Porphyrias (from Greek porphura meaning purple pigment; the name is probably derived from purple discoloration of some body fluids during the attack) are a heterogeneous group of rare disorders resulting from disturbance in the heme biosynthetic pathway leading to the abnormal accumulations of red and purple pigments called as porphyrins in the body. Heme, a component of hemoglobin, is synthesized through various steps as shown in Figure 817.1. Each of the steps is catalyzed by a separate enzyme; if any of these steps fails (due to hereditary or acquired cause), precursors of heme (porphyrin intermediates) accumulate in blood, get deposited in skin and other organs, and excreted in urine and feces. Depending on the site of defect, different types of porphyrias are described with varying clinical features, severity, and the nature of accumulated porphyrin.
Porphyria has been offered as a possible explanation for the medieval tales of vampires and werewolves; this is because of the number of similarities between the behavior of persons suffering from porphyria and the folklore (avoiding sunlight, mutilation of skin on exposure to sunlight, red teeth, psychiatric disturbance, and drinking of blood to obtain heme).
Porphyrias are often missed or wrongly diagnosed as many of them are not associated with definite physical findings, screening tests may yield false-negative results, diagnostic criteria are poorly defined and mild disorders produce an enzyme assay result within ‘normal’ range.
Heme is mainly required in bone marrow (for hemoglobin synthesis) and in liver (for cytochromes). Therefore, porphyrias are divided into erythropoietic and hepatic types, depending on the site of expression of disease. Hepatic porphyrias mainly affect the nervous system, while erythropoietic porphyrias primarily affect the skin. Porphyrias are also classified into acute and nonacute (or cutaneous) types depending on clinical presentation (Table 817.1).
Table 817.1 Various classification schemes for porphyrias
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Classification based on predominant clinical manifestations
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Classification based on site of expression of disease
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Classification based on mode of clinical presentation
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Neuropsychiatric
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Hepatic
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Acute
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1. Acute intermittent porphyria
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1. ALA-dehydratase porphyria
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1. ALA-dehydratase porphyria (Plumboporphyria)
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2. ALA-dehydratase porphyria (Plumboporphyria)
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2. Acute intermittent porphyria
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2. Acute intermittent porphyria
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Cutaneous (Photosensitivity)
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3. Hereditary coproporphyria
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3. Hereditary coproporphyria
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1. Congenital erythropoietic porphyria
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4. Variegate porphyria
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4. Variegate porphyria
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2. Porphyria cutanea tarda
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Erythropoietic porphyria
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Non-acute (cutaneous)
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3. Erythropoietic protoporphyria
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1. Congenital erythropoietic porphyria
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1. Porphyria cutanea tarda
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Mixed (Neuropsychiatric and cutaneous)
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2. Erythropoietic protoporphyria
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2. Congenital erythropoietic porphyria
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1. Hereditary coproporphyria
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Hepatic/Erythropoietic
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3. Erythropoietic protoporphyria
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2. Variegate porphyria
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1. Porphyria cutanea tarda
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Inheritance of porphyrias may be autosomal dominant or recessive. Most acute porphyrias are inherited in an autosomal dominant manner (i.e. inheritance of one abnormal copy of gene). Therefore, the activity of the deficient enzyme is 50%. When the level of heme falls in the liver due to some cause, activity of ALA synthase is stimulated leading to increase in the levels of heme precursors up to the point of enzyme defect. Increased levels of heme precursors cause symptoms of acute porphyria. When the heme level returns back to normal, symptoms subside.
Accumulation of porphyrin precursors can occur in lead poisoning due to inhibition of enzyme aminolevulinic acid dehydratase in heme biosynthetic pathway. This can mimick acute intermittent porphyria.
CLINICAL FEATURES
Clinical features of porphyrias are variable and depend on type. Acute porphyrias present with symptoms like acute and severe abdominal pain/vomiting/constipation, chest pain, emotional and mental disorders, seizures, hypertension, tachycardia, sensory loss, and muscle weakness. Cutaneous porphyrias present with photosensitivity (redness and blistering of skin on exposure to sunlight), itching, necrosis of skin and gums, and increased hair growth over the temples (Table 817.2).
