Bleeding disorders are the result of a generalized defect in hemostasis due to abnormalities of blood vessels, platelets, or coagulation factors.
Initial tests, which should be performed in a suspected bleeding disorder, are complete blood count including blood smear, platelet count, bleeding time, clotting time, prothrombin time, and activated partial thromboplastin time. Depending on the results of these screening tests, one or more specific tests are carried out for definitive diagnosis (e.g. platelet function studies, assays of
coagulation factors, and test for fibrin degradation products). Abnormalities of blood vessels are usually not detectable by laboratory tests for hemostasis, and their diagnosis requires correlation of clinical and other investigations.
A complete blood count and a blood smear can provide information in the form of:
• Presence of cytopenia (anemia, leukopenia, thrombocytopenia)
• Red cell abnormalities (especially fragmented red cells which may indicate disseminated intravascular
• White cell abnormalities (like abnormal cells in leukemias)
• Abnormalities of platelets: thrombocytopenia (normally there is 1 platelets per 500-1000 red cells), giant platelets (seen in myeloproliferative disorders and Bernard-Soulier syndrome), and isolated discrete platelets without clumping in finger-prick smear (seen in uremia, Glanzmann’s thrombasthenia).
(2) Platelet Count
(3) Bleeding Time (BT)
(4) Clotting Time (CT)
(5) Prothrombin Time (PT)
(6) Activated Partial Thromboplastin Time (APTT)
(7) Thrombin Time (TT)
(8) Platelet Function Analyzer-100
|LICENSE: This article uses material from the Wikipedia article "HEMOSTASIS", which is released under the Creative Commons Attribution-Share-Alike License 3.0.|
New methylene blue: 1.0 gm
Sodium citrate: 0.6 gm
Sodium chloride: 0.7 gm
Distilled water: 100 ml
Suitable alternatives to new methylene blue are brilliant cresyl blue and azure B.
(2) Add equal amount of blood and mix well.
(3) Keep the mixture at room temperature or at 37°C for 15 minutes.
(4) After gentle mixing, place a small drop from the mixture on a glass slide, prepare a thin smear, and allow to dry in the air.
(5) Examine under the microscope using oil-immersion objective. Mature red cells stain pale green blue. Reticulocytes show deep blue precipitates of fine granules and filaments in the form of a network (reticulum). Most immature reticulocytes show a large amount of precipitated material in the form of a mass, while the most mature reticulocytes show only a few granules or strands. Any nonnucleated red cell is considered as a reticulocyte if it contains 2 or more blue-stained particles of ribosomal RNA.
(6) Count 1000 red cells and note the number of red cells that are reticulocytes. Counting error is minimized if size of the microscopic field is reduced. This is achieved by using a Miller ocular disk inserted in the eyepiece; it divides the field into two squares (one nine times larger in size than the other). Reticulocytes are counted in both the squares and the red cells are counted in the smaller square.
Normal: 50,000 to 85,000/cmm
Estimated maturation time in days
• Absolute reticulocyte count: 50,000-85,000/cmm
2. The Expert Panel on Cytometry of the International Council for Standardization in Hematology: ICSH guidelines for reticulocyte counting by microscopy on supravitally stained preparations. World Health Organization. WHO/LBS/92.3 1992.
- As one of the baseline studies in anemia with no obvious cause
- To diagnose anemia due to ineffective erythropoiesis (premature destruction of red cell precursors in bone marrow seen in megaloblastic anemia and thalassemia) or due to decreased production of red cells: In hypoplastic anemia or in ineffective erythropoiesis, reticulocyte count is low as compared to the degree of anemia. Increased erythropoiesis (e.g. in hemolytic anemia, blood loss, or specific treatment of nutritional anemia) is associated with increased reticulocyte count. Thus reticulocyte count is used to differentiate hypoproliferative anemia from hyperproliferative anemia.
- To assess response to specific therapy in iron deficiency and megaloblastic anemias.
- To assess response to erythropoietin therapy in anemia of chronic renal failure.
