Sherrie L. Perkins

Specimen Collection
Reliability of Tests
Cell Counts
Red Blood Cell Analysis
Leukocyte Analysis
Platelet Analysis
Advantages and Sources of Error with Automated Hematology Analyzers
Morphologic Analysis of Blood Cells
Bone Marrow Examination
Special Stains
Other Laboratory Studies

Careful assessment of the blood elements is often the first step in assessment of hematologic function and diagnosis of disease. Many hematologic disorders are defined by specific findings gleaned from tests on the blood. Blood examination often yields important diagnostic information and allows broad differential diagnostic impressions to be formed, allowing further specific testing. Therefore, careful examination of cellular morphology and quantification of each type of blood elements, as well as evaluation of a variety of parameters relating to cellular size and shape, are required. This chapter introduces the fundamental concepts that underlie laboratory evaluation of the blood and outlines additional testing that may aid in evaluating a hematologic disorder, including special stains and bone marrow examination. Limitations of such tests are also addressed.

Blood elements include erythrocytes, or red blood cells; leukocytes, or white blood cells; and platelets. Although detailed morphologic descriptions and functional characteristics of each of the cell types are included in subsequent chapters, basic features necessary for blood smear evaluation are covered in this chapter. Red blood cells are the most numerous cells in the blood and are required for tissue respiration. Erythrocytes lack nuclei and contain hemoglobin, an iron-containing protein that acts in the transport of oxygen and carbon dioxide. White blood cells serve in immune function, and include a variety of cell types that have specific functions and characteristic morphologic appearances. In contrast to red blood cells, white blood cells are nucleated. The five types of white blood cells seen normally in blood smears are neutrophils, lymphocytes, monocytes, eosinophils, and basophils. Platelets are cytoplasmic fragments that function in coagulation and hemostasis.

Evaluation of the blood requires quantification of each of the blood elements, by either manual or automated methods. Automated methods, using properly calibrated equipment, are usually more precise than manual procedures. In addition, automated methods may provide additional data describing characteristics such as cell volume. However, the automated measurements describe average cell characteristics, but do not adequately describe the scatter of values around the average value. Hence, a bimodal population of small (microcytic) and large (macrocytic) red blood cells might register a normal value for cell size. Therefore, a thorough examination of the blood also requires microscopic evaluation of a stained blood film to complement the automated blood data.


Proper specimen collection is required for reliable and accurate laboratory data to be obtained on any hematologic specimen. Before a specimen is obtained, careful thought as to what studies are needed will aid in proper handling of the material and prevent collection of inadequate or unusable specimens. Communication with laboratory personnel who will analyze the specimen is often helpful in ensuring that specimens will be handled properly and that the requested testing can be performed.

A number of factors may affect hematologic measurements, and each specimen should be collected in a standardized manner to reduce variability. Factors such as patient activity, level of patient hydration, medications, sex, age, race, smoking, and anxiety may affect hematologic parameters significantly (1,2 and 3). Thus, data such as patient age, sex, and time of specimen collection should be noted. Correlative clinical information is also extremely important in evaluating hematologic specimens. For example, a patient who has had severe diarrhea or vomiting before admission may be sufficiently dehydrated to have an erroneous increase in red blood cell concentration.

Most often, blood is collected by venipuncture into tubes containing anticoagulant. The three most commonly used anticoagulants are tripotassium or disodium salts of ethylenediaminetetraacetic acid (EDTA), trisodium citrate, and heparin. EDTA and disodium citrate act to remove calcium, which is essential for the initiation of coagulation, from the blood. Heparin acts by forming a complex with antithrombin III in the plasma to prevent the formation of thrombin. EDTA is the preferred anticoagulant for blood cell counts because it produces complete anticoagulation with minimal effects on all blood cells (4). Heparin causes a bluish coloration of the background when a blood smear is stained with one of the Romanowsky dyes but does not affect cell size or shape. Heparin is most often used for prevention of red blood cell hemolysis, for osmotic fragility testing, and for functional and immunologic analysis of leukocytes. Heparin does not completely inhibit white blood cell or platelet clumping. Trisodium citrate is the preferred anticoagulant for platelet and coagulation studies.

The concentration of the anticoagulant used may affect cell concentration measures if it is inappropriate for the volume of blood collected, and may also distort cellular morphology. Most often blood is collected directly into commercially prepared negative-pressure vacuum tubes (Vacutainer tubes), which contain the correct concentration of anticoagulant when filled appropriately, thereby minimizing error (5). Anticoagulated blood may be stored at 4° C for a 24-hour period without significantly altering cell counts or cellular morphology (6 and 7). However, it is preferable to perform hematologic analysis as soon as possible after the blood is obtained.


In addition to proper acquisition of specimens, data reliability requires precise and reproducible testing methods. Both manual and automated testing of hematologic specimens must be interpreted in light of test precision. This becomes especially important when evaluating the significance of small changes. All laboratory tests are evaluated with respect to accuracy and reproducibility. Accuracy is the difference between the measured value and the true value, which implies that a true value is known. Clearly this may present difficulties when dealing with biologic specimens. The National Committee for Clinical Laboratory Standards (NCCLS) and the International Committee for Standards in Haematology (ICSH) have attempted to develop standards to assess the accuracy of hematologic examination (8) and automated blood cell analyzers (9). Automated instrumentation requires regular quality assurance evaluation and calibration to reach expected performance goals (10, 11 and 12).

Reproducibility is the precision of a value and reflects the variability between repetitive determinations as measured on the same specimen. Most test results show a scattering of data points around a mean in a Gaussian distribution. Therefore, reproducibility may be expressed as the standard deviation (SD). If one wishes to compare results of tests that are measured in different units, they may be compared by the coefficient of variation (CV), which is derived from the following formula:

in which CV is expressed as a unitless percentage, SD is the standard deviation, and is the mean of counted values. This allows two independent assays, which have different units of measurement, to be compared (that is, red blood cell count in cells per liter and hemoglobin concentration in grams per liter).


Cell counts remain the basis for many of the parameters used in evaluating the blood. Cell counts may be determined either manually or by automated hematology analyzers. Whether they are performed by manual or automated means, accuracy and precision of the counts depends on proper dilution of the blood sample and precise sample measurement. The blood must be precisely aliquoted and the cells evenly distributed within the sample. Because blood contains large numbers of cells, sample dilution is required for accurate analysis. The type of diluent added depends on the cell type counted. Thus, red cell counts require dilution with an isotonic medium, whereas in white cell or platelet counts a diluent that lyses the more numerous red cells is often used. The extent of dilution also depends on the cell type. In general, red cell counts need more dilution than is required for the less abundant white blood cells. Errors in cell counts are caused primarily by errors in sample measurement, dilution, and enumeration of cells. The highest degree of precision occurs when a very large number of cells is enumerated. Clearly, automated methods provide the best means for counting large numbers of cells and minimizing statistical error.

Manual counts are carried out after appropriate dilution of the sample in a hemocytometer, a specially constructed counting chamber that contains a specific volume. Cells may then be counted with a microscope. Red blood cells, leukocytes, and platelets may be counted by this method (13). Due to the inherent imprecision of manual counts and the amount of technical time required, most cell counting is now performed by automated or semiautomated instruments. These machines increase the accuracy and speed of analysis by the clinical laboratory, particularly as test entry, sampling, and analysis are incorporated into single systems (14). With increasing levels of automation, some hematology analyzers have now moved to complete automation, which can be coupled with other laboratory tests using the same tube of blood.

