticulocytes in 1000
red blood cells is determined. This number is
divided by 10 to obtain the reticulocyte count in
percent.

61. SPECIMEN
 Whole blood that is anticoagulated with either EDTA
or heparin is suitable.
 Capillary blood drawn into heparinized tubes or
immediately mixed with stain may also be used.
 Red blood cells must still be living when the test is
performed therefore it is best to perform it promptly
after blood collection.
 Blood may be used up to 8 hours after collection.
 Stained smears retain their color for a prolonged
period of time.

62. REQUIREMENTS
1. Commercially prepared liquid new methylene blue
solution. It should be stored in a brown bottle. If
precipitate is a problem on the smear, the stain
should be filtered prior to use.
2. Microscope slides
3. Microscope
4. 10 x 75 mm test tubes
5. Pasteur pipets (with bulb if pipets are glass)
6. Capillary tubes
7. Miller ocular (if available)

63. PROCEDURE
Preparation of smears
1. Add 3-4 drops of new methylene blue solution to 3-4
drops of thoroughly mixed EDTA anticoagulated blood to
a glass 10 x 75 mm test tube.
2. Mix the contents by gently shaking and allow to incubate
at room temperature for a minimum of 10 minutes.
3. At the end of 10 minutes, gently mix the blood/stain
solution.
4. Using a capillary tube, place a drop of the mixture on
each of three slides near the frosted edge as you would
when making a peripheral smear.
5. Using the wedge smear technique, make acceptable
smears not too thick or thin.
6. Label the slides with patient name, ID# and date.
7. Allow to air dry. (Do not blow to hasten to drying.)

64. COUNTING PROCEDURE
 Place the first slide on the microscope stage and,
using the low power objective (10x), find an area in
the thin portion of the smear in which the red cells are
evenly distributed and are not touching each other.
 Carefully change to the oil immersion objective (100x)
and further located an area in which there are
approximately 100 red cells per oil immersion field.
 Do this by finding a field where the cells are evenly
distributed and mentally divide the field into 4
quadrants. Count the cells in 1 quadrant. If there are
about 25, you are in a good area. There will be a lot of
open space between the red cells.

65. 2. Be sure to count all cells that contain a blue-staining
filament or at least 2 or more discrete blue aggregates
of reticulum in the erythrocyte.
3. Count 1000 red cells in consecutive oil immersion
fields. Record the number reticulocytes seen.
4. You may count 500 cells on two slides each. They
should agree within ± 15% of each other. If they do
not, repeat the reticulocyte count on the third smear.
5. Calculate the result as follow:
%Retix= Reticulocyte counted/10

66. MILLER DISCS METHOD
1. Use a 100x objective and a 10x ocular secured with a Miller
disc.
 The Miller disc imposes two squares (one 9 times the area of the
other) onto the field of view.
 Find a suitable area of the smear. A good area will show 3-10
RBCs in the smaller square of the Miller disc.
2. Count the reticulocytes within the entire large square
including those that are touching the lines on the left and
bottom of the ruled area. Count RBCs in the smaller square
whether they contain stained RNA or not. A retic in the
smaller square should be counted as an RBC and a retic.
Record RBC # counted and retic # counted separately.
3. Continue counting until a minimum of 111 RBCs have been
observed (usually 15-20 fields). This would correspond to
999 RBCs counted with the standard procedure.

67. The Miller disc may be placed in one of the ocular
lenses to aid in the counting of the reticulocytes.
%Reticulocyte= total reticulocyte in square A*100
total RBC in square B*9

68. REFERENCE VALUES
 RBCs life span ~ 100 days, ± 20 days
 Reticulocyte ~ 1 day in peripheral blood, Then
the B.M. replaces approximately 1 % of the adult
red blood cells every day.
 Normal value :
 0.5 to 1.5/100 red blood cells (or
, 0.5 to
1.5%)
 Absolute count :25 to 75 X 109/L
 Newborn (0-2 weeks):2.5-6.0%
 Normal Reticulocyte I
ndex :1-3%

69. RETICULOCYTE

70. REPORTING PATTERN
🟔Absolute Reticulocyte Count (ARC): is the actual
number of reticulocytes in 1L of whole blood. This is
calculated by multiplying the reticulocytes % by the
RBCs count and dividing by 100.
🟔Corrected Reticulocyte Count is calculated based
on a normal hematocrit of 45%.
🟔 Reticulocyte Production Index (RPI) = Corrected retic
count (%) / # Days (Maturation time)

71. CORRECTED RETICULOCYTE COUNT
 In states of anemia, the reticulocyte percentage is
not a true reflection of reticulocyte production. A
correction factor must be used so as not to
overestimate marrow production, because each
reticulocyte is released into whole blood containing
few RBCs - a low hematocrit (Hct) - thus relatively
increasing the percentage.
 The corrected reticulocyte count my be calculated
by the following formula:

72. RETICULOCYTE PRODUCTION INDEX(RPI)
 The RI is a measurement for reticulocytes when anemia is
present
 Estimating RBC production by using the corrected
reticulocyte count may yield erroneously high values in
patients when there is a premature release of younger
reticulocytes from the marrow (owing to increased
erythropoietin stimulation).
 The premature reticulocytes are called “stress or shift”
reticulocytes. These result when the reticulocytes of the
bone marrow pool are shifted to the circulation pool to
compensate for anemia.
 The younger stress reticulocytes present with more
filamentous reticulum. The mature reticulocyte may present
with granular dots representing reticulum. Normally,
reticulocytes lose their reticulum within 24 to 27 hours after
entering the peripheral circulation.

