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Automated hematology analyzers work on different principles:
 
  • Electrical impedance
  • Light scatter
  • Fluorescence
  • Light absorption
  • Electrical conductivity.
 
Most analyzers are based on a combination of different principles.
 
(1) Electrical impedance: This is the classic and timetested technology for counting cellular elements of blood. As this method of cell counting was first developed by Coulter Electronics, it is also called as Coulter principle (see Figure 811.1). Two electrodes placed in isotonic solutions are separated by a glass tube having a small aperture. A vacuum is applied and as a cell passes through the aperture, flow of current is impeded and a voltage pulse is generated.
 
Figure 811.1 Coulter principle of electrical impedance
Figure 811.1 Coulter principle of electrical impedance
 
The requisite condition for cell counting by this method is high dilution of sample so that minimal numbers of cells pass through the aperture at one point of time. There are two electrodes on either side of the aperture; as the solution in which the cells are suspended is an electrolyte solution, an electric current is generated between the two electrodes. When a cell passes through this narrow aperture across which a current is flowing, change in electrical resistance (i.e. momentary interruption of electrical current between the two electrodes) occurs. A small pulse is generated due to a temporary increase in impedance. This pulse is amplified, measured, and counted. The height of the pulse is proportional to cell volume. The width of the pulse corresponds with the time required for the cell to traverse the aperture. Cells that do not pass through the center of the aperture generate a distorted pulse that is not representative of the cell volume. Some analyzers use hydrodynamic focusing to force the cells through the central path so that all cells take the same path for volume measurement.
 
An anticoagulated whole blood sample is aspirated into the system, divided into two portions, and mixed with a diluent. One dilution is passed to the red cell aperture bath (for red cell and platelet counting), and the other is delivered to the WBC aperture bath (where a reagent is added for lysis of red cells and release hemoglobin; this portion is used for leukocyte counting followed by estimation of hemoglobin). Particles between 2-20 fl are counted as platelets, while those between 36-360 fl are counted as red cells. Hemoglobin is estimated by light transmission at 535 nm.
 
(2) Light scatter: Each cell flows in a single line through a flow cell. A laser device is focused on the flow cell; as the laser light beam strikes a cell it is scattered in various directions. One detector captures the forward scatter light (forward angle light scatter or FALS) that is proportional to cell size and a second detector captures side scatter (SS) light (90°) that corresponds to the nuclear complexity and granularity of cytoplasm. This simultaneous measurement of light scattered in two directions is used for distinguishing between granulocytes, lymphocytes, and monocytes.
 
(3) Fluorescence: Cellular fluorescence is used to measure RNA (reticulocytes), DNA (nucleated red cells), and cell surface antigens.
 
(4) Light absorption: Concentration of hemoglobin is measured by absorption spectrophotometry, after conversion of hemoglobin to cyanmethemoglobin or some other compound. In some analyzers, peroxidase cytochemistry is used to classify leukocytes; the peroxidase activity is determined by absorbance.
 
(5) Electrical conductivity: Some analyzers use conductivity of high frequency current to determine physical and chemical composition of leucocytes for their classification.
 
Further Reading:
 
AUTOMATED HEMOTOLOGY ANALYZER
 
Automation is a process of replacement of tasks hitherto performed by humans by computerized methods.
 
Until recently, hematological tests were performed only by manual methods. These methods, though still performed in many peripheral laboratories, are laborintensive, and involve use of hemocytometers (counting chambers), centrifuges, Wintrobe tubes, photometers, and stained blood smears. Hematology cell analyzers can generate the blood test results rapidly and also perform additional tests not possible by manual technology.
 
Both manual and automated laboratory techniques have advantages and disadvantages, and it is unlikely that one will completely replace the other.
 
Advantages of Automated Hematology Analyzer
 
  • Speed with efficient handling of a large number of samples.
  • Accuracy and precision in quantitative blood tests.
  • Ability to perform multiple tests on a single platform.
  • Significant reduction of labor requirements.
  • Invaluable for accurate determination of red cell indices.
 
Disadvantages of Automated Hematology Analyzer
 
  • Flags: Flagging of a laboratory test result demands labour-intensive manual examination of a blood smear.
  • Comments on red cell morphology cannot be generated. Abnormal red cell shapes (such as fragmented cells) cannot be recognized.
  • Erroneously increased or decreased results due to interfering factors.
  • Expensive with high running costs.
 
Automated hematology analyzers are of two main types:
 
  • Semi-automated: Some steps like dilution of blood sample are performed by the technologist; can measure only a few parameters.
  • Fully automated: Require only anticoagulated blood sample; measure multiple parameters.
CAUSES OF ERRONEOUS RESULTS (INTERFERENCES CAUSING ABNORMAL RESULT)
 
These are listed in Table 809.1
 
Table 809.1 Causes of erroneous results with hematology analyzer
Parameter Interfering factors
  Erroneous increase Erroneous decrease
0. All parameters
  • Clotted sample
1. WBC count
  • Nucleated red cells
  • Large platelet clumps
  • Unlysed red cells (some abnormal red cells resist lysing)
  • Cryoglobulins
  • Clotted sample
2. RBC count
  • Very high WBC*
  • Large numbers of giant platelets
  • Clotted sample
  • Microcytic red cells
  • Autoagglutination
3. Hemoglobin
  • Clotted sample
4. MCV
  • Very high WBC
  • Hyperglycemia
  • Autoagglutination (cold agglutinins)
  • Cryoglobulins
5. MCHC
  • Hyperlipidemia
  • Autoagglutination (cold agglutinins)
  • Very high WBC
6. Platelets
  • Microcytic red cells
  • WBC fragments
  • Cryoglobulins
  • Platelet satellitism
  • Platelet clumping
*: WBCs are counted along with RBCs, but normally their number is statistically insignificant
 
 
FLAGGING
 
‘Flags’ are signals that occur when an abnormal result is detected by the analyzer. Flags are displayed to reduce false-positive and false-negative results by mandating a review of blood smear examination.
Parameters measured by hematology analyzers and their derivation are shown in Tables 808.1 and 808.2. Most automated hematology analyzers measure red cell count, red cell indices (mean cell volume, mean cell hemoglobin, mean cell hemoglobin concentration), hemoglobin, hematocrit, total leukocyte count, differential leukocyte count (three-part or five-part), and platelet count.
 
Table 808.1 Parameters measured by hematology analyzers
Parameters measured by most analyzers Parameters measured by some analyzers
  • RBC count
  • Hemoglobin
  • Mean cell volume
  • Mean cell hemoglobin
  • Mean cell hemoglobin concentration
  • WBC count
  • WBC differential
  • Platelet count
  • Red cell distribution width
  • Reticulocyte count
  • Reticulocyte hemoglobin content
  • Mean platelet volume
  • Platelet distribution width
  • Reticulated platelets
 
Table 808.2 Parameters reported by hematology analyzers
Parameters measured directly or derived through histogram Parameters measured through calculation
  • RBC count
  • Mean cell volume (Derived from RBC histogram)
  • Red cell distribution width (Derived from RBC histogram)
  • Hemoglobin
  • Reticulocyte count
  • WBC count
  • Differential WBC count (Derived through WBC histogram)
  • Platelet count
  • Mean platelet volume (Derived from platelet histogram)
  • Hematocrit
  • Mean cell hemoglobin
  • Mean cell hemoglobin concentration
 
Estimation of Hemoglobin
 
Hemoglobin is measured directly by a modification of cyanmethemoglobin method (all hemoglobins are converted to cyanmethemoglobin by potassium ferricyanide; cyanmethemoglobin has a broad absorbance peak at 540 nm). Some analyzers use a nonhazardous reagent such as sodium lauryl sulphate. A non-ionic detergent is added for rapid red cell lysis and to minimize turbidity caused by cell membranes and plasma lipids.
 
Estimation of Red Blood Cell Count and Mean Cell Volume (MCV)
 
Red cell count and cell volume are directly measured by aperture impedance or light scatter analysis. In a red cell histogram, cell numbers are plotted on Y-axis, while cell volume is indicated on Xaxis (see Figure 808.1). The analyzer counts those cells as red cells volume of which ranges between 36 fl and 360 fl. MCV is used for morphological classification of anemia into microcytic, macrocytic, and normocytic types.
 
Figure 808.1 Diagrammatic representation of red cell histogram obtained by aperture impedance
Figure 808.1 Diagrammatic representation of red cell histogram obtained by aperture impedance. The analyzer counts cells between 36 fl and 360 fl as red cells. Although leukocytes are present and counted along with red cells in the diluting fluid, their number is not statistically significant. Only if leukocyte count is markedly elevated (>50,000/μl), histogram and the red cell count will be affected. Area of the peak between 60 fl and 125 fl is used for calculation of mean cell volume and red cell distribution width. Abnormalities in red cell histogram include: (1) Left shift of the curve in microcytosis, (2) Right shift of the curve in macrocytosis, and (3) Bimodal peak of the curve in double (dimorphic) population of red cells
 
Estimation of MCH, MCHC, and Hematocrit (HCT/PCV)
 
These parameters are obtained indirectly through calculations.
 
 
MCH (pg) = Hemoglobin (g/l)
                     RBC count (10⁶/μl)
 
 
MCHC (g/dl) = Hemoglobin (g/dl)
                         Hematocrit (%)
 
 
Hematocrit (%) = Mean Cell Volume (fl)
                              RBC count (10⁶/μl)
 
 
Estimation of Red Cell Distribution Width (RDW)
 
RDW is a quantitative measure of variation in sizes of red cells and is expressed as coefficient of variation of red cell size distribution. It is equivalent to anisocytosis observed on blood smear. It is derived from red cell histogram in some analyzers. RDW is usually elevated in iron deficiency anemia, but not in β-thalassemia minor and anemia of chronic disease (other causes of microcytic anemia). However, this distinction is not absolute and there is a significant overlap between values among patients. Raised RDW requires examination of blood smear.
 