Table 817.2 Clinical characteristics of porphyrias
| Porphyria | Deficient enzyme | Clinical features | Inheritance | Initial test |
| 1. Acute intermittent porphyria (AIP)* | PBG deaminase | Acute neurovisceral attacks; triggering factors+ (e.g. drugs, diet restriction) | Autosomal dominant | Urinary PBG; urine becomes brown, red, or black on standing |
| 2. Variegate porphyria | Protoporphyrinogen oxidase | Acute neurovisceral attacks + skin fragility, bullae | Autosomal dominant | Urinary PBG |
| 3. Hereditary coproporphyria | Coproporphyrinogen oxidase | Acute neurovisceral attacks + skin fragility, bullae | Autosomal dominant | Urinary PBG |
| 4. Congenital erythropoietic porphyria | Uroporphyrinogen cosynthase | Onset in infancy; skin fragility, bullae; extreme photosensitivity with mutilation; red teeth and urine (pink red urinestaining of diapers) | Autosomal recessive | Urinary/fecal total porphyrins; ultraviolet fluorescence of urine, feces, and bones |
| 5. Porphyria cutanea tarda* | Uroporphyrinogen decarboxylase | Skin fragility, bullae | Autosomal dominant (some cases) | Urinary/fecal total porphyrins |
| 6. Erythropoietic protoporphyria* | Ferrochelatase | Acute photosensitivity | Autosomal dominant | Free erythrocyte protoporphyrin |
| Disorders marked with * are the three most common porphyrias. PBG: Porphobilinogen |
Symptoms can be triggered by drugs (barbiturates, oral contraceptives, diazepam, phenytoin, carbamazepine, methyldopa, sulfonamides, chloramphenicol, and antihistamines), emotional or physical stress, infection, dieting, fasting, substance abuse, premenstrual period, smoking, and alcohol. Autosomal dominant porphyrias include acute intermittent porphyria, variegate porphyria, porphyria cutanea tarda, erythropoietic protoporphyria (most cases), and hereditary coproporphyria. Autosomal recessive porphyrias include: congenital erythropoietic porphyria, erythropoietic protoporphyria (few cases), and ALAdehydratase porphyria (plumboporphyria).
LABORATORY DIAGNOSIS
Porphyria can be diagnosed through tests done on blood, urine, and feces during symptomatic period. Timely and accurate diagnosis is required for effective management of porphyrias. Due to the variability and a broad range of clinical features, porphyrias are included under differential diagnosis of many conditions. All routine hospital laboratories usually have facilities for initial investigations in suspected cases of porphyrias; laboratory tests for identification of specific type of porphyrias are available in specialized laboratories.
INITIAL STUDIES
In suspected acute porphyrias (acute neurovisceral attack), a fresh randomly collected urine sample (10-20 ml) should be submitted for detection of excessive urinary excretion of porphobilinogen (PBG) (see Figure 817.2). In AIP, urine becomes red or brown on standing (see Figure 817.3). In suspected cases of cutaneous porphyrias (acute photosensitivity without skin fragility), free erythrocyte protporphyrin or FEP in EDTA blood (for diagnosis of erythrocytic protoporphyria) and for all other cutaneous porphyrias (skin fragility and bullae), examination of fresh, random urine (10-20 ml) and either feces (5-10 g) or plasma for excess porphyrins are necessary (see Figure 817.4 and Table 817.2).
Figure 817.2 Evaluation of acute neurovisceral porphyria
Figure 817.3 Red coloration of urine on standing in acute intermittent porphyria
Figure 817.4 Evaluation of cutaneous porphyrias
Apart from diagnosis, the detection of excretion of a particular heme intermediate in urine or feces can help in detecting site of defect in porphyria. Heme precursors up to coproporphyrinogen III are water-soluble and thus can be detected in urine. Protoporphyrinogen and Protoporphyrin are insoluble in water and are excreted in bile and can be detected in feces. All samples should be protected from light.
Samples required are
- 10-20 ml of fresh random urine sample without any preservative;
- 5-10 g wet weight of fecal sample, and
- blood anticoagulated with EDTA.
Test for Porphobilinogen in Urine
Ehrlich’s aldehyde test is done for detection of PBG. Ehrlich’s reagent (p-dimethylaminobenzaldehyde) reacts with PBG in urine to produce a red color. The red product has an absorption spectrum with a peak at 553 nm and a shoulder at 540 nm. Since both urobilinogen and porphobilinogen produce similar reaction, further testing is required to distinguish between the two. Urobilinogen can be removed by solvent extraction. (See Watson-Schwartz test). Levels of PBG may be normal or near normal in between attacks. Therefore, samples should be tested during an attack to avoid false-negative results.