Automated methods for cell counts use several types of technology, each with unique features (15). Two major types of automated cell counters are available: those that depend on changes in impedance in electrical flow (16) and those that use differences in light scatter properties (17). Most automated hematology analyzers also perform other measurements, such as hemoglobin concentration, red cell size, and leukocyte differentials. Newer instruments may also perform more specialized testing, such as reticulocyte counts (18). Table 2.1 lists the comparable values of reproducibility for automated and manual (hemacytometer) counting methods.

Table 2.1. Reproducibility of Blood Counting Procedures

Aperture-Impedance Counters

This type of analyzer, which includes the Coulter (Hialeah, FL), the Sysmex (Baxter Diagnostics, Waukegan, IL), and some Cell-Dyn (Abbott Diagnostics, Santa Clara, CA) instruments, enumerates cells in a small aperture by measuring changes in electrical resistance as the cell passes through the orifice (Fig. 2.1). A constant current passes between two platinum electrodes on either side of the orifice. The diluent that suspends the cells is more electrically conductive than are the cells. Hence, as each cell passes through the orifice there is a momentary decrease in electrical conductance so that an electrical impulse is generated and recorded electronically. The drop in voltage is proportional to cell size, allowing average cell size to be determined simultaneously (19, 20 and 21).

Instruments using aperture-impedance technology require even cell suspensions so that cells pass individually through the electrical current. Distortion of the electrical pulses may occur when the cells do not pass through the center of the aperture or when more than one cell enters the aperture at a time. The data may be electronically adjusted to exclude distorted peaks, and both upper and lower limits of particle size can be set to exclude cellular clumps or debris. Using size limitation parameters, the instrument can be used to count particles of different sizes, thereby allowing different blood elements to be counted (22).

The Coulter type counters are probably the most widely used example of hematology analyzers that use electrical impedance methods. Most models print data in numerical form as well as providing histograms of blood element size (Fig. 2.2), and the newer models include a white cell differential in addition to red cell counts, white cell counts, platelet counts, reticulocyte counts, hemoglobin, hematocrit, mean corpuscular volume (MCV), mean corpuscular hemoglobin (MCH), mean corpuscular hemoglobin concentration (MCHC), red cell distribution width (RDW), and mean platelet volume (MPV). This type of instrumentation fully analyzes up to 109 samples per hour, depending on the model used, and flags abnormal red cell and white cell populations, including blasts and atypical cells.

Optical Method Counters

The other method used by some hematology analyzers depends on the light scatter properties of blood cells (17, 23 and 24). The primary instruments that use this technology include the Technicon series (H6000, H*1, H*2, H*3) (Bayer Diagnostics, Kent, WA) and the Cell-Dyn 3500 (Abbott Diagnostics, Santa Clara, CA). In these systems, diluted blood passes through a flow cell detector placed in the path of a narrowly focused beam of light (usually a laser) (Fig. 2.3). When the blood cells pass through the counting chamber, they interrupt or alter the beam of light, thereby generating an electrical impulse that may be recorded. The pattern of light scattering using different angles of detection may also be used to determine cell size, volume, shape, and complexity (17 and 24). Optical systems count red cells, white cells, and platelets with precision equivalent to that observed in electrical impedance methods (25). The Cell-Dyn 3500 can provide complete blood counts and white cell differential on a maximum of 90 samples per hour, and the Technicon® H*3 RTC can process up to 102 samples per hour. Depending on the model of analyzer used, deviations in red cell size, white cell left shift, atypical lymphocytes, blasts, and immature granulocytes are flagged.

FIG. 2.1 Impedance type of automated hematology analyzer. As the cells pass through the aperture, they alter the current flow between the electrodes, generating an electronic pulse. Each pulse is recorded electronically. The magnitude of the pulse is proportional to the cell's volume.

FIG. 2.2 Histograms and printout generated by the Coulter STKR automated hematology analyzer.

FIG. 2.3 Optical type of automated hematology analyzer. A suspension of cells is passed through a flow chamber and focused into a single cell sample stream. The cells pass through a chamber and interact with a laser light beam. The scatter of the laser light beam at different angles is recorded, generating signals that are converted to electronic signals giving information about cell size, structure, internal structure, and granularity. (Adapted from Cell-Dyn 3500 Operator's Manual, Abbott Diagnostics, Santa Clara, CA 1993.)


Red blood cells are defined by three quantitative values: the volume of packed red cells (VPRC), or hematocrit; the hemoglobin concentration (Hb); and the red cell concentration per unit volume (RBC). Three additional indices describing average qualitative characteristics of the red cell population are also collected. These include the MCV, the MCH, and the MCHC. All of these values are normally collected and calculated by automated counters, and this method has largely replaced many of the previously used manual or semiautomated methods of red blood cell measurement, with certain exceptions as noted below.

Volume of Packed Red Cells (Hematocrit)

The volume of packed red cells (VPRC), or hematocrit, is the proportion of the volume of a blood sample that is occupied by red blood cells. The hematocrit may be determined manually by centrifugation of blood at a given speed and time in a standardized glass tube with a uniform bore, as was originally described by Wintrobe (26). The height of the column of red cells compared with that of the total column of blood following centrifugation yields the hematocrit. Both macro (using 3-mm test tubes) methods with low-speed centrifugation or micro methods using capillary tubes and high-speed centrifugation may be used.

The manual method of measuring hematocrit has proved to be an accurate method of assessing red cell status. It is easily performed with little specialized equipment, allowing it to be adapted for situations where automated cell analysis is not readily available, or for office use. However, several sources of error are inherent in the technique. The spun hematocrit measures the red cell concentration, not red cell mass. Therefore, patients in shock or with volume depletion may have normal or high hematocrit measurements due to hemoconcentration, despite a decreased red cell mass. Technical sources of error in manual hematocrit determinations usually arise from inappropriate concentrations of anticoagulants (27), poor mixing of samples, or insufficient centrifugation. Another inherent error in manual hematocrit determinations arises from trapping of plasma in the red cell column. This may account for 1 to 3% of the volume in microcapillary tube methods, with macrotube methods trapping more plasma (28 and 29). In addition, it should be noted that abnormal red cells (such as sickle cells, microcytic cells, macrocytic cells, or spherocytes) may trap higher levels of plasma because of increased cellular rigidity, possibly accounting for up to 6% of the red cell volume (28). Very high hematocrits, as in polycythemia, may also have excess plasma trapping. Manual hematocrit methods typically have a precision of about 2% (CV) (28).

Automated determination of hematocrit usually does not depend on centrifugation techniques, but instead measures or calculates hematocrit dependent on direct measurements of red cell number and red cell volume (hematocrit = red cell number × red cell volume). The automated values closely parallel manually obtained hematocrit values, and the manual hematocrit method is used as the reference method for automated methods (with correction for the error induced by plasma trapping) (30). It should be noted that errors of automated methods are more common in patients with polycythemia (31) or abnormal plasma osmotic pressures (32). Manual methods of hematocrit determination may be preferable in these cases. The precision of most automated determinations of hematocrit is about 1% (CV) (25).