Among the red cell values generated by the analyzer (red cell count, hemoglobin, hematocrit, MCV, MCH, MCHC, and RDW), most important for decision-making are hemoglobin, hematocrit, and MCV.
 
WBC Differential
 
Difference between 3-part and 5-part hemotology analyzer...
 
Hematology analyzers can either generate a 3-part differential (differential count reported as lymphocytes, monocytes, and granulocytes) or a 5-part differential (lymphocytes, monocytes, neutrophils, eosinophils, and basophils). The 3-part differential counting is based on electrical impedance volume measurement of leukocytes. In volume histogram for WBCs, approximate numbers of cells are plotted on Y-axis and cell size on X-axis. Those cells with volume 35-90 fl are designated as lymphocytes, cells with volume 90-160 fl as mononuclear cells, and cells with volume 160-450 fl as neutrophils (see Figure 808.2). Any deviation from the expected histogram is flagged by the analyzer, mandating review of blood smear. A large proportion of 3-part differential counts are ‘flagged’ to avoid missing abnormal cells.
 
Instruments measuring a 5-part differential work on a combination of different principles, e.g. light scatter, impedance, and electrical conductivity, a combination of light scatter, peroxidase staining, and resistance of basophils to lysis in acid buffer, etc.
 
Figure 808.2 Diagrammatic representation of WBC histogram
Figure 808.2 Diagrammatic representation of WBC histogram. WBC histogram analysis shows relative numbers of cells on Y-axis and cell size on X-axis. The lytic agent lyses the cytoplasm that collapses around the nucleus causing differential shrinkage. The analyzer sorts the WBCs according to the nuclear size into 3 main groups (3-part differential): Cells with 35-90 fl volume are designated as lymphocytes, cells with 90-160 fl volume are designated as monocytes, and cells with 160-450 fl volume are designated as neutrophils. Abnormalities in WBC histogram include: (1) Peak to the left of lymphocyte peak: Nucleated red cells, (2) Peak between lymphocytes and monocytes: Blast cells, eosinophilia, basophilia, plasma cells, and atypical lymphocytes, and (3) Peak between monocytes and neutrophils: Left shift
 
Platelet Count
 
Platelets are difficult to count because of their small size, marked variation in size, tendency to aggregation, and overlapping of size with microcytic red cells, cellular fragments, and other debris. In hematology analyzers, this difficulty is addressed by mathematical analysis of platelet volume distribution so that it corresponds to lognormal distribution. Platelets are counted by electrical impedance method in the RBC aperture, and a histogram is generated with platelet volume on X-axis and relative cell frequency on Y-axis (see Figure 808.3). Normal platelet histogram consists of a right-skewed single peak. Particles greater than 2 fl and less than 20 fl are classified as platelets by the analyzer.
 
Figure 808.3 Diagrammatic representation of normal platelet histogram
Figure 808.3 Diagrammatic representation of normal platelet histogram: Counting and sizing of platelets by electrical impedance method occurs in the RBC aperture. The counter designates particles between sizes 2 fl and 20 fl as platelets. Abnormalities in platelet histogram result from interferences such as cytoplasmic fragments (peak at left end of histogram) or severely microcytic red cells and giant platelets (peak at right end of histogram)
 
Two other platelet parameters can be obtained from platelet histogram using computer technology: mean platelet volume (MPV) and platelet distribution width (PDW). Some analyzers can generate another parameter called as reticulated platelets.
 
MPV refers to the average size of platelets and is obtained from mathematical calculation. Normal MPV is 7-10 fl. Increased MPV (> 10 fl) results from presence of immature platelets in circulation; peripheral destruction of platelets stimulates megakaryocytes to produce such platelets (e.g. in idiopathic thrombocytopenic purpura). Decreased MPV (< 7 fl) is due to presence of small platelets in circulation (in conditions associated with reduced production of platelets in bone marrow).
 
PDW is analogous to RDW and is a measure of variation in size of platelets (normal <20%). Increased PDW is observed in megaloblastic anemia, chronic myeloid leukemia, and after chemotherapy.
 
Some analyzers measure reticulated platelets or young platelets that contain RNA (similar to reticulocytes). Increased numbers of reticulated platelets are seen in thrombocytopenia due to peripheral destruction of platelets.

Reticulocyte Count
 
Various fluorescent dyes can combine with RNA of reticulocytes; the fluorescence then is counted in a flow cytometer. More immature reticulocytes fluoresce more strongly as they contain more RNA.
 
Reticulocyte hemoglobin content is a parameter that estimates hemoglobinization of most recently produced red cells. It is a predictor of iron deficiency.
 
WBC Cytogram (Scattergram)
 
In the scattergram, each dot represents a cell of a given volume and density, and the positions of dots in the graph are determined by the degree of side scatter, degree of forward scatter, light absorption by the cell, and cytochemical staining (if used). The forward angle light scatter (FALS) is represented on Y-axis, and the side scatter (SS) is represented on X-axis. Low FALS and low SS are indicative of lymphocytes; with subsequent increasing FALS and SS, monocytes, neutrophils, and lastly eosinophils are designated in the graph. Counting of basophils is based on a different technology.
 
Further Reading:
 

FLOW CYTOMETRY

  • 29 Jul 2017
FLOW CYTOMETRY
 
Box 807.1 Properties of a cell measured by a flow cytometerFlow cytometry is a procedure used for measuring multiple cellular and fluorescent properties of cells when they flow as a single cell suspension through a laser beam by a specialized instrument called as a flow cytometer. Flow cytometry can analyze numerous cells in a short time and multiple parameters of a single cell can be analyzed simultaneously. From the measured parameters, specific cell populations are defined. Cells or particles with size 0.2-150 μm are suitable for flow cytometer analysis.
 
Flow cytometry can provide following information about a cell (Box 807.1):
 
  • Cell size (forward scatter)
  • Internal complexity or granularity (side scatter)
  • Relative fluorescence intensity.
 
A flow cytometer consists of three main components or systems: fluidics, optics, and electronics.
 
(1) Fluidics: The function of the fluidics system is to transport cells in a stream to the laser beam for interrogation. Cells (fluorescence-tagged) are introduced into the cytometer (injected into the sheath fluid within the flow chamber) and made to flow in a single file past a laser (light amplification by stimulated emission of radiation) beam. The stream transporting the cells should be positioned in the center of the laser beam. The portion of the fluid stream where the cells are located is called as the sample core. Only a single cell or particle should pass through the laser beam at one time. Flow cytometers use the principle of hydrodynamic focusing (process of centering the sample core within the sheath fluid) for presenting cells to the laser.
 
(2) Optics: This system consists of lasers for illumination of cells in the sample, and filters to direct the generated light signals to the appropriate detectors.
 
The light source used in most flow cytometers is laser.
 
The laser most commonly used in flow cytometry is Argon-ion laser. The light signals are generated when the laser beam strikes the cell, which are then collected by appropriately positioned lenses. A system of optical mirrors and filters then directs the specified wavelengths of light to the designated detectors. Two types of light signals are generated when the laser beam strikes cells tagged with fluorescent molecules: fluorescence and light scatter. The cells tagged with fluorescence emit a momentary pulse of fluorescence; in addition, two types of light scatter are measured: forward scatter (proportional to cell diameter) and side scatter (proportional to granularity of cell).
 
(3) Electronics: The optical signals (photons) are converted to corresponding electronic signals (electrons) by the photodetectors (photodiodes and photomultiplier tubes). The electronic signal produced is proportional to the amount of light striking a cell. The electric current travels to the amplifier and is converted to a voltage pulse. The voltage pulse is assigned a digital value representing a channel by the Analog-to Digital Converter (ADC). The channel number is then transferred to the computer which displays it to the appropriate position on the data plot.
 
Further Reading:
 
  1. Leukemias and lympomas: Immunophenotyping (evaluation of cell surface markers), diagnosis, detection of minimal residual disease, and to identify prognostically important subgroups.
  2. Paroxysmal nocturnal hemoglobinuria: Deficiency of CD 55 and CD 59.
  3. Hematopoietic stem cell transplantation: Enumeration of CD34+ stem cells.
  4. Feto-maternal hemorrhage: Detection and quantitation of foetal hemoglobin in maternal blood sample.
  5. Anemias: Reticulocyte count.
  6. Human immunodeficiency virus infection: For enumeration of CD4+ lymphocytes.
  7. Histocompatibility cross matching.
Platelet aggregation tests are carried out in specialized hematology laboratories if platelet dysfunction is suspected. These tests are usually indicated in patients presenting with mucocutaneous type of bleeding and in whom screening tests reveal normal platelet count, prolonged bleeding time, normal prothrombin time, and normal activated partial thromboplastin time. Platelet aggregation studies are carried out on platelet-rich plasma using aggregometer. When a platelet aggregating agent is added to platelet-rich plasma, platelets form aggregates and optical density falls (or light transmission increases); this is recorded by a chart recorder on a strip chart. Commonly used platelet aggregating agents are ADP (adenosine 5-diphosphate), epinephrine (adrenaline), collagen, arachidonic acid, and ristocetin. ADP (low dose) and epinephrine induce primary and secondary waves of aggregation (biphasic curve). Primary wave is due to the direct action of aggregating agent on platelets. Secondary wave is due to thromboxane A2 synthesis and secretion from platelets. Collagen, arachidonic acid and ristocetin induce a single wave of aggregation (monophasic curve) Normal aggregation curve is shown in Figure 804.1. Aggregation patterns in various platelet function defects are shown in Figures 804.2 to 804.4, and in Table 804.1.
 