Test for Total Porphyrins in Urine
Total porphyrins can be detected in acidified urine sample by spectrophotometry (Porphyrins have an intense absorbance peak around 400 nm). Semiquantitative estimation of porphyrins is possible.
Test for Total Porphyrins in Feces
Total porphyrins in feces can be determined in acidic extract of fecal sample by spectrophotometry; it is necessary to first remove dietary chlorophyll (that also absorbs light around 400 nm) by diethyl ether extraction.
Tests for Porphyrins in Erythrocytes and Plasma
Visual examination for porphyrin fluorescence, and solvent fractionation and spectrophotometry have now been replaced by fluorometric methods.
Further Testing
If the initial testing for porphyria is positive, then concentrations of porphyrins should be estimated in urine, feces, and blood to arrive at specific diagnosis (Tables 817.3 and 817.4).
Table 817.3 Diagnostic patterns of concentrations of heme precursors in acute porphyrias
| Porphyria | Urine | Feces |
| Acute intermittent porphyria | PBG, Copro III | – |
| Variegate porphyria | PBG, Copro III | Proto IX |
| Hereditary coproporphyria | PBG, Copro III | Copro III |
| PBG: Porphobilinogen; Copro III: Coproporphyrinogen III; Proto IX: Protoporphyrin IX |
Table 817.4 Diagnostic patterns of concentrations of heme precursors in cutaneous porphyrias
| Porphyria | Urine | Feces | Erythrocytes |
| Congenital erythropoietic porphyria | Uro I, Copro I | Copro I | – |
| Porphyria cutanea tarda | Uroporphyrin | Isocopro | – |
| Erythropoietic protoporphyria | – | – | Protoporphyrin |
| Uro I: Uroporphyrinogen I; Copro I: Coproporphyrinogen I; Isocopro: Isocoproporphyrinogen |
In latent porphyrias and in patients during remission, porphyrin levels may be normal; in such cases, enzymatic and DNA testing is necessary for diagnosis.
If porphyria is diagnosed, then it is necessary to investigate close family members for the disorder. Positive family members should be counseled regarding triggering factors.
This is done by flow cytometric analysis for detection of lack of GpIb/IX in Bernanrd Soulier syndrome (deficiency of CD42), and lack of GpIIb/IIIa in Glanzmann’s thrombasthenia (deficiency of CD41, CD61).
What is the best protocol for platelet glycoprotein (GPIIb/IIIa) analysis using flow cytometry?
Fresh platelets should always be used. Storing platelets dramatically changes the level of transmembrane proteins. The best way is to follow one of standardized protocols defined in: Immunophenotypic analysis of platelets. Krueger LA, Barnard MR, Frelinger AL 3rd, Furman MI, Michelson AD.Curr Protoc Cytom. 2002 Feb;Chapter 6:Unit 6.10.
D-dimer is derived from the breakdown of fibrin by plasmin and D-dimer test is used to evaluate fibrin degradation. Blood sample can be either serum or plasma. Latex or polystyrene microparticles coated with monoclonal antibody to D-dimer are mixed with patient’s sample and observed for microparticle agglutination. As the particle is small, turbidometric endpoint can be determined in automated instruments. D-dimer and FDPs are raised in disseminated intravascular coagulation, intravascular thrombosis (myocardial infarction, stroke, venous thrombosis, pulmonary embolism), and during postoperative period or following trauma. D-dimer test is commonly used for exclusion of thrombosis and thrombotic tendencies.
Further Reading:
FDPs are fragments produced by proteolytic digestion of fibrinogen or fibrin by plasmin. For determination of FDPs, blood is collected in a tube containing thrombin (to remove all fibrinogen by converting it into a clot) and soybean trypsin inhibitor (to inhibit plasmin and thus prevent in vitro breakdown of fibrin). A suspension of latex particles linked to antifibrinogen antibodies (or fragments D and E) is mixed with dilutions of patient’s serum on a glass slide. If FDPs are present, agglutination of latex particles occurs (see Figure 814.1). The highest dilution of serum at which agglutination is detected is used to determine concentration of FDPs. Increased levels of FDPs occur in fibrinogenolysis or fibrinolysis. This occurs in disseminated intravascular coagulation, deep venous thrombosis, severe pneumonia, and recent myocardial infarction.