Hemoglobin Concentration

Hemoglobin is an intensely colored protein, which allows its measurement by a variety of colorimetric and spectrophotometric techniques. Hemoglobin is found in the blood in a variety of forms including oxyhemoglobin, carboxyhemoglobin, methemoglobin, and other minor components. These may be converted to a single stable compound, cyanmethemoglobin, by mixing blood with Drabkin solution, which contains potassium ferricyanide and potassium cyanide (33 and 34). Sulfhemoglobin is not converted, but is rarely present in significant amounts. The absorbance of the cyanhemoglobin is measured in a spectrophotometer at 540 nm and hemoglobin concentration determined. Similar methods are used in manual methods and automated cell analyzers. Hemoglobin concentration is expressed in grams per deciliter (g/dL) of whole blood. The main errors in measurement arise from dilution errors or increased sample turbidity due to improperly lysed red cells, leukocytosis, or increased levels of lipid or protein in the plasma (35, 36, 37 and 38). Using automated methods, the precision for hemoglobin determinations is less than 1% (CV) (25).

Red Cell Counts

Manual methods for counting red cells have proven to be very inaccurate, and automated counters provide a much more accurate reflection of red cell numbers (25). Both erythrocytes and leukocytes are counted in whole blood that has been diluted in an isotonic medium. As the number of red cells greatly exceeds the number of white cells (by a factor of 500 or more), the error introduced by counting both cell types is negligible. However, in patients with marked leukocytosis, red cell counts and volume determinations may be erroneous unless corrected for white cell effects. The observed precision for red cell counts using automated hematology analyzers is less than 1% (CV) (25), compared with a minimal estimated value of 11% using manual methods (26).

Mean Corpuscular Volume

The average volume of the red blood cells is a useful red cell index that is used in classification of anemias (Chapter 31). The MCV is usually measured directly with automated instruments, but may also be calculated from the erythrocyte count and the hematocrit by means of the following formula (26):

The MCV is measured in femtoliters (fL, or 10–15 L). Using automated methods this value is derived by dividing the summation of the red cell volumes by the erythrocyte count. The CV in most automated systems is approximately 1% (25).

Agglutination of red blood cells, as in cold agglutinin disease, may result in a falsely elevated MCV (39). Most automated systems gate out MCVs above 360 fL, thereby excluding most red cell clumps, although this may falsely lower hematocrit determinations. In addition, severe hyperglycemia (glucose greater than 600 mg/dL) may cause osmotic swelling of the red cell during analysis, leading to a falsely elevated MCV (32 and 40). The CV for automated MCV measurements is less than 1%, compared with approximately 10% for manual methods (29).

Mean Corpuscular Hemoglobin

MCH is a measure of the hemoglobin content per red cell. It may be calculated manually or by automated methods using the following formula (26):

MCH is expressed in picograms (pg, or 10–12 g). Thus, the MCH reflects the mass of hemoglobin. In anemias where hemoglobin synthesis is impaired, hemoglobin mass per red cell decreases, leading to a decrease in the MCH. MCH measurements may be falsely elevated by hyperlipidemia (35 and 36) or leukocytosis (36), because increased plasma turbidity may erroneously elevate the hemoglobin measurement. Centrifugation of the blood sample to eliminate the turbidity followed by manual hemoglobin determination can allow correction of the MCH value. The CV for automated analysis of MCH ranges from 0.6 to 1.2 (32), compared with approximately 10% for manual methods (29).

Mean Corpuscular Hemoglobin Concentration

The average concentration of hemoglobin per red cell may be calculated by the following formula (26):

The MCHC is expressed in grams of hemoglobin per deciliter of packed red blood cells. This measures the concentration of hemoglobin (that is, the ratio of hemoglobin mass to the volume in which it is contained). Thus, if decreased synthesis of hemoglobin exceeds the decrease in red cell size in certain microcytic anemias, the MCHC will be decreased. The determination of MCHC is affected by conditions that affect either hematocrit (plasma trapping or presence of abnormal red cells) or hemoglobin (hyperlipidemia, leukocytosis) determinations. The CV for MCHC for automated methods ranges between 1 and 1.5 (32).

With the exception of hereditary spherocytosis and some cases of homozygous sickle cell or hemoglobin C disease, MCHC values will not exceed 37 g/dL. This level is close to the solubility value for hemoglobin, and further increases in hemoglobin concentration may lead to crystallization.

As noted above, the MCV, MCH, and MCHC reflect average values and may not adequately describe blood samples when mixed population of cells are present. For example, in sideroblastic anemias a dimorphic red cell population of both hypochromic and normochromic cells may be present, yet the indices may be normochromic. Thus, it is important to examine the blood smear as well as red cell histograms in order to detect the bimodal populations. The MCV is an extremely useful value in classification of anemias, but the MCH and MCHC often do not add significant clinical information. However, the MCH and MCHC play an important role in laboratory quality control because these values will remain stable for a given specimen.

Red Cell Distribution Width

The RDW is a newer red cell measurement that, along with a histogram of red cell volume heterogeneity, is provided by many of the more recent automated hematology analyzers (41, 42 and 43). The measurement reflects the range of red cell sizes measured within a sample. RDW has been proposed to be useful in early classification of anemias because it becomes abnormal earlier in nutritional deficiency anemias than any of the other red cell parameters, especially in cases of iron deficiency anemia (41,44 and 45). Thus, the RDW may be useful when characterizing microcytic anemia, particularly distinguishing between iron deficiency anemia (high RDW, normal to low MCV) and uncomplicated heterozygous thalassemia (normal RDW, low MCV) (41,46 and 47). However, the reliability of using RDW as a sole method for diagnosis of anemia is uncertain, and other methods are required to confirm the diagnosis (48).

In addition to providing information about the etiology of an anemia, the RDW is useful in identifying red cell fragmentation, agglutination, or dimorphic cell populations (46 and 49). Blood from patients with cold agglutinin disease yields increased MCV and RDW values, and the red cell histogram shows a bimodal population of cells: one population of cells in the normal size range and one population with an apparent size approximately double that of the single cells (50). Bimodal distributions of cell size may also be seen in patients who have had transfusions or have been recently been treated for a nutritional deficiency.

Complementing the multiparameter analyses performed by the instruments outlined above, several instruments have been developed that allow patient bedside point-of-care analysis for limited hematologic parameters, including hemoglobin concentration and hematocrit. Initial evaluation of this technology shows good levels of accuracy, although limitations exist in the complexity of the hematological analysis performed (51 and 52).


White Blood Cell Counts

Leukocytes are enumerated by either manual methods or automated hematology analyzers. Leukocytes are counted following dilution of blood in a diluent that lyses the red blood cells (usually acid or detergent). The much lower numbers of leukocytes present require less dilution of the blood than is needed for red blood cell counts. Manual counts, as with red cell counts, have more inherent error, with coefficients of variation ranging from 6.5% in cases with normal or increased white cell counts to 15% in cases with decreased white cell counts. Automated methods characteristically yield coefficients of variation in the 1 to 3% range (25). Leukocyte counts may be falsely elevated in the presence of cryoglobulins or cryofibrinogen (53), aggregated platelets (54), and nucleated red blood cells, or when there is incomplete lysis of red cells. Falsely low neutrophil counts have also been reported due to granulocyte agglutination secondary to surface immunoglobulin interactions (55).

Leukocyte Differentials

Following the white blood cell number, the white cells are analyzed to find the percentage of each white blood cell type by doing a differential leukocyte count. Manual methods for performing differential leukocyte counts have been proposed by the NCCLS (56). In manual leukocyte counts, three main sources of error are encountered: distribution of cells on the slide, cell recognition errors, and statistical sampling errors (57 and 58). In wedge-pushed smears, leukocytes tend to aggregate in the feathered edge and side of the blood smear rather than in the center of the slide. Larger cells (blasts, monocytes) also tend to aggregate at the edges of the blood smear (59). Use of coverslip preparations and spinner systems tends to minimize this artifact of cell distribution. For wedge-push smears, it is recommended that a battlement pattern of smear scanning be used in which one counts fields in one direction, then changes direction and counts an equal number of fields before changing direction again to minimize distributional errors (13). However, it should be noted that scanning the entire blood film, especially at the edges, is extremely important so that abnormal cell populations are not missed. Poor preparation and staining of peripheral smears are major contributors to cell recognition errors (59). Statistical errors are the main source of error inherent in manual counts, due to the small sample size in counts of 100 or 200 cells (60). Automated methods of differential counting tend to be more accurate because of the much larger sample of cells counted (61, 62 , 63 and 64).