Figure 804.1 Normal platelet aggregation curves
Figure 804.1 Normal platelet aggregation curves
 
Figure 804.2 Platelet aggregation curves in von Willebrand disease and Bernard Soulier syndrome absent aggregation with ristocetin normal aggregation with ADP epinephrine and arachidonic acid
Figure 804.2 Platelet aggregation curves in von Willebrand disease and Bernard-Soulier syndrome (absent aggregation with ristocetin, normal aggregation with ADP, epinephrine, and arachidonic acid)
 
Figure 804.3 Platelet aggregation curves in storage pool defect absent second wave of aggregation with ADP and epinephrine absent or greatly diminished aggregation with collagen and normal ristocetin aggregation
Figure 804.3 Platelet aggregation curves in storage pool defect (absent second wave of aggregation with ADP and epinephrine, absent or greatly diminished aggregation with collagen, and normal ristocetin aggregation)
 
Figure 804.4 Platelet aggregation curves in Glanzmanns thrombasthenia absent aggregation with all agonists except ristocetin
Figure 804.4 Platelet aggregation curves in Glanzmann’s thrombasthenia (absent aggregation with all agonists except ristocetin)
A blood smear is examined for:
 
 
A peripheral blood smear has three parts: Head, body, and tail. Also read: PREPARATION OF BLOOD SMEAR BY WEDGE METHOD.
 
A blood smear should be examined in an orderly manner. Initially, blood smear should be observed under low power objective (10×) to assess whether the film is properly spread and stained, to assess cell distribution, and to select an area for examination of blood cells. Best morphologic details are seen in the area where red cells are just touching one another. Low power view is also helpful for the identification of Rouleaux formation, autoagglutination of red cells, and microfilaria. High power objective (45×) is suitable for examination of red cell morphology and for differential leukocyte count. A rough estimate of total leukocyte count can be obtained which also serves to crosscheck the total leukocyte count done by manual counting or automated method. Oil-immersion objective (100×) is used for more detailed examination of any abnormal cells.
 
Further Reading:
 
Box 802.1 Role of blood smear in thrombocytopeniaPlatelets are small, 1-3 μm in diameter, purple structures with tiny irregular projections on surface. In blood films prepared from non-anticoagulated blood (i.e. direct fingerstick), they occur in clumps. If platelet count is done on automated blood cell counters using EDTA-anticoagulated blood sample, about 1% of persons show falsely low count due to the presence in them of EDTA dependent antiplatelet antibody. Examination of a parallel blood film is useful in avoiding the false diagnosis of thrombocytopenia in such cases. Occasionally, platelets show rosetting around neutrophils (platelet satellitism) (see Figure 802.1). This is seen in patients with platelet antibodies and in apparently normal persons. Blood smear examination can be helpful in determining underlying cause of thrombocytopenia such as leukemia, lymphoma, or microangiopathic hemolytic anemia (Box 802.1).
 
Also Read:
 
For meaningful interpretation, absolute count of leukocytes should be reported. These are obtained as follows:
 
Absolute Leukocyte Count = Leukocyte% × Total Leukocyte Count/ml
 
 
Neutrophilia:
 
An absolute neutrophil count greater than 7500/μl is termed as neutrophilia or neutrophilic leukocytosis.
 
Causes
 
  1. Acute bacterial infections: Abscess, pneumonia, meningitis, septicemia, acute rheumatic fever, urinary tract infection.
  2. Tissue necrosis: Burns, injury, myocardial infarction.
  3. Acute blood loss
  4. Acute hemorrhage
  5. Myeloproliferative disorders
  6. Metabolic disorders: Uremia, acidosis, gout
  7. Poisoning
  8. Malignant tumors
  9. Physiologic causes: Exercise, labor, pregnancy, emotional stress.
 
Leukemoid reaction: This refers to the presence of markedly increased total leukocyte count (>50,000/cmm) with immature cells in peripheral blood resembling leukaemia but occurring in non-leukemic disorders (see Figure 801.2). Its causes are:
 
  • Severe bacterial infections, e.g. septicemia, pneumonia
  • Severe hemorrhage
  • Severe acute hemolysis
  • Poisoning
  • Burns
  • Carcinoma metastatic to bone marrow Leukemoid reaction should be differentiated from chronic myeloid leukemia (Table 801.1).
 
Table 801.1 Differences between leukemoid reaction and leukemia
Table 801.1 Differences between leukemoid reaction and leukemia
 
Figure 801.2 Leukemoid reaction in blood smear
Figure 801.2 Leukemoid reaction in blood smear
 
 
Absolute neutrophil count less than 2000/μl is neutropenia. It is graded as mild (2000-1000/μl), moderate (1000-500/μl), and severe (< 500/μl).
 
Causes
 
I. Decreased or ineffective production in bone marrow:
 
  1. Infections 
    (a) Bacterial: typhoid, paratyphoid, miliary tuberculosis, septicemia
    (b) Viral: influenza, measles, rubella, infectious mononucleosis, infective hepatitis.
    (c) Protozoal: malaria, kala azar
    (d) Overwhelming infection by any organism
  2. Hematologic disorders: megaloblastic anemia, aplastic anemia, aleukemic leukemia, myelophthisis.
  3. Drugs:
    (a) Idiosyncratic action: Analgesics, antibiotics, sulfonamides, phenothiazines, antithyroid drugs, anticonvulsants.
    (b) Dose-related: Anticancer drugs
  4. Ionizing radiation
  5. Congenital disorders: Kostman's syndrome, cyclic neutropenia, reticular dysgenesis.
 
II. Increased destruction in peripheral blood:
 
  1. Neonatal isoimmune neutropaenia
  2. Systemic lupus erythematosus
  3. Felty's syndrome
 
III. Increased sequestration in spleen:
 
  1. Hypersplenism
 
Eosinophilia:
 
This refers to absolute eosinophil count greater than 600/μl.
 
Causes
 
  1. Allergic diseases: Bronchial asthma, rhinitis, urticaria, drugs.
  2. Skin diseases: Eczema, pemphigus, dermatitis herpetiformis.
  3. Parasitic infection with tissue invasion: Filariasis, trichinosis, echinococcosis.
  4. Hematologic disorders: Chronic Myeloproliferative disorders, Hodgkin's disease, peripheral T cell lymphoma.
  5. Carcinoma with necrosis.
  6. Radiation therapy.
  7. Lung diseases: Loeffler's syndrome, tropical eosinophilia
  8. Hypereosinophilic syndrome.
 
Basophilia:
 
Increased numbers of basophils in blood (>100/μl) occurs in chronic myeloid leukemia, polycythemia vera, idiopathic myelofibrosis, basophilic leukemia, myxedema, and hypersensitivity to food or drugs.
 
Monocytosis:
 
This is an increase in the absolute monocyte count above 1000/μl.
 
Causes
 
  1. Infections: Tuberculosis, subacute bacterial endocarditis, malaria, kala azar.
  2. Recovery from neutropenia.
  3. Autoimmune disorders.
  4. Hematologic diseases: Myeloproliferative disorders, monocytic leukemia, Hodgkin's disease.
  5. Others: Chronic ulcerative colitis, Crohn's disease, sarcoidosis.
 
Lymphocytosis:
 
Box 801.1 Differential diagnosis of LymphocytosisThis is an increase in absolute lymphocyte count above upper limit of normal for age (4000/μl in adults, >7200/μl in adolescents, >9000/μl in children and infants) (Box 801.1).
 
Causes
 
  1. Infections: 
    (a) Viral: Acute infectious lymphocytosis, infective hepatitis, cytomegalovirus, mumps, rubella, varicella
    (b) Bacterial: Pertussis, tuberculosis
    (c) Protozoal: Toxoplasmosis
  2. Hematological disorders: Acute lymphoblastic leukemia, chronic lymphocytic leukemia, multiple myeloma, lymphoma.
  3. Other: Serum sickness, post-vaccination, drug reactions.
Approximate idea about total leukocyte count can be gained from the examination of the smear under high power objective (40× or 45×). A differential leukocyte count should be carried out. Abnormal appearing white cells are evaluated under oil-immersion objective.
 
Morphology of normal leukocytes (see Figure 800.1):
 
  1. Polymorphonuclear neutrophil: Neutrophil measures 14-15 μm in size. Its cytoplasm is colorless or lightly eosinophilic and contains multiple, small, fine, mauve granules. Nucleus has 2-5 lobes that are connected by fine chromatin strands. Nuclear chromatin is condensed and stains deep purple in color. A segmented neutrophil has at least 2 lobes connected by a chromatin strand. A band neutrophil shows non-segmented U-shaped nucleus of even width. Normally band neutrophils comprise less than 3% of all leukocytes. Majority of neutrophils have 3 lobes, while less than 5% have 5 lobes. In females, 2-3% of neutrophils show a small projection (called drumstick) on the nuclear lobe. It represents one inactivated X chromosome.
  2. Eosinophil: Eosinophils are slightly larger than neutrophils (15-16 μm). The nucleus is often bilobed and the cytoplasm is packed with numerous, large, bright orange-red granules. On blood smears, some of the eosinophils are often ruptured.
  3. Basophils: Basophils are seen rarely on normal smears. They are small (9-12 μm), round to oval cells, which contain very large, coarse, deep purple granules. It is difficult to make out the nucleus since granules cover it.
  4. Monocytes: Monocyte is the largest of the leukocytes (15-20 μm). It is irregular in shape, with oval or clefted (kidney-shaped) nucleus and fine, delicate chromatin. Cytoplasm is abundant, bluegray with ground glass appearance and often contains fine azurophil granules and vacuoles. After migration to the tissues from blood, they are called as macrophages.
  5. Lymphocytes: On peripheral blood smear, two types of lymphocytes are distinguished: small and large. The majority of lymphocytes are small (7-8 μm). These cells have a high nuclearcytoplasmic ratio with a thin rim of deep blue cytoplasm. The nucleus is round or slightly clefted with coarsely clumped chromatin. Large lymphocytes (10-15 μm) have a more abundant, pale blue cytoplasm, which may contain a few azurophil granules. Nucleus is oval or round and often placed on one side of the cell.
 