- To follow the course of bone marrow transplantation for engraftment
- To assess recovery from myelosuppressive therapy
- To assess anemia in neonate
Reticulocyte Count Test
Reticulocyte Values In Sickle Cell Anemia
Reticulocyte Count Reference Range
Supravital Stain Reticulocytes
Reticulocyte Count Method
Reticulocyte Staining Procedure
Reticulocytes With Supravital Stain
Miller Disc Retic Count
Reticulocyte Count Procedure Miller Disc
(2) A 'spreader' slide is placed at an angle of 30° in front of the drop and then drawn back to touch the drop of blood. Blood spreads across the line of contact of two slides.
(3) Smear is made by smooth, forward movement of the 'spreader' along the slide. The whole drop should be used up 1 cm before the end of the slide. The length of the smear should be about 3 cm. The 'spreader' should not be raised above the slide surface till whole drop of blood is spread out.
(5) Patient's name or laboratory number and date are written (with a lead pencil, a permanent marker pen, or a diamond pencil) on the thicker end of the smear.
(6) The smear is fixed immediately with absolute methyl alcohol (which should be moisture- and acetone-free) for 2-3 minutes in a covered jar (Absolute ethyl alcohol can also be used, but not methylated spirit as it contains water). Aim of fixation is to prevent washing off of the smear from the slide. Following this, color of the smear becomes light brown. This fixation is desirable even when Leishman stain is used which contains methyl alcohol. This is because Leishman stain may have absorbed moisture leading to poor fixation. If methanol is contaminated with water, sharpness of cell morphology is lost and there is vacuolation of red cells. Methanol should be acetone-free since acetone washes out nuclear stain. (In many laboratories, slide is stained immediately after air-drying without prior fixation, and the results are satisfactory; however, if delay of >4 hours is anticipated between air-drying and staining, the slide should be fixed. If not, a background gray-blue staining of plasma occurs).
(1) Making a 'spreader' slide—a glass slide with absolutely smooth edges should be selected and one or both corners at one end of the slide should be broken off. The 'spreader' slide should be narrower (width of about 15 mm) so that edges of the smear can be examined microscopically. The 'spreader' slide should be discarded after use. If the same is to be reused, its edge should be thoroughly cleaned and dried (otherwise carryover of cells or parasites can occur).
(2) A well-spread blood smear (a) is tongue-shaped with a smooth tail, (b) does not cover the entire area of the slide, (c) has both thick and thin areas with gradual transition, and (d) does not contain any lines or holes.
(3) By changing the angle of the 'spreader' and its speed, thickness of the blood smear can be controlled. In patients with anemia, a thicker smear can be obtained by increasing the angle and the speed of spreading. In patients with polycythemia, a thinner smear is obtained by decreasing the 'spreader' angle and the speed of spreading.
(5) It is recommended to stain blood films in reagent filled Coplin jars (rather than covering them with the staining solution) to avoid formation of stain precipitates due to evaporation.
(6) A drawback of this method is uneven distribution of leukocytes (i.e. monocytes, neutrophils, and abnormal cells are pushed towards the extreme tail end of the smear) and distortion of red cell morphology at the edges.
(7) Blood smear is covered with a coverslip and mounted in a mounting medium (e.g. DPX) for protection against mechanical damage and deterioration of staining with time on exposure to air.
STAINING OF BLOOD SMEAR
Blood smears are routinely stained by one of the Romanowsky stains. Romanowsky stains consist of a combination of acidic and basic dyes and after staining various intermediate shades are obtained between the two polar (red and blue) stains. Romanowsky stains include May-Grunwald-Giemsa, Jenner, Wright's, Leishman's, and Field's stains. Staining properties of the Romanowsky stains are dependent on two synthetic dyes: methylene blue and eosin. International Committee for Standardization in Haematology has recommended a highly purified standardized stain, which contains azure B and eosin Y; it, however, is very expensive. Romanowsky stains are insoluble in water but soluble in methyl alcohol. Methyl alcohol acts as a solvent as well as a fixative. Staining reaction is pH-dependent. These stains have a tendency towards precipitation and should be filtered before use.