Automated methods of obtaining a leukocyte differential have developed that markedly decrease the time and cost of performing routine examinations. However, this technology is incapable of identifying and classifying all abnormal or immature cells. Therefore, most machines identify abnormal white cell populations by flagging, indicating the need for examination by a skilled morphologist (64). The automated instruments used for performing automated leukocyte differentials are of two general types: cell identification on the basis of pattern recognition using stained slides and automated microscopy; and a flow-through system that identifies cells on the basis of cell size and staining characteristics.

Pattern recognition systems were first available in the early 1970s and included such instruments as the Hematrack (Geometric Data); Coulter diff 3 and diff 4, Abbott® ADC 500, and the Leukocyte Automatic Recognition Counter (Corning Laboratories) (65 and 66). This technology used a blood film on a glass slide that was stained and loaded onto the machine. A computer drove a microscopic mechanical stage until a dark staining area, corresponding to a leukocyte nucleus, was detected. Using data collected for each cell on cell size, nuclear and cytoplasmic coloration, and density, the computer matched the data patterns with specifications for each white cell type and identified the cell. Most pattern recognition technology was hampered by many of the same limitations of accuracy–limited numbers of cells counted, difficulties in classifying abnormal cell types, and cell distribution characteristics–as are manual counts (67). Although the automated pattern recognition systems do decrease technician time, they are significantly slower than the flow-through methods. Hence, pattern recognition systems are now rarely used, and the instruments are no longer manufactured.

Because of the ability to link the automated differential to the rest of the automated hematologic analysis, most recent systems use a flow-through system that generates a leukocyte differential as a part of the complete blood count (CBC) (63, 68 and 69). Flow-through systems collect and analyze data from large numbers of white blood cells to provide a differential count that has a high degree of precision. The determination of white blood cell type depends on cell size and  cytochemical  staining  characteristics  (Technicon® H6000, H*1, H*2, H*3 series) (25, 70, 71 and 72) or on the basis of cell volume and internal complexity, as measured by electrical impedance and light scatter characteristics (Coulter STKR and S-Plus series [25 and 73], TOA E5000 [42], Cell-Dyn 3000 [25], Sysmex NE-8000 [25, 74 and 75], and Cobas-Helios [76] systems).

Systems that use myeloperoxidase staining characteristics of cells perform cell counts on specimens via continuous-flow cytometric analysis of blood samples in which the red cells have been lysed and white cells fixed. The cells are suspended in diluent and passed through a flow cell in a continuous stream so that single cells are analyzed for cell size (dark field light scatter) and cytochemical characteristics of myeloperoxidase staining (bright field detector). The data are plotted as a scattergram reflecting cell size (light scatter) on the y axis and myeloperoxidase staining intensity or activity on the x axis (Fig. 2.4), which gives rise to a six-part differential (neutrophils, lymphocytes, monocytes, eosinophils, basophils, and large unstained cells).

FIG. 2.4. Histograms and printout generated by the H*1 automated hematology analyzer.

The total white blood cell count as well as the neutrophil, lymphocyte, monocyte, and eosinophil counts are enumerated in the myeloperoxidase channel. Lymphocytes are characterized as small (low-scatter) unstained cells. Larger atypical lymphocytes, blasts, or circulating plasma cells fall into the large unstained cells channel. Neutrophils have stronger peroxidase staining and appear as larger cells. Eosinophils have very strong peroxidase activity, but appear smaller than neutrophils because they tend to absorb some of their own light scatter. Monocytes have lower levels of peroxidase activity and are usually found between neutrophils and the large unstained cell areas. The system uses floating myeloperoxidase staining thresholds to bracket the neutrophil area, which helps adjust for individual sample differences in myeloperoxidase staining. To enumerate basophils, which are difficult to enumerate with automated flow-through techniques, the later models (Technicon H*1, H*2, and H*3) use a basophil-nuclear lobularity channel. For this determination, red blood cells and white blood cells are differentially lysed, leaving bare leukocyte nuclei, with the exception of basophils, which are resistant to lysis and can then be counted based on cell size. Light scatter data obtained from the leukocyte nuclei may also help identify blasts, which have a lower light scatter than do mature lymphocyte nuclei. The nuclear lobularity index is a measurement of the number of mononuclear and polynuclear cells that may help identify immature neutrophils or nucleated red blood cells when correlated with mean peroxidase activity and cell count data. These abnormal cell populations generate a flag, indicating a need for morphologic review of the peripheral smear. Studies using these systems have shown good ability to identify acute leukemias (77, 78 and 79), myelodysplastic syndromes (80), and acute infection or inflammation (72 and 81). Analysis using this technique examines thousands of cells per sample, increasing statistical accuracy (25, 57 and 62). The H*3 analyzers may perform 60 or more leukocyte differentials per hour.

The remaining systems use leukocyte volume determinations based on electrical impedance, in some cases coupled with light scatter data to generate a leukocyte differential. Initially, this type of methodology gave rise to a three-part differential that enumerated only neutrophils, monocytes, and lymphocytes exemplified by the Coulter S-Plus series of analyzers. This count was based on white cells that had been lysed, with subsequent collapse of the cellular cytoplasm around the nucleus and cytoplasmic granules (73 and 82). The cells were divided into three distinct size populations: large cells (neutrophils), intermediate cells (monocytes), and small cells (lymphocytes). When clear-cut size populations were not discernible, the machine generated a flag to indicate that the peripheral smear needed to be reviewed. This type of technology is best at enumerating neutrophils and lymphocytes, with high levels of correlation between manual and instrument determinations (67). However, there was a poor correlation on monocytic counts because of lower cell numbers. In addition,  other  cell populations  including eosinophils, basophils, atypical lymphocytes, blasts, immature granulocytes, and plasma cells tended to fall into the monocytic region or granulocyte region and confounded the data. Depending on the patient population studied (that is, the percentage of normal versus abnormal samples) the proportion of false negatives (samples in which a true abnormal population was not detected by the analyzer) varied from 4 to 16% (67, 82, 83, 84 and 85). This value is similar to those of the manual methods, where the false negative rate is estimated to be 9% (67). Although the three-part differential was useful as a screening tool, its limitations in providing full analysis of the white blood cell component was of concern.

More recently, most impedance-type hematology analyzers incorporate improved methods of identifying leukocyte subsets using size data enhanced by additional data of internal cell complexity provided by conductivity or light scatter measurements. This modification has greatly improved the ability of later model analyzers to provide a full differential white blood cell counts. Three of the most commonly used hematology analyzers of this later generation include the Coulter STKS, the Sysmex NE-8000, and the Cell-Dyn 3000 or 3500, although new upgrades and models appear with great rapidity (86).