Figure 800.1 Normal mature white blood cells in peripheral blood
Figure 800.1 Normal mature white blood cells in peripheral blood
 
Morphology of abnormal leukocytes:
 
  1. Box 800.1 Role of blood smear in leukemiaToxic granules: These are darkly staining, bluepurple, coarse granules in the cytoplasm of neutrophils. They are commonly seen in severe bacterial infections.
  2. Döhle inclusion bodies: These are small, oval, pale blue cytoplasmic inclusions in the periphery of neutrophils. They represent remnants of ribosomes and rough endoplasmic reticulum. They are often associated with toxic granules and are seen in bacterial infections.
  3. Cytoplasmic vacuoles: Vacuoles in neutrophils are indicative of phagocytosis and are seen in bacterial infections.
  4. Shift to left of neutrophils: This refers to presence of immature cells of neutrophil series (band forms and metamyelocytes) in peripheral blood and occurs in infections and inflammatory disorders.
  5. Hypersegmented neutrophils: Hypersegmentation of neutrophils is said to be present when >5% of neutrophils have 5 or more lobes. They are large in size and are also called as macropolycytes. They are seen in folate or vitamin B12 deficiency and represent one of the earliest signs.
  6. Pelger-Huet cells: In Pelger-Huet anomaly (a benign autosomal dominant condition), there is failure of nuclear segmentation of granulocytes so that nuclei are rod-like, round, or have two segments. Such granulocytes are also observed in myeloproliferative disorders (pseudo-Pelger-Huet cells).
  7. Atypical lymphocytes: These are seen in viral infections, especially infectious mononucleosis. Atypical lymphocytes are large, irregularly shaped lymphocytes with abundant cytoplasm and irregular nuclei. Cytoplasm shows deep basophilia at the edges and scalloping of borders. Nuclear chromatin is less dense and occasional nucleolus may be present.
  8. Blast cells: These are most premature of the leukocytes. They are large (15-25 μm), round to oval cells, with high nuclear cytoplasmic ratio. Nucleus shows one or more nucleoli and nuclear chromatin is immature. These cells are seen in severe infections, infiltrative disorders, and leukemia. In leukemia and lymphoma, blood smear suggests the diagnosis or differential diagnosis and helps in ordering further tests (see Figure 800.2 and Box 800.1).
 
Figure 800.2 Morphological abnormalities of white blood cells
Figure 800.2 Morphological abnormalities of white blood cells: (A) Toxic granules; (B) Döhle inclusion body; (C) Shift to left in neutrophil series; (D) Hypersegmented neutrophil in megaloblastic anemia; (E) Atypical lymphocyte in infectious mononucleosis; (F) Blast cell in acute leukemia
 
Further Reading:
 
Role of blood smear in anemiasRed cells are best examined in an area where they are just touching one another (towards the tail of the film). Normal red cells are 7-8 μm in size, round with smooth contours, and stain deep pink at the periphery and paler in the center. Area of central pallor is about 1/3rd the diameter of the red cell. Size of a normal red cell corresponds roughly with the size of the nucleus of a small lymphocyte. Normal red cells are described as normocytic (of normal size) and normochromic (with normal staining intensity i.e. hemoglobin content).
 
Morphologic abnormalities of red cells in peripheral blood smear can be grouped as follows:
 
  • Red cells with abnormal size (see Figure 799.1)
  • Red cells with abnormal staining
  • Red cells with abnormal shape (see Figure 799.1)
  • Red cell inclusions (see Figure 799.2)
  • Immature red cells (see Figure799.3)
  • Abnormal red cell arrangement(see Figure 799.4).
 
Red cells with abnormal size:
 
Mild variation in red cell size is normal. Increased variation in red cell size is called as anisocytosis. This is a feature of most anemias and is non-specific. Anisocytosis is due to the presence of microcytes, macrocytes, or both in addition to red cells of normal size.
 
Microcytes are red cells smaller in size than normal. They are seen when hemoglobin synthesis is defective i.e. in iron deficiency anemia, thalassemias, anemia of chronic disease, and sideroblastic anemia.

Macrocytes are red cells larger in size than normal. Oval macrocytes (macro-ovalocytes) are seen in megaloblastic anemia, myelodysplastic syndrome, and in patients being treated with cancer chemotherapy. Round macrocytes are seen in liver disease, alcoholism, and hypothyroidism.
 
Red cells with abnormal staining (hemoglobin content):

Staining intensity of red cells depends on hemoglobin content. Red cells with increased area of central pallor (i.e. containing less hemoglobin) are called as hypochromic. They are seen when hemoglobin synthesis is defective, i.e. in iron deficiency, thalassemias, anaemia of chronic disease, and sideroblastic anemia.
 
In dimorphic anemia, there are two distinct populations of red cells in the same smear. An example is presence of both normochromic and hypochromic red cells seen in sideroblastic anemia, iron deficiency anemia responding to treatment, and following blood transfusion in a patient of hypochromic anemia. In myelodysplastic syndrome, dimorphic picture results from admixture of microcytic hypochromic cells and macrocytes.
 
Red cells with abnormal shape:
 
Increased variation in red cell shape is called as poikilocytosis and is a feature of many anemias. A red cell that is abnormal in shape is called as a poikilocyte.
 
Sickle cells are narrow and elongated red cells with one or both ends pointed. Sickle form is assumed when a red cell containing hemoglobin S is deprived of oxygen. Sickle cells are seen in sickle cell disorders, particularly sickle cell anemia. Sickle cells are not seen on blood smear in neonates with sickle cell disease because high percentage of fetal hemoglobin in red cells prevents sickling.
 
Spherocytes are red cells, which are slightly smaller in size than normal, round, stain intensely, and do not have central area of pallor. The surface area of spherocytes is less as compared to the volume. They are seen in hereditary spherocytosis, autoimmune hemolytic anemia (warm antibody type), and ABO hemolytic disease of newborn.
 
Schistocytes are fragmented red cells, which take various forms like helmet, crescent, triangle, etc. and usually have surface projections or spicules. They are seen in microangiopathic hemolytic anemia, cardiac valve prosthesis, and severe burns.
 
Target cells are red cells with bull's eye appearance. These red cells show a central stained area and a peripheral stained rim with unstained cytoplasm in between. They are seen in hemoglobinopathies (e.g. thalassemias, hemoglobin disease, sickle cell disease), obstructive jaundice, and following splenectomy.disease, sickle cell disease), obstructive jaundice, and following splenectomy.
 
Burr cells or echinocytes are small red cells with regularly placed small projections on surface. They are seen in uremia.
 
Acanthocytes are red cells with irregularly spaced sharp projections of variable length on surface. They are seen in spur cell anemia of liver disease, McLeod phenotype, and following splenectomy.
 
Teardrop cells or dacryocytes have a tapering droplike shape. Numerous teardrop red cells are seen in myelofibrosis and myelophthisic anemia.
 
Blister cells or hemi ghost cells are irregularly contracted cells in which hemoglobin is contracted and condensed away from the cell membrane. This is seen in glucose-6-phosphate dehydrogenase defici-ency during acute hemolytic episode.
 
Bite cells result from removal of Heinz bodies by the pitting action of the spleen (i.e. a part of red cell is bitten off by the splenic macrophages). They are seen in glucose-6-phosphate dehydrogena-se deficiency and unstable hemoglobin disease.
 
Red cell inclusions:
 
Those inclusions that can be visualized on Romanowsky-stained smears are basophilic stippling, Howell-Jolly bodies, Pappenheimer bodies, and Cabot's rings.

Basophilic stippling or punctate basophilia refers to the presence of numerous, irregular basophilic (purple-blue) granules which are uniformly distributed in the red cell. These granules represent aggregates of ribosomes. Their presence is indicative of impaired erythropoiesis and they are seen in thalassemias, megaloblastic anemia, heavy metal poisoning (e.g. lead), and liver disease.cell. These granules represent aggregates of ribosomes. Their presence is indicative of impaired erythropoiesis and they are seen in thalassemias, megaloblastic anemia, heavy metal poisoning (e.g. lead), and liver disease.
 
Red cell inclusions
Figure 799.2 Red cell inclusions: (A) Basophilic stippling; (B) Howell-Jolly bodies; (C) Pappenheimer bodies; (D) Cabot’s ring
 
Howell-Jolly bodies are small, round, purple-staining nuclear remnants located peripherally in red cells. They are seen in megaloblastic anemia, thalasse-mias, hemolytic anemia, and following splenectomy.

Pappenheimer bodies are basophilic, small, ironcontaining granules in red cells. They give positive Perl's Prussian blue reaction. Unlike basophilic stippling, Pappenheimer bodies are few in number and are not distributed throughout the red cell. They are seen following splenectomy and in thalassemias and sideroblastic anemia.

Cabot's rings are fine, reddish-purple or red, ring-like structures. They appear like loops or figure of eight structures. They indicate impaired erythropoiesis and are seen in megaloblastic anemia and lead poisoning.
 
Immature red cells:
 
Polychromatic cells are young red cells containing remnants of ribonucleic acid. These cells are slightly larger than normal red cells and have a diffuse bluishgrey tint. (They represent reticulocytes when stained with a supravital stain like new methylene blue). Polychromasia is due to the uptake of acid stain by hemoglobin and basic stain by ribonucleic acid. Presence of polychromatic cells is indicative of active erythropoiesis and are increased in hemolytic anemia, acute blood loss, and following specific therapy for nutritional anemia.and are increased in hemolytic anemia, acute blood loss, and following specific therapy for nutritional anemia.
 