Methylene blue and azure B are basic (cationic) dyes and have affinity for acidic components of the cells (like nucleic acids or basophil granules) and impart purpleviolet color to the nuclear chromatin, dark blue-violet color to the basophil granules, and deep blue color to the cytoplasm of lymphocytes. Eosin is an acidic (anionic) dye and has affinity for basic components like hemoglobin (stained pink-red), and granules in eosinophils (stained orange-red). Neutrophil granules are slightly basic and stain violet-pink or lilac.
Romanowsky stains impart more colours than just blue (from methylene blue or azure B) and red-orange (from eosin Y). Usefulness of the Romanowsky stains lies in their ability to differentially stain leucocyte granules.
Method of Leishman staining is given below:
(1) Leishman stain: William Boog Leishman, a British pathologist, modified the original Romanowsky method and devised a stain which is widely known as Leishman's stain. This consists of methylene blue and eosin dissolved in absolute methyl alcohol. Commercially available Leishman stain powder (0.6 gram) is mixed with water-free absolute methyl alcohol (400 ml). Prepared stain should be kept tightly stoppered in a brown bottle and stored in a cool, dark place at room temperature. Exposure to direct sunlight causes deterioration of the stain. After preparation, stain should be kept for 3-5 days before using since it improves the quality of the stain.
(2) Buffered water (pH 6.8).
(1) Air-dry the smear and fix with methanol for 2-3 minutes.
(2) Cover the smear with Leishman stain for 2 minutes.
(3) After 2 minutes, add twice the volume of buffered water and leave for 5-7 minutes. A scum of metallic sheen forms on the surface.
(4) Wash the stain away in a stream of buffered water. Tap water can also be used for washing if it is not highly alkaline or highly acid.
(5) Wipe the back of the slide clean and set it upright in the draining rack to dry.
(6) Mount the slide in a suitable mounting medium (e.g. DPX) with a clean and dry 25 × 25 mm coverslip.
• Red cells: pink-red or deep pink
• Polychromatic cells (Reticulocyt-es): Gray-blue
• Neutrophils: Pale pink cytopla-sm; mauve-purple granules
• Eosinophils: Pale-pink cytoplasm; orange-red granules
• Basophils: Blue cytoplasm; dark blue-violet granules
• Monocytes: Gray-blue cytoplasm; fine reddish (azurophil) granules
• Small lymphocytes: Dark blue cy-toplasm
• Platelets: Purple
• Nuclei of all cells: Purple-violet
- Stage 1: Lag pahse or rouleaux formation: The RBCs stack together and form a structure like package of coins in a shape of canned. (10 Minutes)
- Stage 2: Submerged of rouleaux. (40 Minutes)
- Stage 3: Slow sedimentation. (10 Minutes)
COMPONENTS INFLUENCING ON ERYTHROCYTE SEDIMENTATION RATE
- Red Blood Cells: In polycythemia, the mass of the red blood cells increases which cause the decrease of ESR. In Anemia the mass of the red cells decreases which cause the increase of ESR. In other words, Erythrocyte Sedimentation Rate is indirectly proportional to ratio between mass of the red cells and plasma.
- Plasma: The most important component influencing on Erythrocyte Sedimentation Rate is the composition of plasma. High level of C-Reactive Protein, fibrinogen, heptoglobin, α1-antitrypsin, ceruloplasmin and immunoglobulins causes the elevation of Erythrocyte Sedimentation Rate. When the level of proteins increases in plasma, it reduce the negative charge from the surface of red cells and depreciate the zeta potential; this facilitate the attraction between red blood cells, and form rouleaux.
- Technical Issues: Elevation in room temperature also affects the Erythrocyte Sedimentation Rate. Moving tube in sloping position, length and calibre of the tube also affect Erythrocyte Sedimentation Rate.
- Wintrobe method
- Westergren method
- Zeta Sedimentation Ratio
- Trisodium citrate dihydrate 32.08 gm
- Distilled water upto 1000 ml
- Mix the anticoagulated blood smaple thoroughly. Fill the Westergren tube with blood upto “ZERO” mark. Note that there is no air bubbles in the blood.