The Coulter STKS uses electronic impedance to measure volume, high-frequency electromagnetic fields to measure conductivity and light scatter with a monochromatic laser to determine cell cytoplasmic complexity or granule content, analyzing up to 144 specimens per hour. These generate a three-dimensional scatter plot (Fig. 2.2) that can separate the leukocytes into neutrophils, lymphocytes, monocytes, eosinophils, and basophils with flags for abnormal populations. The Sysmex NE-8000 uses electrical impedance and electromagnetic data to identify the monocytes, neutrophils, and lymphocytes, then identifies eosinophils and basophils based on a proprietary lysing agent (87 and 88). It may analyze up to 120 samples per hour. The Cell-Dyn 3000 identifies all of the leukocyte classes based on light scatter properties (small-angle forward light scatter, wide-angle light scatter, orthogonal light scatter, and depolarized light scatter [89]). The Cell-Dyn 3500 uses both impedance and laser light scatter at 0°, 10°, and 90° angles (90 and 91). When compared among themselves and with the Technicon H*1 or H*2, all of the automated hematology analyzers mentioned above had excellent accuracy and precision for typical clinical laboratory usage, with slight differences between the different technologies but a marked improvement over manual methods. Most studies find a poor correlation value for basophil counts (25, 62 and 91), probably reflecting the very low levels of these cells available for manual counts. A newer instrument developed in Europe, the Cobas analyzer (Roche Diagnostic Systems, Branchburg, NJ), uses a flow cytometric and light scatter technology that allows somewhat improved detection of band neutrophils over other systems with similar accuracy and precision with regard to other white and red blood cell parameters (92, 93, 94 and 95).

In addition to their utility in providing a differential count of white blood cells, the flow-through techniques of automated cell counting also can provide reproducible and accurate absolute numbers of each cell type because they analyze large populations of cells. Use of percentages of cell types may mask some cytopenias or excessive numbers of cells. Absolute counts are used to define some disease states, such as chronic lymphocytic leukemia and chronic myelomonocytic leukemia. Absolute neutrophil counts are often useful when monitoring bone marrow recovery after chemotherapy or bone marrow transplant.


Platelets are counted in automated hematology analyzers once red cells have been removed by sedimentation or centrifugation, as well as by techniques using whole blood (96 and 97). These give highly reliable platelet numbers when compared with manual methods of counting using a hemacytometer (98). Falsely low platelet counts may be caused by the presence of platelet clumps, platelet agglutinins (54), or adsorption of platelets to leukocytes (99). Fragments of red or white blood cells may falsely elevate the automated platelet count, but this usually gives rise to an abnormal histogram that identifies the spurious result (100 and 101).

Automated hematology analyzers also determine mean platelet volume (MPV), which has been correlated with several disease states (102 and 103). In general, MPV has an inverse relationship with platelet number, with larger platelet volumes seen in thrombocytopenic patients (104 and 105). MPV is characteristically increased in hyperthyroidism (106) and myeloproliferative disorders (107). However, it should be noted that platelets tend to swell during the first 2 hours in EDTA anticoagulant, shrinking again with longer storage (108 and 109). Decreased MPV has been associated with megakaryocytic hypoplasia and cytotoxic drug therapy (110).


Clearly the use of automated hematology analyzers has reduced laboratory costs and turnaround time while improving the accuracy and reproducibility of blood counts. The coefficient of variation for most of the parameters measured is in the range of 1 to 2%. This degree of reproducibility is not achievable with the use of most manual techniques (Tables 2.1 and 2.2).

Despite this high degree of accuracy, several potential errors may invalidate automated collection of data. Proper calibration of instrumentation is essential for collection of accurate data. Faulty current settings, which determine threshold counting values as well as variation in either the counting volumes or flow characteristics of a sample, will negatively affect data accuracy. Electrical or mechanical failures induce marked errors in data collection, as well as relatively minor voltage fluctuations. Careful calibration of the instrumentation initially, followed by frequent evaluation of reproducibility by analysis of samples with known cell concentrations, is an essential quality control measure (13 and 58). Reference methods for instrument calibration have been developed by both the NCCLS and ICSH (9, 22, 43, 56 and 111).

Table 2.2. Reproducbility of Red Cell Indices

Certain disease states are also associated with spuriously high or low results, although some of these are specific to a particular type of instrumentation (summarized in Table 2.3). Therefore, the individual values obtained from the automated hematology analyzer must be interpreted in context with the clinical findings. In addition, careful examination of the stained blood film often imparts additional information that may not be reflected in the average values that constitute the automated data. For example, decreased red blood cell counts, macrocytosis, and extremely high MCHC have been observed in patients with cold agglutinin disease with a higher thermal amplitude and in some patients with elevated serum viscosity (50). High levels of paraprotein may lead to falsely elevated hemoglobin levels, therefore affecting MCH and MCHC calculations (37). Many analyzers report spurious increases in hemoglobin levels when white cell counts exceed 30 ×109/L, due to increased turbidity. This has been addressed in the Sysmex systems by use of two lysing agents and redesign of the flow system, so that hemoglobin levels remain extremely accurate in the face of white blood cell counts as high as 100 × 109/L (75 and 112). Extremely high white cell counts may also may falsely raise the red cell count and hematocrit as the white cell count is incorporated into the red cell count. High glucose levels (greater than 400 to 600 mg/dL) and the associated hyperosmolarity cause red cell swelling and generate a high MCV and hematocrit with a falsely low MCHC (32, 40 and 113). The increased turbidity associated with hyperlipidemia may also cause falsely elevated hemoglobin determinations, MCH, and MCHC (35 and 36).

Despite the high level of accuracy and precision, the automated hematology analyzers usually have false positive rates (flagging) of 10 to 25% of patients, requiring manual examination of the blood smear (63 and 114). Blood smear examination still plays an important role in characterizing samples that raise flags or show findings outside the parameters set in a particular laboratory. In addition, some cells require morphologic examination to identify, such as Sézary cells (114), and red cell morphology is best analyzed by direct smear examination (68).


Careful evaluation of a well-prepared blood smear is an important part of the evaluation of hematologic disease. Although a specific diagnosis may be suggested by the data obtained from an automated hematology analyzer, many diseases may have normal blood counts but abnormal cellular morphology. Examples of abnormal red cells that may be seen in the peripheral blood smear examination and are associated with specific disease states are found in Table 2.4. However, morphologic analysis may be greatly hampered by poorly prepared or stained blood smears. Preparation of satisfactory blood smears requires careful attention to preparation of the blood smear, staining techniques, and familiarity with the morphologic appearances of normal and pathologic cell types.

Table 2.3. Disorders and Conditions That May Reduce the Accuracy of Blood Cell Counting*

Preparation of Blood Smears

Blood films may be prepared on either glass slides or coverslips. Each method has specific advantages and disadvantages. Blood smears are often prepared from samples of anticoagulated blood remaining from automated hematological analysis. However, artifacts in cell appearance and staining may be induced by the anticoagulant. Optimal morphology and staining are obtained from noncoagulated blood, most often from a fingerstick procedure. Mechanical dragging of the cells across the glass of the slide or coverslip also distorts the cells; however, this artifact may be minimized with proper technique.

Coverslip smears (Fig. 2.5A) are prepared using a good grade of flat, number 1 1/2-inch square (or 22 × 22 mm) coverslips that are free of lint, dust, and grease. Such coverslips allow optimal spreading of the blood over the surface and minimal artifact. Usually, high-quality coverslips do not require additional cleaning, although there may be some deterioration with age. Plastic "nonwettable" coverslips are not satisfactory for these preparations. The smear is prepared by holding the coverslip by two adjacent corners between the thumb and index finger. A small drop of either fresh or anticoagulated blood is placed in the center of the coverslip. The size of the drop of blood is critical. If the drop is too large, a thick smear results. If the drop of blood is too small, a very thin smear is obtained. A second coverslip is then grasped in a similar fashion with the other hand and placed across the first coverslip and rotated 45 degrees with a steady, rapid and gentle motion. The two coverslips are then immediately pulled apart and allowed to air dry. If done properly, this procedure produces two coverslips with even dispersion of blood without holes or thick areas.