Nucleated red cells are red cell precursors (erythroblasts), which are released prematurely in peripheral blood from the bone marrow. They are a normal finding in cord blood of newborns. Large number of nucleated red cells in blood smear is seen in hemolytic disease of newborn, hemolytic anemia, leukemias, myelophthisic anemia, and myelofibrosis.
 
Immature red cells
Figure 799.3 Immature red cells: (A) Polychromatic red cell; (B) Nucleated red cell
 
Abnormal red cell arrangement:
 
Rouleaux formation refers to alignment of red cells on top of each other like a stack of coins. It occurs in multiple myeloma, Waldenström's macroglobulinemia, hypergammaglobulinemia, and hyper fibrinogenemia.
 
Abnormal red cell arrangement
Figure 799.4 Abnormal red cell arrangement: (A) Rouleaux formation; (B) Autoagglutination

Autoagglutination refers to the clumping of red cells in large, irregular groups on blood smear. It is seen in cold agglutinin disease. Role of blood smear in anemia is shown in Box 799.1 and Figures 799.5 to 799.7.
 
Figure 799.5 Differential diagnosis of macrocytic anemia on blood smear
Figure 799.5 Differential diagnosis of macrocytic anemia on blood smear: (A) Megaloblastic anemia; (B) Hemolytic anemia; (C) Liver disease; (D) Myelodysplastic syndrome
 
Figure 799.6 Differential diagnosis of microcytic anemia on blood smear
Figure 799.6 Differential diagnosis of microcytic anemia on blood smear: (A) Iron deficiency anemia; (B) Thalassemia minor; (C) Thalassemia major; (D) Sideroblastic anemia
 
Figure 799.7 Differential diagnosis of hemolytic anemia on blood smear
Figure 799.7 Differential diagnosis of hemolytic anemia on blood smear. (A) Microangiopathic hemolytic anemia showing fragmented red cells, (B) Hereditary spherocytosis showing spherocytes and a polychromatic red cell, and (C) Glucose-6-phosphate dehydrogenase deficiency showing a blister cell and a bite cell
 
Further Reading:
 
The microscope is the most important piece of equipment in the clinic laboratory. The microscope is used to review fecal, urine, blood, and cytology samples on a daily basis (see Figure). Understanding how the microscope functions, how it operates, and how to care for it will improve the reliability of your results and prolong the life of this valuable piece of equipment.

Parts and functions of a compound microscope

Compound Microscope(A) Arm: Used to carry the microscope.
 
(B) Base: Supports the microscope and houses the light source.
 
(C) Oculars (or eyepieces): The lens of the microscope you look through. The ocular also magnifies the image. The total magnification can be calculated by multiplying the objective power by the ocular power. Oculars come in different magnifications, but 10× magnification is common.
 
(D) Diopter adjustment: The purpose of the diopter adjustment is to correct the differences in vision an individual may have between their left and right eyes.
 
(E) Interpupillary adjustment: This allows the oculars to move closer or further away from one another to match the width of an individual’s eyes. When looking through the microscope, one should see only a single field of view. When viewing a sample, always use both eyes. Using one eye can cause eye strain over a period of time.
 
(F) Nosepiece: The nosepiece holds the objective lenses. The objectives are mounted on a rotating turret so they can be moved into place as needed. Most nosepieces can hold up to five objectives.
 
(G) Objective lenses: The objective lens is the lens closest to the object being viewed, and its function is to magnify it. Objective lenses are available in many powers, but 4×, 10×, 40×, and 100× are standard. 4× objective is used mainly for scanning. 10× objective is considered “low power,” 40× is “high power” and 100× objective is referred to as “oil immersion.” Once magnified by the objective lens, the image is viewed through the oculars, which magnify it further. Total magnification can be calculated by multiplying the objective power by the ocular lens power.
 
For example: 100× objective lens with 10× oculars = 1000× total magnification.
 
(H) Stage: The platform on which the slide or object is placed for viewing.
 
(I) Stage brackets: Spring-loaded brackets, or clips, hold the slide or specimen in place on the stage.
 
(J) Stage control knobs: Located just below the stage are the stage control knobs. These knobs move the slide or specimen either horizontally (x-axis) or vertically (y-axis) when it is being viewed.
 
(K) Condenser: The condenser is located under the stage. As light travels from the illuminator, it passes through the condenser, where it is focused and directed at the specimen.
 
(L) Condenser control knob: Allows the condenser to be raised or lowered.
 
(M) Condenser centering screws: These crews center the condenser, and therefore the beam of light. Generally, they do not need much adjustment unless the microscope is moved or transported frequently.
 
(N) Iris diaphragm: This structure controls the amount of light that reaches the specimen. Opening and closing the iris diaphragm adjusts the diameter of the light beam.
 
(O) Coarse and fine focus adjustment knobs: These knobs bring the object into focus by raising and lowering the stage. Care should be taken when adjusting the stage height. When a higher power objective is in place (100× objective for example), there is a risk of raising the stage and slide and hitting the objective lens. This can break the slide and scratch the lens surface. Coarse adjustment is used for finding focus under low power and adjusting the stage height. Fine adjustment is used for more delicate, high power adjustment that would require fine tuning.
 
(P) Illuminator: The illuminator is the light source for the microscope, usually situated in the base. The brightness of the light from the illuminator can be adjusted to suit your preference and the object you are viewing.
What is Kohler illumination?

Kohler illumination is a method of adjusting a microscope in order to provide optimal illumination by focusing the light on the specimen. When a microscope is in Kohler, specimens will appear clearer, and in more detail.

Process of setting Kohler
 
Materials required
 
  • Specimen slide (will need tofocus under 10× power)
  • Compound microscope.
 
Kohler illumination
 
  1. Mount the specimen slide onthe stage and focus under 10×.
  2. Close the iris diaphragm completely.
  3. If the ball of light is not in the center, use the condenser centering screws to move it so that it is centered.
  4. Using the condenser adjustment knobs, raise or lower the condenser until the edges of the field becomes sharp (see Figure 797.1 and Figure 797.2).
  5. Open the iris diaphragm until the entire field is illuminated.
 
Note the blurry edges of the unfocused light
Figure 797.1 Note the blurry edges of the unfocused light
 
Adjusting the condenser height sharpens the edges of the ball of light
Figure 797.2 Adjusting the condenser height sharpens the edges of the “ball of light.”
 
When should you set/check Kohler?
 
  • During regular microscope maintenance
  • After the microscope is moved/transported
  • Whenever you suspect objects do not appear as sharp as they could be.
 
Further Reading:
 

COLLECTION OF BLOOD

It is necessary to follow a standard procedure for specimen collection to get the most accurate and trustworthy results of the laboratory test. The blood sample can be collected from the venipuncture or skin puncture for the hematological investigations.

SKIN PUNCTURE

This method is most common and mostly used in infants and small children and if the small amount of blood is required. This method is suitable for the estimation of hemoglobin, cell counts, determination of hematocrit (HCT) or packed cell volume (PCV) by micro method and preparation of blood films. Blood obtained by this method is also called as capillary blood. However, it is the mixture of blood from arterioles, venules, and capillaries. It also contains small amount of tissue fluid. In infants, blood is collected from the heel (the medial or lateral aspect of plantar surface or great toe). In adults, it is collected from the side of a middle or ring finger (distal digit) or from the earlobe. (see Figure 796.1).

A. Blood lancet and sites of B. finger puncture cross and C. heel puncture shaded areas
Figure 796.1 (A) Blood lancet and sites of (B) finger puncture (cross) and (C) heel puncture (shaded areas)

The puncture site is cleansed with the 70% ethanol or another suitable disinfectant. After drying, a puncture is made with a sterile, dry, disposable lancet, in deep to allow free flow of blood. The first drop of blood is wiped away with the dry and sterile cotton as it contains tissue fluid. After wiping the first drop of blood, next few drops of blood are collected. Excessive pressing should be avoided, as it may dilute the blood with the tissue fluid. After collection of blood, a piece of dry and sterile cotton is pressed over the puncture site till the bleeding ends. Hemoglobin, red cell count and hematocrit (HCT) or packed cell volume (PCV) are moderately higher in the blood collected from skin puncture, as compared to the venous blood. The reason behind this scenario is that platelets adhere to the puncture site and cause the lower count of platelet, and due to small sample size, instant repeat testing is not possible if the result is abnormal or unusual.

Avoid collecting blood from cold, cyanosed skin since the false elevation of values of red blood cells, white blood cells and hemoglobin will be obtained.

VENOUS BLOOD COLLECTION

Venous blood is obtained when the larger quantity of blood is needed to perform multiple tests. Different test tubes are filled with blood as per requirement of anticoagulant and blood ratio for the test. Anticoagulant is not required for the test performed by the serum.