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
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 | |
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0. All parameters
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—
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1. WBC count
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2. RBC count
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3. Hemoglobin
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4. MCV
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5. MCHC
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6. Platelets
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| *: 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 |
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Table 808.2 Parameters reported by hematology analyzers
| Parameters measured directly or derived through histogram | Parameters measured through calculation |
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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. 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. 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: 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
Flow 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:
Fluorescence
A fluorochrome absorbs light energy and emits excess energy in the form of photon light (fluorescence). Fluorescence is the property of molecules to absorb light at one wavelength and emit light at a longer wavelength. The fluorescent dyes commonly used in flow cytometry are fluorescein isothiocyanate (FITC) and phycoerythrin (PE). The fluorochrome-labeled antibodies are used for detection of antigenic markers on the surface of cells. A particular cell type can be identified on the basis of the antigenic profile expressed. Multiple fluorochromes can be used to identify different cell types in a mixed population of cells.
Light Scatter
Light is scattered when the incident light is deflected by a particle traversing through a beam of light. This depends on the physical properties of the cell. Two forms of light scatter are used to identify different cell types: forward scatter and side scatter. Forward scatter (light scattered in the same direction as the laser beam) is related to cell size. Side scatter (light scattered at a 90° angle to the laser beam) is related to internal granularity of the cell. Main subpopulations of leukocytes are identified on the basis of correlated measurements of forward and side scatters. When a cell passes through laser beam, side scatter and fluorescent signals that are emitted by the cell are directed to photomultiplier tubes, while the forward scatter signals are directed to a photodiode. Photomultiplier tubes and photodiodes are called as detectors. Optical filters are placed before the detectors that allow only a narrow range of wavelengths to reach the detectors (see Figure 806.1).
Figure 806.1 Principle of working of a flow cytometer
Data Analysis
The data collected and stored in the computer can be displayed in various formats. A parameter means forward scatter, or side scatter, or emitted fluorescence from a particle as it passes through a laser beam. A histogram is a data plot of a single parameter, with the parameter’s signal value in channel numbers or relative fluorescence intensity on X-axis (horizontal axis) and number of events on the Y-axis. A dot plot is a two parameter data graph in which each dot represents one event that the flow cytometer analyzed; one parameter is displayed on the X-axis and the other on the Y-axis. A 3-D plot represents one parameter on X-axis, another parameter on Y-axis, and number of events per channel on Z-axis.
Gating
A gate is a boundary that can be set to restrict the analysis to a specific population within the sample. For example, a gate boundary can be drawn on a dot plot or histogram to restrict the analysis only to cells with the size of lymphocytes. Gates can be inclusive (selection of events that fall within the boundary) or exclusive (selection of events that fall outside the boundary). Data selected by the gate is then displayed in subsequent plots.
Sorting
Usually, when a cell passes through the laser beam, it is sent to waste. Sorting consists of collecting cells of interest (defined through criteria of size and fluorescence) for further analysis (such as microscopy or functional or chemical analysis). Sorting feature is available only in some flow cytometers.
- Leukemias and lympomas: Immunophenotyping (evaluation of cell surface markers), diagnosis, detection of minimal residual disease, and to identify prognostically important subgroups.
- Paroxysmal nocturnal hemoglobinuria: Deficiency of CD 55 and CD 59.
- Hematopoietic stem cell transplantation: Enumeration of CD34+ stem cells.
- Feto-maternal hemorrhage: Detection and quantitation of foetal hemoglobin in maternal blood sample.
- Anemias: Reticulocyte count.
- Human immunodeficiency virus infection: For enumeration of CD4+ lymphocytes.
- 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 804.2 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 804.4 Platelet aggregation curves in Glanzmann’s thrombasthenia (absent aggregation with all agonists except ristocetin)
A blood smear is examined for:
- Red cells: Morphology, immature forms, inclusion bodies, arrangement of cells.
- White cells: Differential count, abnormal or immature forms.
- Platelets: Adequacy, abnormal forms.
- Parasites: Malaria, filaria.
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:
Platelets 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
- Acute bacterial infections: Abscess, pneumonia, meningitis, septicemia, acute rheumatic fever, urinary tract infection.
- Tissue necrosis: Burns, injury, myocardial infarction.
- Acute blood loss
- Acute hemorrhage
- Myeloproliferative disorders
- Metabolic disorders: Uremia, acidosis, gout
- Poisoning
- Malignant tumors
- 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
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:
- 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 - Hematologic disorders: megaloblastic anemia, aplastic anemia, aleukemic leukemia, myelophthisis.