Blood smears may also be prepared on clean glass slides by the wedge method (Fig. 2.5B). This often leads to irregular distribution of cells on the slide, a distinct disadvantage over the coverslip procedure. However, glass slides are less fragile and easier to handle, and may be labeled more easily than coverslips. To prepare a slide blood smear, a drop of blood is placed in the middle of the slide about 1 to 2 cm from one end. A second spreading slide is placed at a 30 to 45o angle and moved backward to make contact with the blood drop. The blood drop will spread along the slide edge, then the spreader slide is moved rapidly forward. This technique creates a film of blood that is 3 to 4 cm long. Artifact may be introduced by irregular edges in the spreader and by the speed at which the spreader is moved. Glass slide preparations have increased incidence of accumulation of the larger white cells at the edges of the film, increasing cellular distribution errors. Fast movement of the spreader results in a more uniformly distributed population of cells.

Automated techniques for blood smear preparation have also been developed that produce very uniform blood smears. Two major types of instruments are used: those that use centrifugation and those that mechanically spread the blood. Centrifugation techniques are often most useful when a small number of cells must be concentrated in a small area, as in preparing smears of cells in fluids such as cerebrospinal fluid. Mechanical spreaders mimic the manual technique, and are useful when large numbers of blood smears are prepared. In general, smears made by automated techniques are usually inferior to those made by an experienced technician.

Table 2.4. Pathologic Red Cells in Blood Smears

Routine Staining of Blood Smears

Blood smears are usually stained with either Wright or May–Grunwald–Giemsa stains. Both stains are modifications of the Romanowsky procedure (115). The stain may be purchased commercially, or may be made in the laboratory. The basic stain is formulated from methylene blue and eosin. The Wright stain formulation uses sodium bicarbonate to convert methylene blue to methylene azure, which stains the cell. Giesma stains use known quantities of acid bichromate to form the converted azure compounds. All types of Romanowsky stains are water insoluble but are dissolved in methyl alcohol. The stain must be free of water, which induces red blood cell artifacts. Water artifacts may be avoided by fixation of slides or coverslips in anhydrous methanol before staining.

Optimal staining conditions must be established for each new batch of stain. The methylene blue conversion to azure compounds continues to occur while the stain is in the bottle, so staining conditions may change over time. Methyl azures are basic dyes that impart a violet-blue coloration to the acidic components of the cell, such as nucleic acids and proteins. The eosin reacts with the basic cellular elements, imparting a reddish hue to cytoplasmic components and hemoglobin. A properly stained slide has a pink tint. The red cells have an orange to pink coloration and leukocytes have purplish-blue nuclei. The Romanowsky stains differentially stain leukocyte granules, which aids in morphologic analysis of the cells. Thus, neutrophil granules are slightly basic and stain weakly with the azurophilic component. The eosinophils contain a strongly basic spermine derivative and stain strongly with eosin. In contrast, basophil granules contain predominately acidic proteins and stain a deep blue-violet. No precipitate should overlie the cells, because this indicates use of slides or coverslips that were not cleaned properly. Dust on slides may also induce artifacts. Staining solutions should be filtered or replaced weekly if used heavily.

Occasionally an excessive blue coloration of the cells is seen. This may be caused by excessive staining times, improperly prepared or aged buffer that is too alkaline, old blood smears, or blood smears that are too thick. The quality of the staining may be improved by quick and vigorous rinsing with distilled water. If the areas between cells are staining, it usually indicates inadequate washing of the slide, heparin anticoagulation, or possible paraproteinemia. When the staining appears too pink or red, the usual problem is buffer that is too acidic. This results in pale-stained leukocyte nuclei, excessively orange-red blood cells, and bright red eosinophil granules. Other causes of excessive red coloration include inadequate staining times and excessive washing of the slide. Most often problems with staining are caused by problems with the acidity or alkalinity of the solutions, and new buffers often correct the problem.

FIG. 2.5. Preparation of blood smears. Blood smears may be prepared by the coverslip (A) or slide wedge method (B). Coverslip smears are prepared by placing a drop of blood in the center of a coverslip and spreading the blood by rotating a second coverslip over it. Wedge smears are prepared by placing a drop of blood on a slide and using a second slide to push the blood out along the length of the slide. (Adapted from Bauer JD. Clinical laboratory methods. 9th ed. St.Louis: CV Mosby, 1982; 270.)

Examination of the Blood Smear

The blood smear should be initially examined under an intermediate power (10 to 20× objective) to assess the adequacy of cellular distribution and staining. An estimate of the white blood cell count may also be made at this power, and scanning for abnormal cellular elements such as blasts or nucleated red blood cells can be performed. It is important to scan over the entire blood smear to ensure that abnormal populations, which may be concentrated at the edges of the smear, are not missed. Use of an oil immersion lens (50× or 63×) is usually sufficient for performing leukocyte differential counts, although a 100× oil lens may be necessary for study of cellular inclusions or cytoplasmic granules. Systematic evaluation of the blood smear is essential so that all cell types are examined and characterized. Each cell type should be evaluated for both quantitative and qualitative abnormalities.

It is difficult to evaluate quantitative abnormalities of red cells on a blood smear; however, the red blood cells should be evaluated for variations in size, shape, hemoglobin distribution, and the presence of cellular inclusions. The red cells are usually unevenly distributed throughout the blood film. Optimal red cell morphology is seen in an area of the smear where the red cells are close together but do not overlap. Areas where the red cells are spread too thinly or thickly have increased artifacts. In some blood smears the red cells appear to stick together, forming what appear to be stacks of red cells, called rouleaux. This phenomenon may be seen in areas of the smear where the red cells are too close together in normal blood, but if it persists in thinner areas of the blood film, it suggests the presence of a paraprotein coating the red cells and causing agglutination due to loss of normal electrostatic repulsion between red cells.

Red cells should be uniform in size and shape, with an average diameter of 7.2 to 7.9 µm. This may be measured by use of a micrometer or by using the diameter of a small lymphocyte nucleus for comparison. Variation in red cell size is called anisocytosis. Cells that are larger than 9 µm and well hemoglobinated are considered macrocytes. Less mature erythrocytes are macrocytic and have a bluish tint to the hemoglobin (polychromatophilia) or have fine basophilic stippling of the cell due to remnant RNA and ribosomes. Microcytes are cells with a diameter less than 6 µm.

Normal erythroid cells are round. Variations in shape are called poikilocytosis. The red cell should have a pale central area with a rim of red to orange hemoglobin. Hypochromia reflects poor hemoglobinization and results in a very thin rim of hemoglobin or an increased area of central pallor. Abnormal distribution of hemoglobin may result in formation of a cell with a central spot of hemoglobin surrounded by an area of pallor, called a target cell. Abnormal hemoglobins may also form crystals. Spherocytes and macrocytes lack an area of central pallor because of increased thickness of the cell. Red cells may also contain inclusions such as remnants of nuclear material (Howell–Jolly bodies), remnants of mitochondria or siderosomes (Pappenheimer bodies), or infectious agents (malarial parasites).