Method

  1. Common sites of venepuncture in antecubital fossaThe best site for obtaining blood is the veins of antecubital fossa. A rubber tourniquet is applied to the upper arm (see Figure; Common sites of venepuncture in antecubital fossa (red circles)). It should not be too much tight and should not remain in a place for more than 120 seconds. To get veins more palpable and prominent, the patient is asked to make a fist.
  2. The puncture site is cleansed with the 70% ethanol or other suitable disinfectant and allowed to dry.
  3. The preferred vein is anchored by squeezing and pulling the soft tissues below the prick site with the left hand.
  4. Sterile, dry, disposable needles and syringes should be used for the collection of blood. Needle size should be 23-gauge in children and 19- to 21-gauge in adults. Venepuncture is made along with the direction of the vein and with the bevel of the needle up. Blood is withdrawn slowly. Pulling the piston quickly can cause hemolysis and collapse the vein. The tourniquet should be released as soon as the blood begins to flow into the syringe.
  5. When the required blood is collected, the patient is asked to open his/her fist. The needle is removed from the vein. A sterile alcohol swab is pressed over the puncture site. The patient is asked to press the alcohol swab over the site till the bleeding ends.
  6. The needle is removed from the syringe and the required amount of blood is carefully transferred into the test tube containing anticoagulant as per requirement of the laboratory test. If the blood is forced through the syringe without removing the needle, hemolysis can occur. Containers may be glass bottles or disposable plastic tubes with corks and flat bottom.
  7. Blood is mixed with the anticoagulant in the container thoroughly by gently inverting the container several times. The container should not be shaken strenuously as it can cause hemolysis and fizzing.
  8. Check whether the patient is dizzy and bleeding has stopped. Cover the site of puncture with a sticky bandage strip. Recapping the needle by hand can cause needle-prick injury. After the usage of disposable syringe, needles are crashed by the syringe needle destroyer and the syringe is disposed into the biohazard box. The blood container is labeled properly with the patient’s name, age, gender and the time of collection. The sample should be sent without delay to the laboratory with accompanying properly filled laboratory requisition form.

Precautions

  1. The tourniquet should not be too tight and should not be applied for more than 120 seconds as it will cause hemoconcentration and variation of test results.
  2. The tourniquet should be released before removing the needle from the vein to prevent the formation of a hematoma.
  3. Blood is never collected from the arm being used for the intravenous line since it will dilute the blood sample.
  4. Blood is never collected from an area with hematoma and from a sclerosed vein.
  5. A small bore needle should not be used, blood is withdrawn gradually and the needle is removed from the syringe before transferring blood into the container to avoid hemolysis.
  6. Proper precautions should be noticed while collecting blood either from a skin or a vein puncture since all blood samples are considered as infectious.
  7. The anticoagulated blood sample should be tested within 1-2 hours of collection. If this is not possible, the sample can be stored for 24 hours in a refrigerator at 4-6° C. After the sample is taken out of the refrigerator, it should be allowed to return to room temperature, mixed properly, and then laboratory test is performed.

Complications

  1. Failure to obtain blood: This is very common and usually painful for the patient. This happens if the vein is missed, or excessive pull is applied to the piston causing collapse of the vein.
  2. Formation of hematoma, abscess, thrombosis, thrombophlebitis, or bleeding.
  3. Transmission of infection like human immunodeficiency virus (HIV) or hepatitis B virus (HBV) if reusable syringes and needles, which are not properly sterilized, are used.

Further Reading:

SEQUENCE OF FILLING OF TUBES
 
Following order of filling of tubes should be followed after withdrawal of blood from the patient if multiple investigations are ordered:
 
  1. First tube: Blood culture.
  2. Second tube: Plain tube (serum).
  3. Third tube: Tube containing anticoagulant (EDTA, citrate, or heparin).
  4. Fourth tube: Tube containing additional stabilizing agent like fluoride.
 
Further Reading:
 
Plasma is the supernatant liquid obtained after centrifugation of anticoagulated whole blood.
 
Serum is the liquid obtained after clotting of whole blood sample collected in a plain tube.
 
Some of the differences between the two are as follows:
 
  1. Plasma contains fibrinogen as well as all the other proteins, while serum does not contain fibrinogen.
  2. Plasma can be obtained immediately after sample collection by centrifugation, while minimum of 30 minutes are required for separation of serum from the clotted blood.
  3. Amount of sample is greater with plasma than with serum for a given amount of blood.
  4. Use of anticoagulant may alter the concentration of some constituents if they are to be measured like sodium, potassium, lithium, etc.
Plain tubes (i.e. without any anticoagulant) are used for chemistry studies after separation of serum: liver function tests (total proteins, albumin, aspartate aminotransferase, alanine aminotransferase, bilirubin), renal function tests (blood urea nitrogen, creatinine), calcium, lipid profile, electrolytes, hormones, and serum osmolality. Fluoride bulb is used for collection of whole blood for estimation of blood glucose. Addition of sodium fluoride (2.5 mg/ml of blood) maintains stable glucose level by inhibiting glycolysis. Sodium fluoride is commonly used along with an anticoagulant such as potassium oxalate or EDTA.
The International Council for Standardization in Haematology (ICSH) was initiated as a standardization committee by the European Society of Haematology (ESH) in 1963 and officially constituted by the International Society of Hematology (ISH) and the ESH in Stockholm in 1964. The ICSH is recognised as a Non-Governmental Organisation with official relations to the World Health Organisation (WHO).
 
The ICSH is a not-for-profit organisation that aims to achieve reliable and reproducible results in laboratory analysis in the field of diagnostic haematology.
 
The ICSH coordinates Working Groups of experts to examine laboratory methods and instruments for haematological analyses, to deliberate on issues of standardization and to stimulate and coordinate scientific work as necessary towards the development of international standardization materials and guidelines.
Anticoagulants used for hematological investigations are ethylene diamine tetra-acetic acid (EDTA), heparin, double oxalate, and trisodium citrate (Table 791.1).
 
Table 791.1 Salient features of three main anticoagulants used in the hematology laboratory
Salient features of three main anticoagulants used in the hematology laboratory
 
Ethylene Diamine Tetra-acetic Acid (EDTA)
 
Changes occurring due to prolonged storage of blood in EDTAThis is also called as Sequestrene or Versene. This is the recommended anticoagulant for routine hematological investigations. However, it cannot be used for coagulation studies. Disodium and dipotassium salts of EDTA are in common use. International Committee for Standardization in Hematology recommends dipotassium EDTA since it is more soluble. It is used in a concentration of 1.5 mg/ml of blood. Dried form of anticoagulant is used as it avoids dilution of sample. Its mechanism of action is chelation of calcium. Proportion of anticoagulant to blood should be maintained. EDTA in excess of 2mg/ml causes shrinkage of and degenerative changes in red and white blood cells, decrease in hematocrit, and increase in mean corpuscular hemoglobin concentration. Excess EDTA also causess welling and fragmentation of platelets, which leads to erroneously high platelet counts. Prolonged storage of blood in EDTA anticoagulant leads to alterations as shown in Figure 791.1 and Box 791.1. EDTA is used for estimation of hemoglobin, hematocrit, cell counts, making blood films, sickling test, reticulocyte count, and hemoglobin electrophoresis.
 
Preparation
 
Dipotassium EDTA 20 gm
Distilled water 200 ml
 
Mix to dissolve. Place 0.04 ml of this solution in a bottle for 2.5 ml of blood. Anticoagulant should be dried on a warm bench or in an incubator at 37°C before use. For routine hematological investigations, 2-3 ml of EDTA blood is required.
 
Changes in blood cell morphology crenation of red cells separation of nuclear lobes of neutrophil vacuoles in cytoplasm and irregular lobulation of monocyte and lymphocyte nuclei due to storage of blood in EDTA anti
Figure 791.1 Changes in blood cell morphology (crenation of red cells, separation of nuclear lobes of neutrophil, vacuoles in cytoplasm, and irregular lobulation of monocyte and lymphocyte nuclei) due to storage of blood in EDTA anticoagulant for prolonged time
 
Heparin
 
Heparin prevents coagulation by enhancing the activity of anti-thrombin III (AT III). AT III inhibits thrombin and some other coagulation factors. It is used in the proportion of 15-20 IU/ ml of blood. Sodium, lithium, or ammonium salt of heparin is used. Heparin should not be used for total leukocyte count (since it causes leukocyte clumping) and for making of blood films (since it imparts a blue background). It is used for osmotic fragility test (since it does not alter the size of cells) and for immunophenotyping.
 
Double Oxalate (Wintrobe Mixture)
 
This consists of ammonium oxalate and potassium oxalate in 3:2 proportion. This combination is used to balance the swelling of red cells caused by ammonium oxalate and shrinkage caused by potassium oxalate. Mechanism of anticoagulant action is removal of calcium. It is used for routine hematological tests and for estimation of erythrocyte sedimentation rate by Wintrobe method. As it causes crenation of red cells and morphologic alteration in white blood cells, it cannot be used for making of blood films.
 
Preparation
 
Ammonium oxalate 1.2 gm
Potassium oxalate 0.8 gm
Distilled water upto 100 ml
 
Place 0.5 ml of this solution in a bottle for 5 ml of blood. Anticoagulant should be dried in an incubator at 37°C or on a warm bench before use.
 
Trisodium Citrate (3.2%)
 
This is the anticoagulant of choice for coagulation studies and for estimation of erythrocyte sedimentation rate by Westergren method.
 
Preparation
 
Trisodium citrate 3.2 gm
Distilled water upto 100 ml
 
Mix well to dissolve. Store in a refrigerator at 2-8°C.
 
Use 1:9 (anticoagulant: blood) proportion for coagulation studies; for ESR, 1:4 proportion is recommended.
 
ESR should be measured within 4 hours of collection of blood, while coagulation studies should be performed within 2 hours.
 
Further Reading:
 
ABO Grouping

There are two methods for ABO grouping:
 
  • Cell grouping (forward grouping): Red cells are tested for the presence of A and B antigens employing known specific anti-A and anti-B (and sometimes anti-A, B) sera.
  • Serum grouping (reverse grouping): Serum is tested for the presence of anti-A and anti-B antibodies by employing known group A and group B reagent red cells.

Both cell and serum grouping should be done since each test acts as a check on the other.
 
There are three methods for blood grouping: slide, tube and microplate. Tube and microplate methods are better and are employed in blood banks.
 