73.  The premature stress retics have increased reticulum
and require 2 to 2.5 days to lose their reticulum,
resulting in a longer peripheral blood maturation time.
 The peripheral blood smear should be reviewed
carefully for the presence of many polychromatophilic
macrocytes, thus indicating stress reticulocytes and
the need for correction for both the RBC count and the
presence of stress reticulocytes. The value obtained is
called the reticulocyte production index (RPI).

74. Maturation Time Hematocrit %
1 day 45
1.5 day 35
2 day 24
3 day 15

75. INTERPRETATION
 The Reticulocyte count is an important diagnostic
tool: The number of Reticulocytes is a good indicator of
bone marrow activity, because it represents recent
production. It is used to differentiate anemia's caused by
bone marrow failure from those caused by hemorrhage
or hemolysis.
 It used also to check the effectiveness of treatment in
prenicious anemia and folate and iron deficiency.
 To assess the recovery of bone marrow function in
aplastic anemia and to determine the effects of
radioactive substance on exposed workers.
 A low reticulocyte count may mean a need for a bone
marrow biopsy. This can tell if is a problem with how new
reticulocytes are made by the bone marrow.

76.  Reticulocytosis (Increased RBC Production)
⚫ Reticulocyte Index >3%, Reticulocyte Count >1.5%
1. Acute blood loss or hemorrhage
2. Post-Splenectomy
3. Acute Hemolytic Anemia (Microangiopathic
Anemia)
4. Hemoglobinopathy
 Sickle Cell Anemia
 Thalassemia major
5. Post-Anemia Treatment
 Folate Supplementation
 Iron Supplementation
 Vitamin B12 Supplementation

77.  Reticulocytopenia (Decreased RBC Production)
⚫ Reticulocyte Index <1%, Reticulocyte Count <0.5%
1. Aplastic Anemia
2. Bone Marrow infiltrate
3. Bone Marrow suppression or failure
1. Sepsis
2. Chemotherapy or radiotherapy
4. Disordered RBC maturation
1. Iron Deficiency Anemia
2. Vitamin B12 Deficiency
3. Folate Deficiency
4. Sideroblastic Anemia
5. Anemia of Chronic Disease
6. Hypothyroidism
5. Blood transfusion
6. Liver disease

78. FACTORS AFFECTING THE TEST
Reasons you may not be able to have the
test or why the results may not be helpful
include:
⚫ Taking medicines, such as levodopa,
corticotropin, azathioprine (Imuran),
chloramphenicol (Chloromycetin), dactinomycin
(Cosmegen), medicines to reduce a fever,
medicines to treat malaria, and methotrexate and
other cancer chemotherapy medicines.
⚫ Getting radiation therapy
⚫ Taking sulfonamide antibiotics (such as Bactrim or
Septra)
⚫ Being pregnant
⚫ Having a recent blood transfusion

79. ERROR SOURCES
1. A refractile appearance of erythrocytes should not be
confused with reticulocytes.
2. Filtration of the stain is necessary when precipitated
material is present which can resemble a reticulocyte.
3. Erythrocyte inclusions should not be mistaken for
Reticulocytes.
 Howell-Jolly bodies appear as one or sometime two, deep-purple
dense structures.
 Heinz bodies stain a light blue-green and are usually present at the
edge of the erythrocyte.
 Pappenheimer bodies are more often confused with reticulocytes
and are the most difficult to distinguish. These purple-staining iron
deposits generally appear as several granules in a small cluster. If
Pappenheimer bodies are suspected, stain with Wright-Giemsa to
verify their presence. Hemoglobin H inclusions will appear as multiple
small dots in every cell.

80. 4. Falsely decreased reticulocyte counts can result
from under staining the blood with new methylene
blue. Be sure the stain/blood mixture incubates the
full 10 minutes.
5. High glucose levels can cause reticulocytes to stain
poorly.
6. There is high degree of inaccuracy in the manual
reticulocyte count owing to error (± 2%) in low
counts and ± 7% in high counts) and a lack of
reproducibility because of the inaccuracy of the
blood film. This inaccuracy has been overcome by
the use of automated instruments using flow
cytometry.
7. If no reticulocytes are observed after scanning at
least two slides, report “none seen”.