Platelet numbers and morphology are then evaluated. Platelets appear as small blue cytoplasmic fragments with red to purple granules. Platelets are usually 1 to 2 µm in diameter with wide variation in shape. Platelet numbers may be estimated from the blood film. Normal platelet counts should have several (5 to 15) platelets per oil immersion field or approximately 1 platelet for 10 to 20 red blood cells. It should be noted that platelets may aggregate if uncoagulated blood is used, and this may cause the spurious impression of a low platelet count.

Leukocyte morphology and distribution are analyzed last. The number of leukocytes may be estimated by scanning the blood film at an intermediate power. Abnormal distribution of larger cells should be excluded by examination of the edges of the blood film in particular (59). White cells seen at the edges of the blood smear may appear artifactually smaller (because of cellular shrinkage and poor spreading of the cell) or larger (because of cellular disruption). Care must be taken when making the smear, because cells, particularly neoplastic cells, may be disrupted by too much mechanical pressure. Optimal morphology of the leukocytes requires that blood smears be made promptly. Significant artifact begins to be observed in blood that has been held for several hours and includes cytoplasmic vacuolation, nuclear karyorrhexis, and cytoplasmic disruption.

The white blood cells normally seen in the peripheral smear include neutrophils, eosinophils, basophils, lymphocytes, and monocytes. The presence of immature myeloid cells (myelocytes, metamyelocytes, promyelocytes, and blasts) is distinctly abnormal. At least 100 cells should be identified and counted to yield a manual white blood cell differential (56 and 57). In addition to identifying relative populations of white cells by performing a differential count, the cells should be closely examined for morphologic abnormalities of the cytoplasm and nucleus. For example, infection or growth factor therapy often leads to increased prominence of the primary (azurophilic) granules in neutrophils, which is called toxic granulation (116). In contrast, many myelodysplastic disorders are characterized by hypogranularity of neutrophils in addition to abnormal nuclear segmentation. Cytoplasmic inclusions may be seen in some storage disorders or lysosomal disorders.

Other Means of Examining Blood

Occasionally it is necessary to examine fresh blood as a wet mount. Wet preparations are made by placing a drop of blood on a slide, covering the drop with a coverslip, and surrounding the coverslip with petroleum jelly or paraffin wax to seal the edges. If needed, the blood may be diluted with isotonic saline or in some cases it may be fixed with buffered glutaraldehyde for later examination. The blood may then be viewed with light or phase contrast microscopy. Wet mounts are used to detect sickling of red cells, spherocytes, and parasites within erythrocytes. Other organisms, such as spirochetes and trypanosomes, may be detected by their movement. Dark field illumination enhances the refractile qualities of leukocyte granules or malarial pigment (117). Phase contrast microscopy accentuates the fine cellular details of cells, especially cytoplasmic granules and intracellular inclusions (118). Platelets are well visualized with phase contrast, and this aids in performance of manual platelet counts.

Supravital staining is performed on living, motile cells, and helps avoid artifacts induced by smear preparation, fixation, and staining (119, 120 and 121). However, such preparations are not permanent, a distinct disadvantage. Supravital stains are often used to detect red cell inclusions. These include crystal violet staining that detects Heinz bodies, which are denatured hemoglobin inclusions that appear as irregularly shaped purple bodies within the red cell. Brilliant cresyl blue may be used to precipitate and stain unstable hemoglobins, such as hemoglobin Zurich and hemoglobin H (122).

The most commonly used supravital stain is new methylene blue or brilliant cresyl blue used for reticulocyte determinations. These stains allow visualization of the reticulin network of erythrocyte ribosomes in newly formed red blood cells (123). Reticulocyte counts are used in evaluation of new red cell production and are helpful in determining the hematopoietic activity of the bone marrow and marrow response to anemia (124 and 125). Reticulocytes are not identified positively on Wright-stained blood smears, although their presence is suggested by polychromatophilia of the red blood cells. Recently, many automated hematology analyzers have incorporated staining to detect reticulocytes. These automated procedures appear to perform reticulocyte counting with a higher degree of precision than can be achieved manually (18, 126, 127, 128, 129 and 130). The degree of ribosomal staining may also be quantitated to allow assessment of reticulocyte age (125). Automated reticulocyte counts may have increased errors in the presence of Heinz bodies (131) or Howell–Jolly bodies (132) in the red cells. Normal values for reticulocyte counts may be affected by the age and sex of the patient (133).


Diagnosis and management of many hematologic diseases depend on examination of the bone marrow. Bone marrow examination usually involves two separate but interrelated specimens. The first is a cytological preparation of bone marrow cells obtained by aspiration of the marrow and a smear of the cells, allowing excellent visualization of cell morphology and enumeration of the marrow cellular elements. The second specimen is a needle biopsy of the bone and associated marrow, which allows optimal evaluation of bone marrow cellularity, fibrosis, infections, or infiltrative diseases.

There are several indications for performing a bone marrow examination. These include further workup of hematologic abnormalities observed in the peripheral blood smear, evaluation of primary bone marrow tumors, staging for bone marrow involvement by metastatic tumors, assessment of infectious disease processes including fever of unknown origin, and evaluation of metabolic storage diseases. Before a bone marrow examination is carried out, clear diagnostic goals about the information to be obtained from the procedure should be defined. Before the procedure one should decide whether any special studies are needed, so all the necessary specimens may be collected and handled correctly. Clearly, the decision to perform a bone marrow examination as well as the choice of tests to be performed using the material should be made on an individualized basis.

Several sites may be used for bone marrow aspiration and biopsy (134). In part, the site chosen reflects the normal distribution of bone marrow with the age of the patient. At birth, hematopoietic marrow is found in all of the bones of the body. However, by early childhood, fat cells begin to replace the bone marrow hematopoietic cells in the extremities. In adults, hematopoiesis is limited to the axial skeleton and proximal portions of the extremities (135). Thus, younger children may have marrow examinations from the anterior medial tibial area, whereas adult marrow is best sampled from the sternum at the second intercostal space or from either the anterior or posterior iliac crest area. Sternal marrows do not allow a biopsy to be performed, and several possible complications, including hemorrhage and pericardial tamponade, may occur if the inner table of the sternum is penetrated by the needle at areas other than the second intercostal space. The sternal marrow space in an adult is only about 1 cm thick at the second intercostal space, so care must be taken to avoid penetrating the chest cavity. In contrast, little morbidity is associated with iliac crest aspiration and biopsy, and the posterior iliac crest is the most common site for bone marrow sampling. The anterior iliac crest may be used if previous radiation, surgery, or patient discomfort do not allow a posterior approach.

Bone Marrow Aspiration and Biopsy

Bone marrow is semifluid and easily aspirated through a needle. Many types of needles have been used for performing marrow aspiration. Most are 14 to 18 gauge, and many have a removable obturator, which prevents plugging of the needle before aspiration, and a stylet that may be used to express the bone marrow biopsy sample (Fig. 2.6). Some models, primarily used for sternal bone marrow aspiration procedures, have adjustable guards that limit the extent of needle penetration.

In most cases, marrow aspiration and biopsy may be carried out with little risk or patient discomfort, provided adequate local anesthesia is used. Apprehensive patients may be sedated before the procedure, but this is usually not necessary. The procedure is performed under sterile conditions. The skin at the site of the biopsy is shaved, if necessary, and cleaned with a disinfectant solution. The skin, subcutaneous tissues, and periosteum in the area of the biopsy are anesthetized with a local anesthetic, such as 1% lidocaine, using a 25-gauge needle. Care must be taken to fully anesthetize the periosteum where most of the bone pain fibers are located. After the anesthetic has taken effect, a small cut is made in the skin overlying the biopsy site and the marrow aspiration needle is inserted through the skin, subcutaneous tissues, and bone cortex with a slight rotating motion. Entrance of the needle into the bone marrow cavity should be sensed as a slight give or increase in the speed of needle advancement. The needle obturator is removed and the needle is attached to a 10- or 20-mL syringe. Aspiration of the marrow is achieved by rapid suctioning with the syringe so that 0.2 to 2 mL of bloody fluid is obtained. Aspiration may cause a very brief, sharp pain. If no pain is noted and no marrow is obtained, the needle may be rotated and suction applied again. If no marrow is obtained, another sampling site may be required.