Further Reading:
 
  1. Autoagglutination: Presence of IgM autoantibodies reactive at room temperature in patient’s serum can lead to autoagglutination. If autocontrol is not used, blood group in such a case will be wrongly typed as AB. Therefore, for correct result, if autocontrol is also showing agglutination, cell grouping should be repeated after washing red cells with warm saline, and serum grouping should be repeated at 37°C.
  2. Rouleaux formation: Rouleux formation refers to red cells adhering to each other like a stack of coins and can be mistaken for agglutination. Rouleaux formation is caused by high levels of fibrinogen, immunoglobulins, or intravenous administration of a plasma expander such as dextran. Rouleaux formation (but not agglutination) can be dispersed by addition of normal saline during serum grouping.
  3. False-negative result due to inactivated antisera: For preservation of potency of antisera, they should be kept stored at 4°-6°C. If kept at room temperature for long, antisera are inactivated and will give false-negative result.
  4. Age: Infants start producing ABO antibodies by 3-6 months of age and serum grouping done before this age will yield false-negative result. Elderly individuals also have low antibody levels.
D antigen is the most immunogenic after ABO antigens and therefore red cells are routinely tested for D. Individuals are called as Rh-positive or Rh-negative depending on presence or absence of D antigen on their red cells. Following transfusion of Rhpositive blood to Rh-negative persons, 70% of them will develop anti Rh-D antibodies. This is of particular importance in women of childbearing age as anti-D antibodies can crosss the placenta during pregnancy and destroy Dpositive fetal red cells and cause hemolytic disease of newborn. In other sensitized individuals, reexposure to D antigen can cause hemolytic transfusion reaction.
 
In Rh D grouping, patient’s red cells are mixed with anti-D reagent. Serum or reverse grouping is not carried out because most Rhnegative persons do not have anti-D antibodies; anti-D develops in Rh-negative individuals only following exposure to Rh-positive red cells.
 
Rh typing is done at the same time as ABO grouping. Method of Rh D grouping is similar in principle to ABO grouping. Since serum or reverse grouping is not possible, each sample is tested in duplicate. Dosage effect (stronger antigenantibody reaction in homozygous cells i.e. stronger reaction with DD) is observed with antigens of the Rh system. Autocontrol (patient’s red cell + patient’s serum) and positive and negative controls are included in every test run. Monoclonal IgM anti-D antiserum should be used for cell grouping, which allows Rh grouping to be caried out at the same time as ABO grouping at room temperature. With monoclonal antisera, most weak and variant forms of D antigen are detected and further testing for weak forms of D antigen (Du) is not required. Differences between ABO and Rh grouping are shown in Table 788.1.
 
Table 788.1 Comparison of ABO grouping and Rh typing
Comparison of ABO grouping and Rh typing
Microplate is a polystyrene plate consisting of 96 micro wells of either U- or V-shape. Grouping is carried out in micro wells. This method is sensitive and ideal for large number of samples (see Figure 787.1).
 
Further reading: Rh D GROUPING METHOD
Principle
 
Red cells from the specimen are reacted with reagent antisera (anti-A and anti-B). Agglutination of red cells indicates presence of corresponding antigen (agglutinogen) on red cells.
 
Specimen
 
Capillary blood from finger prick, or venous blood collected in EDTA anticoagulant.
 
Reagents
 
ABO antisera: See box 786.1 and Figure 786.1.
 
BOX ABO antisera
Box 786.1: ABO antisera
 
Anti A and anti B sera used for cell grouping
 Figure 786.1 Anti-A and anti-B sera used for cell grouping
 
Method
 
  1. A clean and dry glass slide is divided into two sections with a glass marking pencil. The sections are labeled as anti-A and anti-B to identify the antisera (see Figure 786.2).
  2. Place one drop of anti-A serum and one drop of anti-B serum in the center of the corresponding section of the slide. Antiserum must be taken first to ensure that no reagents are missed.
  3. Add one drop of blood sample to be tested to each drop of antiserum.
  4. Mix antiserum and blood by using a separate stick or a separate corner of a slide for each section over an area about 1 inch in diameter.
  5. By tilting the slide backwards and forwards, examine for agglutination after exactly two minutes.
  6. Result:
    Positive (+): Little clumps of red cells are seen floating in a clear liquid.
    Negative (–): Red cells are floating homogeneously in a uniform suspension.
  7. Interpretation: Interpret the result as shown in the Table 786.1 and Figure 786.2.
 
Table 786.1 Interpretation of cell grouping (forward grouping) by slide test
Anti-A Anti-B Blood Group
+ - A
- + B
+ + AB
- - O
 
Cell grouping by slide method
Figure 786.2 Cell grouping by slide method
 
Slide test is quick and needs only simple equipment. It can be used in blood donation camps and in case of an emergency. However, it is not recommended as a routine test in blood banks since weakly reactive antigens on cells on forward grouping and low titer anti-A and anti-B on reverse grouping may be missed. Also, drying of the reaction mixture at the edges causes aggregation that may be mistaken for agglutination. Results of slide test should always be confirmed by cell and serum grouping by tube method.
Test tube method is more reliable than slide test, but takes longer time and more equipment. For cell grouping, patient’s saline-washed red cells are mixed with known antiserum in a test tube; the mixture is incubated at room temperature, and centrifuged. For serum grouping, patient’s serum is mixed with reagent red cells of known group (available commercially or prepared in the laboratory), incubated at room temperature, and centrifuged (See Table). Following centrifugation, a red cell button (sediment) will be seen at the bottom of the tube. Cell button is dislodged by gently tapping the base of the tube and examined for agglutination.
 
Positive (+) Test
 
Clumps of red cells suspended in a clear fluid. Agglutination in tube test is graded from 1+ to 4+ and read macroscopically (See Figure). 
 
Grading of ABO tube test
Grading of ABO tube test. Negative: Uniform suspension of red cells; Grade 1 (1+): Many small clumps of red cells (fine granular appearance); Grade 2 (2+): Many large clumps with many free red cells; Grade 3 (3+): Three or four individual clumps with few free red cells; and Grade 4 (4+): One solid clump of red cells with no free red cells
 
Negative (–) Test
 
Uniform suspension of red cells.

Separate tubes of auto-control, positive control, and negative control should always be setup along with the test sample tube. Auto-control tube consists of mixture of patient’s red cells and patient’s own serum. This is required to rule out false-positive result due to auto antibodies in patient’s serum causing auto agglutination of patient’s own red cells. Auto-control test is particularly essential when ABO grouping is being done only by forward method and blood group is typed as AB. If there are auto antibodies in recipient’s serum, ABO grouping, Rh typing, antibody screening, and cross matching all will show positive result.
 
In two positive control tubes, anti-A serum is mixed with group. A red cells and anti-B is mixed with group B red cells respectively. In two negative control tubes, anti-A serum is mixed with group B red cells and anti-B serum is mixed with group. A red cells respectively. These controls are necessary to confirm that reagents are working properly.
 
Interpretation of forward (cell) and reverse (serum) grouping
Interpretation of forward cell and reverse serum grouping
 
Why test tube method of blood grouping is more reliable than slide method?
 
Test tube method of blood grouping is more reliable than slide method. This is because centrifugation enhances the reaction by bringing antigen and antibodies closer together and allows detection of weaker antigen antibody reactions; in addition drying is avoided and smaller amounts of reagent are required.
 
If forward grouping, reverse grouping, and autocontrol tests are all positive, then these results are probably indicative of a cold-reactive autoantibody. Before performing forward typing, red cells should be washed with normal saline to elute the antibody. Before performing reverse grouping, autoantibody should be adsorbed by washed cells till autocontrol is negative.
A WBC differential count gives us information regarding the proportion and numbers of individual leukocytes in the patient’s sample, including significant morphological changes. This can provide useful diagnostic information in cases of inflammation, infection, and antigenic responses.

METHOD
 
Equipment
 
Stained PBS, microscope with 100×objective lens and cell counter.
 
Procedure
 
It is important that examinationand counts be performed withinthe monolayer area of your slide
 
  1. Scan the slide in a methodical grid pattern, in order not to cover the same area twice. Counts can be completed quickly under 400×magnification, but if you are also evaluating morphology, 1000×magnification should be used.
  2. Count a minimum of 100 WBCs.
 
(If the total WBC Count is increased, 200 cells should be counted to maintain accuracy.)
 
Calculations
 
Relative count:
 
No. of Cell Type Seen = ___%
100
 
Absolute count:
 
Relative (%) x WBC Count (10³/ L) = ___ x 10³/μL
100
 
Note: Check your math:
 
• Relative counts of each cell type should add up to equal 100
• Absolute counts of each cell type should add up to equal your WBC count.
Erythrocyte (Gr. erythros, red; kytos, cell) or red blood corpuscles are circular, anucleated, highly flexible, biconcave disc-shaped cells with high edges. The sixe of each cell averages 7.2 micrometer in diameter and 2.1 micrometer in thickness. It is 1.0 micrometer thick in the center. A complex membrane surrounds it, which is a bimolecular layer of protein. There is an inner most structure, called stroma, which is composed of lipids and proteins in the form of a fibrous protein. The cell contents are 90% hemoglobin. There are two methods for estimation of erythrocyte count:
 
  • Manual or microscopic method
  • Automated method
 
MANUAL METHOD

Equipment

Hemocytometer with cover glass, compound microscope.
 
 
Hayem’s diluting solution is prepared as follows:

HgCl2               0.05 gm
NaSO4               2.5 gm
NaCl                  0.5 gm
Distilled water     100 ml
 
Specimen
 
EDTA anticoagulated venous blood or blood obtained by skin puncture is used.
 