- Place the tube is vertical position in the ESR stand and left for an hour.
- Just exactly after an hour, read the height of the column of plasma above the red cells column in mm.
- Result is express in the following manner:
Erythrocyte Sedimentation Rate = ________ mm in an hour.
- Always use the correct ratio of anticoagulant and blood. Blood should be check for clots and air bubbles before going to the further process. Blood and anticoagulant should be mix thoroughly.
- Make sure that the temperature of the room is between 18-25°C. If the room temperature is elevated than 25°C, Erythrocyte Sedimentation Rate will increase and different reference range will acquire.
- ESR tube must be in strict vertical position. Even a slight tilting will cause elevation in Erythrocyte Sedimentation Rate.
- Male: 0-9 mm in an hour
- Females: 0-20 mm in an hour
- Children: 0-13 mm in an hour
Erythrocyte Sedimentation Rate by Westergren Method
- Males < 50 years: 0-15 mm in an hour.
- Females < 50 years: 0-20 mm in an hour.
- Children: 0-10 mm in an hour.
- Elderly males > 50 years: 0-20 mm in an hour.
- Elderly females > 50 years: 0-30 mm in an hour
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• Manual or microscopic method
• Automated method
Free-flowing capillary or well-mixed anticoagulated venous blood is added to a diluent at a specific volume in the Unopette reservoir. The diluents (1% ammonium oxalate) lyses the erythrocytes but preserves leukocytes and platelets. A 20 µL pipette is used with 1.98 ml of diluents to make a 1:100 dilution. The diluted blood is added to the hemacytometer chamber. Cells are allowed to settle for 10 minutes before leukocytes and platelets are counted. (Always refer to the manufacturer’s instructions for the procedure.)
Hemocytometer with cover glass, compound microscope. Unopette capillary pipette, lint-free wipe, alcohol pads, hand counter, petri dish with moist filter paper.
Ammonium oxalate: 11.45 gm
Sorensen’s phosphate buffer: 1.0 gm
Thimerosal: 0.1 gm
Distilled water: 1000 ml
EDTA-anticoagulated blood or capillary blood is preferred.
(1) Using the protective shield on the capillary pipette, puncture diaphragm of Unopette reservoir.
(2) Remove shield from pipette assembly by twisting. Holding pipette almost horizontally, touch tip of pipette to blood. Pipette will fill by capillary action. Filling will cease automatically when the blood reaches the end of the capillary bore in the neck of the pipette.
(4) Squeeze reservoir slightly to force out some air while simultaneously maintaining pressure on reservoir.
(5) Cover opening of overflow chamber of pipette with index finger and seat pipet securely in reservoir neck.
(6) Release pressure on reservoir. Then remove finger from pipette opening. At this time negative pressure will draw blood into reservoir.
(7) Squeeze reservoir gently two or three times to rinse capillary bore forcing diluent up int, but not out of, overflow chamber, releasing pressure each time to return mixture to reservoir.
(8) Place index finger over upper opening and gently invert several times to thoroughly mix blood with diuent.
(9) Cover overflow chamber with pipette shield and incubate at room temperature for 10 minutes before charging the hemacytometer.
(10) Meticulously clean the hemacytometer with alcohol or other cleaning solution. This is important because dust particles and other debris can be mistaken for platelets especially on a light microscope. Allow to dry completely before charging with diluted specimen.
(11) To charge the hemacyto-meter, convert to dropper assembly by withdrawing pipette from reservoir and reseating securely in reverse position.
(12) Invert reservoir and discard the first 3 or 4 drops of mixture.
(13) Carefully charge hemacyto-meter with diluted blood by gently squeezing sides of reservoir to expel contents until chamber is properly filled.
(14) Place hemacytometer in moist Petri dish for 10 minutes to allow platelets to settle. (Moistened filter paper retains evaporation of diluted specimen while standing.)
(15) Mount the hemacytometer on the microscope and lower its condenser.
(16) Procedure for counting platelets:
• Under 40x magnification, scan to ensure even distribution. Platelets are counted in all twenty-five small squares within the large center square. Platelets appear greenish, not refractile.