- Drugs:
(a) Idiosyncratic action: Analgesics, antibiotics, sulfonamides, phenothiazines, antithyroid drugs, anticonvulsants.
(b) Dose-related: Anticancer drugs - Ionizing radiation
- Congenital disorders: Kostman's syndrome, cyclic neutropenia, reticular dysgenesis.
II. Increased destruction in peripheral blood:
- Neonatal isoimmune neutropaenia
- Systemic lupus erythematosus
- Felty's syndrome
III. Increased sequestration in spleen:
- Hypersplenism
Eosinophilia:
This refers to absolute eosinophil count greater than 600/μl.
Causes
- Allergic diseases: Bronchial asthma, rhinitis, urticaria, drugs.
- Skin diseases: Eczema, pemphigus, dermatitis herpetiformis.
- Parasitic infection with tissue invasion: Filariasis, trichinosis, echinococcosis.
- Hematologic disorders: Chronic Myeloproliferative disorders, Hodgkin's disease, peripheral T cell lymphoma.
- Carcinoma with necrosis.
- Radiation therapy.
- Lung diseases: Loeffler's syndrome, tropical eosinophilia
- 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
- Infections: Tuberculosis, subacute bacterial endocarditis, malaria, kala azar.
- Recovery from neutropenia.
- Autoimmune disorders.
- Hematologic diseases: Myeloproliferative disorders, monocytic leukemia, Hodgkin's disease.
- Others: Chronic ulcerative colitis, Crohn's disease, sarcoidosis.
Lymphocytosis:
This 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
- Infections:
(a) Viral: Acute infectious lymphocytosis, infective hepatitis, cytomegalovirus, mumps, rubella, varicella
(b) Bacterial: Pertussis, tuberculosis
(c) Protozoal: Toxoplasmosis - Hematological disorders: Acute lymphoblastic leukemia, chronic lymphocytic leukemia, multiple myeloma, lymphoma.
- 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):
- 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.
- 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.
- 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.
- 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.
- 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
Morphology of abnormal leukocytes:
Toxic granules: These are darkly staining, bluepurple, coarse granules in the cytoplasm of neutrophils. They are commonly seen in severe bacterial infections.- 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.
- Cytoplasmic vacuoles: Vacuoles in neutrophils are indicative of phagocytosis and are seen in bacterial infections.
- 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.
- 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.
- 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).
- 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.
- 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: (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:
Red 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.
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.
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.
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: (A) Megaloblastic anemia; (B) Hemolytic anemia; (C) Liver disease; (D) Myelodysplastic syndrome
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. (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:
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).
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
The 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.- The puncture site is cleansed with the 70% ethanol or other suitable disinfectant and allowed to dry.
- The preferred vein is anchored by squeezing and pulling the soft tissues below the prick site with the left hand.
- 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.
- 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.
- 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.
- 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.
- 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
- 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.
- The tourniquet should be released before removing the needle from the vein to prevent the formation of a hematoma.
- Blood is never collected from the arm being used for the intravenous line since it will dilute the blood sample.
- Blood is never collected from an area with hematoma and from a sclerosed vein.
- 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.
- Proper precautions should be noticed while collecting blood either from a skin or a vein puncture since all blood samples are considered as infectious.
- 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
- 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.
- Formation of hematoma, abscess, thrombosis, thrombophlebitis, or bleeding.
- 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:
- First tube: Blood culture.
- Second tube: Plain tube (serum).
- Third tube: Tube containing anticoagulant (EDTA, citrate, or heparin).
- Fourth tube: Tube containing additional stabilizing agent like fluoride.
Further Reading:
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:
- Plasma contains fibrinogen as well as all the other proteins, while serum does not contain fibrinogen.
- 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.
- Amount of sample is greater with plasma than with serum for a given amount of blood.
- 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
Ethylene Diamine Tetra-acetic Acid (EDTA)
This 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.
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:
- 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.
- 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.
- 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.
- 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
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 786.1: ABO antisera
Figure 786.1 Anti-A and anti-B sera used for cell grouping
Method
- 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).
- 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.
- Add one drop of blood sample to be tested to each drop of antiserum.
- 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.
- By tilting the slide backwards and forwards, examine for agglutination after exactly two minutes.
- Result:
Positive (+): Little clumps of red cells are seen floating in a clear liquid.
Negative (–): Red cells are floating homogeneously in a uniform suspension. - 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 |
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. 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
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
- 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.
- 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.