Method
 
  1. Wipe finger with cotton soaked with alcohol, with a sterile lancet do small prick on the finger tip. Use pipette. Aspirate blood to 0.5.
  2. Aspirate diluting Hayem’s solution to the 101 mark. It will give 1:200 dilution of the blood.
  3. Hold the pipette horizontally and role it with both hands between finger and thumb.
  4. Place the counting chamber, absolutely free from dust and grease, on the table and lay the cover glass in place over the ruled area.
  5. Discard the first two or three drops from the pipette. Charge the counting chamber by holding the pipette in an inclined position. Allow 3 minutes for the cells to settle.
  6. Locate the central square, which is divided into 25 medium sized squares. Each of the medium sized squares is further divided into 16 smallest squares.
  7. Count the erythrocytes in medium sized squares (80 smallest squares) using high power objective.
  8. In order to avoid confusion in counting, count all cells wihich touch the upper and left outer double line of the group of 16 squares as if they were inside the square. Neglect all those cells, which touch the lower and right inner line.
 
Calculation
 
You may calculate total number of erythrocytes per cu mm of the blood as shown in the following.
 
Supose number of erythrocytes counted in 5 intermediate squares
 
= E
 
Area of each of the five squares in which cells are counted
 
= 1/25 sq mm
 
Therefore, total area counted
 
= 1/25 sq mm x 5
= 1/5 sq mm
 
Depth of chamber = 1/10 mm
 
Therefore, the volume in which cells are counted
 
= Area x Depth
= 1/5 sqmm x 1/10 mm
= 1/50 cu mm
 
Now, in 1/50 cu mm of diluted blood, the number of erythrocyte counted = E
 
Number of erythrocyte in one cu mm in diluted blood = E x 50
 
Since the dilution of the blood is 1 in 200, the number of erythrocytes in one cu mm of undiluted blood
 
= E x 50 x 200
 
GENERAL NOTES

(1) Increased in numbers of RBC called polycythemia it is due to
 
Congenital heart disease
• Cor pulmonale
Dehydration
• Pulmonary fibrosis
• Polycythemia vera
 
(2) Decreased in numbers of RBC is due to
 
• Anemia
Bone marrow failure
• Erythropoietin deficiency (2ndry to kidney disease)
Hemolysis (RBC destruction) from transfusion reaction
Hemorrhage
• Leukemia
• Multiple myloma
• Nutritional deficiencies of (Iron, Copper, Folate, Vit B12, B6)
 
REFERENCE RANGES

• Newborns: 4.8-7.2 millions
• Children: 3.8-5.5 millions
• Adult ( male): 4.6-6.0 millions
• Adult (Females): 4.2-5.0 millions
• Pregnancy: slightly lower than normal
 
REFERENCES
 
  • Brown, B.A., Haemotology, Principles and Procedures, Lea & Febiger, U.S.A., 1976.
  • Hoffbrand, A. V. and Pettit, 1. E., Essential Haemotology, Blackwell Scientific Publication, U.S.A., 1980.
  • Kassirsky, I. and Alexeev, G., Clinical Haemotology, Mir Publishers, U.S.S.R., 1972.
  • Widmann, F.K., Clinical interpretation of Laboratory tests, F.A. Davis Company, U.S.A., 1985.
  • Kirk, C.J.C. et al, Basic Medical Laboratory Technology, Pitman Book Ltd., U.K. 1982.
  • Green, J.H., An Introduction to human Physiology, Oxford University Press, U.K., 1980.

Principle

Anticoagulated whole blood is centrifuged in a capillary tube of uniform bore to pack the red cells. Centrifugation is done in a special microhematocrit centrifuge till packing of red cells is as complete as possible. The reading (length of packed red cells and total length of the column) is taken using a microhematocrit reader, a ruler, or arithmetic graph paper.

Equipment

  1. Microhematocrit centrifuge: It should provide relative centrifugal force of 12000 g for 5 minutes.
  2. Capillary hematocrit tubes: These are disposable glass tubes 75 mm in length and 1 mm in internal diameter. They are of two types: plain (containing no anticoagulant) and heparinised (coated with a dried film of 2 units of heparin). For plain tubes, anticoagulated venous blood is needed. Heparinised tubes are used for blood obtained from skin puncture.
  3. Tube sealant like plastic sealant or modeling clay; if not available, a spirit lamp for heat sealing.
  4. Microhematocrit reader; if not available, a ruler or arithmetic graph paper.

Specimen

Venous blood collected in EDTA (dipotassium salt) for plain tubes or blood from skin puncture collected directly in heparinised tubes. Venous blood should be collected with minimal stasis to avoid hemoconcentration and false rise in PCV.

Method

  1. Fill the capillary tube by applying its tip to the blood (either from skin puncture or anticoagulated venous blood, depending on the type of tube used). About 2/3rds to 3/4ths length of the capillary tube should be filled with blood.
  2. Seal the other end of the capillary tube (which was not in contact with blood) with a plastic sealant. If it is not available, heatseal the tube using a spirit lamp.
  3. The filled tubes are placed in the radial grooves of the centrifuge with the sealed ends toward the outer rim gasket. Counterbalance by placing the tubes in the grooves opposite to each other.
  4. Centrifuge at relative centrifu-gal force 12000 g for 5 minutes to completely pack the red cells.
  5. Immediately remove the tubes from the centrifuge and stand them upright. The tube will show three layers from top to bottom: column of plasma, thin layer of buffy coat, and column of red cells.
  6. With the microhematocrit reader, hematocrit is directly read from the scale. If hematocrit reader is not available, the tube is held against a ruler and the hematocrit is obtained by the following formula:
Length of red cell column in mm
-------------------------------------------------------
Length of total column in mm

To obtain PCV, the above result is multiplied by 100.

GENERAL NOTES

  1. Prolonged application of tourniquet during venepuncture causes hemoconcentration and rise in hematocrit.
  2. Excess squeezing of the finger during skin puncture dilutes the sample with tissue fluid and lowers the hematocrit.
  3. Correct proportion of blood with anticoagulant should be used. Excess EDTA causes shrinkage of red cells and falsely lowers the hematocrit.
  4. Inadequate mixing of blood with anticoagulant, and inadequate mixing of blood before testing can cause false results.
  5. Low hematocrit can result if there are clots in the sample.
  6. Centrifugation at lower speed and for less time falsely increases PCV.
  7. A small amount of plasma is trapped in the lower part of the red cell column which is usually insignificant. Increased amount of plasma is trapped in microcytosis, macrocytosis, spherocytosis, and sickle cell anemia, which cause an artifactual rise in hematocrit. Larger volume of plasma is trapped in Wintrobe tube than in capillary tube.
  8. As PCV requires whole blood sample, it is affected by plasma volume (e.g. PCV is higher in dehydration, and lower in fluid overload).
  9. Expression of PCV: Occasionally, PCV is expressed as a percentage. In SI units, PCV is expressed as a volume fraction. Conversion factor from conventional to SI units is 0.1 and from SI to conventional units is 100.
  10. Rules of 3 and 9: These rules of thumb are commonly used to check the accuracy of results and are applicable only if red cells are of normal size and shape.
    Hemoglobin (gm/dl) × 3 = PCV
    Red cell count (million/cmm) × 9 = PCV
  11. Automated hematocrit: In automated hematology analyzers, hematocrit is obtained by multiplying red cell count (in millions/cmm) by mean cell volume (in femtoliters).

REFERENCE RANGES

  • Adult males: 40-50%
  • Adult females (nonpregnant): 38 45%
  • Adult females (pregnant): 36-42%
  • Children 6 to 12 years: 37-46%
  • Children 6 months to 6 years: 36 42%
  • Infants 2 to 6 months: 32-42%
  • Newborns: 44-60%

CRITICAL VALUES

  • Packed cell volume: < 20% or > 60%

Principle

Anticoagulated whole blood is centrifuged in a Wintrobe tube to completely pack the red cells. The volume of packed red cells is read directly from the tube. An advantage with this method is that before performing PCV, test for erythrocyte sedimentation rate can be set up.

Equipment

  1. Wintrobe tube: This tube is about 110 mm in length and has 100 markings, each at the interval of 1 mm. Internal diameter is 3 mm. It can hold about 3 ml of blood.
  2. Pasteur pipette with a rubber bulb and a sufficient length of capillary to reach the bottom of the Wintrobe tube.
  3. Centrifuge with a speed of 2300 g.

Specimen

Venous blood collected in EDTA (1.5 mg EDTA for 1 ml of blood) or in double oxalate. Test should be performed within 6 hours of collection.

Method

  1. Mix the anticoagulated blood sample thoroughly.
  2. Draw the blood sample in a Pasteur pipette and introduce the pipette up to the bottom of the Wintrobe tube. Fill the tube from the bottom exactly up to the 100 mark. During filling, tip of the pipette is raised, but should remain under the rising meniscus to avoid foaming.
  3. Centrifuge the sample at 2300 g for 30 min (To counterbalance a second Wintrobe tube filled with blood from another patient or water should be placed in the centrifuge).
  4. Take the reading of the length of the column of red cells.

Hematocrit can be expressed either as a percentage or as a fraction of the total volume of blood sample.

Significance

In anemia, PCV is below the lower level of normal range. PCV is raised in dehydration, shock, burns, and polycythemia.

After centrifugation of anticoagulated whole blood, three zones can be distinguished in the Wintrobe tube from above downwards-plasma, buffy coat layer (a small greyish layer of white cells and platelets, about 1 mm thick), and packed red cells. Normal plasma is straw-colored. It is colorless in iron deficiency anemia, pink in the presence of hemolysis or hemoglobinemia, and yellow if serum bilirubin is raised (jaundice). In hypertriglyceridemia, plasma appears milky. Increased thickness of buffy coat layer occur if white cells or platelets are increased in number (e.g. in leukocytosis, thrombocytosis, or leukemia). Smears can be made from the buffy coat layer for demonstration of lupus erythematosus (LE) cells, malaria parasites, or immature cells.

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