• Count cells starting in the upper left of the large middle square. Continue counting to the right hand square, drop down to the next row; continue counting in this fashion until the total area in that middle square (all 25 squares) have been counted.
• Count all cells that touch any of the upper and left lines, do not count any cell that touches a lower or right line.
• Count both sides of the hemocyt-ometer and take the average.
Ns x As x Ds
Total number of cells= 230
Number of squares counted: 1
Area of each square: 1 mm3
Depth of solution: 0.1mm
cells/mm3 = 230 x 100
1 x 1 mm2 x 0.01 mm
= 230 x 103/L
• 150 - 450 x 109/L
1. Brown, B.A., Haemotology, Principles and Procedures, Lea & Febiger, U.S.A., 1976.
2. Hoffbrand, A. V. and Pettit, 1. E., Essential Haemotology, Blackwell Scientific Publication, U.S.A., 1980.
3. Kassirsky, I. and Alexeev, G., Clinical Haemotology, Mir Publishers, U.S.S.R., 1972.
4. Widmann, F.K., Clinical interpretation of Laboratory tests, F.A. Davis Company, U.S.A., 1985.
5. Kirk, C.J.C. et al, Basic Medical Laboratory Technology, Pitman Book Ltd., U.K. 1982.
6. Green, J.H., An Introduction to human Physiology, Oxford University Press, U.K., 1980.
- Manual or microscopic method
- Automated method
(4) Pasteur pipette
(5) Test tube (75 × 12 mm).
WBC diluting fluid (Turk’s fluid) consists of a weak acid solution (which hemolyzes red cells) and gentian violet (which stains leucocyte nuclei deep violet). Diluting fluid also suspends and disperses the cells and facilitates counting. Its composition is as follows:
• Acetic acid, glacial 2 ml
• Gentian violet, 1% aqueous 1 ml
• Distilled water to make 100 ml
EDTA anticoagulated venous blood or blood obtained by skin puncture is used. (Heparin should not be used since it causes leukocyte clumping). While collecting capillary blood from the finger, excess squeezing should be avoided so as not to dilute blood with tissue fluid.
(1) Dilution of blood: Take 0.38 ml of diluting fluid in a test tube. To this, add exactly 20 μl of blood and mix. This produces 1:20 dilution. Alternatively, 0.1 ml of blood can be added to 1.9 ml of diluting fluid to get the same dilution.
(2) Charging the counting chamber: Place a coverglass over the hemocytometer. Draw some of the diluted blood in a Pasteur pipette. Holding the Pasteur pipette at an angle of 45° and placing its tip between the coverglass and the chamber, fill one of the ruled areas of the hemocytometer with the sample. The sample should cover the entire ruled area, should not contain air bubbles, and should not flow into the side channels. Allow 2 minutes for settling of cells.
= Nw x 20 x 10
= Nw x 50
Where Nw is the number of WBCs counted, Cd is the correction of dilution, Cv is the correction of volume and NLS is the number of large squares counted.
If TLC is more than 50,000/ml, then dilution of blood should be increased to 1:40 to increase the accuracy of the result.
If TLC is less than 2,000/ml then lesser dilution should be used.
Expression of TLC: Conventionally, TLC is expressed as cells/μl or cells/cmm or cells/mm3. In SI units, TLC is expressed as cells × 109/liter. Conversion factors for conventional to SI units is 0.001 and SI to conventional units is 1000.
NRBC + 100
• At birth: 10,000-26000/μl
• 1 year: 6,000-16,000/μl
• 6-12 years: 5,000-13,000/μl
• Pregnancy: up to 15,000/μl
2. Lewis SM, Bain BJ, Bates I. Dacie and Lewis Practical Haematology (9th Ed). London. Churchill Livingstone, 2001.
3. The Expert Panel on Cytometry of the International Council for Standardization in Haematology: Recommended methods for the visual determination of white blood cell count and platelet count. World Health Organization. WHO/DIL/00.3. 2000.
4. World Health Organization. Manual of Basic Techniques for a Health Laboratory (2nd Ed). Geneva: World Health Organization, 2003.