The aspirated material is given to a technical assistant, who makes smears of the material and assesses the quality of the material by noting the presence of marrow spicules. The smears must be made quickly to avoid clotting in a manner similar to that described for blood smears. After several smears are made, the aspirate is allowed to clot for later fixation and processing by the histology laboratory. If additional material is needed for flow cytometry, cytogenetics, culture, or other special studies, additional aspirations may be performed by withdrawing the needle and repositioning it in a new site. Occasionally a portion of an anticoagulated marrow aspirate is spun down to obtain a buffy coat, thereby concentrating the cellular elements in a very hypocellular specimen. EDTA is the best anticoagulant to use because it introduces the least amount of morphologic artifact to the specimen (136 and 137). In some instances, no marrow can be aspirated (dry tap). In these cases it is essential to make smears from material at the tip of the needle and to also make touch preparations from the biopsy, as outlined below, to allow cytologic examination of the bone marrow elements (138 and 139).

Using the same skin incision, the bone marrow biopsy may be performed if the aspirate has been performed in the iliac crest area. A separate biopsy needle that is slightly larger than the needle used for aspiration may be used, or the same needle that was used for the bone marrow aspiration may be reused. Care must be taken to reposition the needle entry site away from the area where the aspiration was performed to avoid collection of a specimen with extensive artifact induced by the aspiration procedure. The biopsy needle may require more pressure to enter the bone because of the larger bore size. Once the needle is in place in the bone, the stylet may be inserted to give an approximation of the size of the bone core within the needle. The biopsy needle is rotated and gently rocked to free the biopsy from the surrounding bone and then advanced slightly further. The biopsy is then removed from the bone by withdrawing the needle. The biopsy is expressed from the needle by the stylet. Touch preparations of the bone biopsy should be made, particularly if no aspirate was obtained, to allow cytologic examination of the bone marrow elements. The bony core is then fixed and processed for histologic examination.

Once the biopsy is completed, manual pressure is applied to the site for several minutes to achieve hemostasis. The site is then bandaged and the patient instructed to remain recumbent so as to apply further pressure for about 60 minutes. If a patient is thrombocytopenic, pressure bandages should be applied and the site checked frequently for prolonged bleeding.

Staining and Evaluation of Bone Marrow Aspirates and Touch Preparations

The bone marrow aspirate or touch preparation slides are stained with either Wright or May-Grünwald–Giemsa stains, similar to the procedure for blood smears. These stains allow excellent morphologic detail and allow differential counts to be performed (134). Unstained smears should be retained for possible special stains, if indicated (140).

Evaluation of bone marrow aspirates gives little information about the total cellularity of the bone marrow because of fluctuations in cell counts induced by peripheral blood contamination of the bone marrow specimen (141). An overall impression of the cellularity may be given (that is cellular or paucicellular). More accurate evaluation of bone marrow cellularity requires examination of a bone marrow biopsy section, although the biopsy represents a tiny fraction of the total marrow and may also be subject to sampling error (142). The stained aspirate smear appears as a central zone of dark marrow particles surrounded by a thinner area of dispersed bone marrow cells and red cells. Low-power examination allows evaluation of the adequacy of cellularity and of the presence of megakaryocytes. Tumor cells or granulomas may also be seen by scanning the aspirate smear at low power.

The aspirate smear allows cytologic examination of the bone marrow cells. A minimum of 500 nucleated cells should be evaluated under oil immersion magnification. Only intact cells are evaluated; all bare nuclei are excluded. Counting is performed in an area where few bare nuclei are present and the cells are not overlapping, found in clusters, or artifactually distorted. Reference ranges for the percentage of bone marrow cell types vary widely between laboratories, and are used only as guides for what is to be expected in normal bone marrow samples (140 and 143). Results of differential counts from sternal bone marrow aspirate smears obtained from 12 healthy men at the University of Utah are presented in Table 2.5 as an example of bone marrow differential count reference ranges. The proportions of each cell type and maturational sequence are determined from the differential counts. In addition, the myeloid to erythroid ratio may be calculated.

Differences in cell differential results among infants, children, and adults exist (Table 2.6) (140). In general, lymphocytes are more commonly seen in the marrow of children, especially those younger than 4 years of age, where they may compose up to 40% of the marrow cellularity (144, 145 and 146). Plasma cells are rare in the marrow of infants. Lymphocytes are much less numerous in adult bone marrows, usually making up less than 20% of adult marrow cellularity. Lymphocyte and plasma cell counts in adults tend to be quite variable, perhaps reflecting the tendency of these cells to be unevenly distributed in the bone marrow of adults. Often lymphoid cells are found in nodular aggregates in older adults, and plasma cells tend to be associated with blood vessels (147 and 148).

FIG. 2.6. Jamshidi bone marrow aspiration and biopsy needle. This type of hollow needle with a beveled tip (A) is satisfactory for percutaneous biopsy of the bone marrow. The needle is inserted with the obturator (B) in place. The biopsy is expressed from the needle using the stylet (C).

Table 2.5. Differental Counts of Bone Marrow Aspirates from 12 Healthy Man

During the first month of life, erythroid cells are prominent because of high levels of erythropoietin (149 and 150); thereafter, the erythroid cells make up 10 to 40% of the marrow cells. Relatively few early erythroid precursors (normoblasts) are usually seen, and more mature forms predominate. Erythroid cells should be examined for abnormalities in morphology as well as iron content because these parameters are often deranged in pathologic states. The myeloid cells are usually the predominant cell within the bone marrow, and more mature cells are most numerous. Increased numbers of immature myeloid cells usually indicate a disease process. Children tend to have higher numbers of eosinophils and eosinophilic precursor cells than do adults, although many medications may increase the bone marrow eosinophil count. Megakaryocytes constitute the least abundant cell type seen in the bone marrow, usually making up less than 1% of the cells.

In addition to the hematopoietic cells mentioned above, a variety of other cells may be seen in bone marrow aspirates in varying proportions. These include macrophages, mast cells, stromal cells, osteoblasts, osteoclasts, and fat cells. Normally, these cells make up less than 1% of the total marrow cellularity; however, they may be increased in a variety of reactive and pathologic processes. Aspirate smears are excellent for evaluation of macrophage hemophagocytosis (151) or storage disorders. Osteoclasts and osteoblasts are most often seen in aspirates obtained from children in whom active bony remodeling is taking place, and are distinctly abnormal when seen in adult aspirates (140).

Examination of Bone Marrow Histologic Sections

Bone marrow core biopsies and the clot obtained from the aspiration procedure are usually fixed in formalin or in a coagulative fixative, such as B5. The material is then processed and embedded in paraffin or plastic, and sections are made for examination. Plastic embedding allows preparation of very thin (2 µm) sections that allow optimal morphologic assessment, but requires additional technical expertise and longer processing times (48 hours) (152 and 153). Well-prepared, thin (3- to 4-µm) sections from paraffin-embedded materials may also be used. Preparation of paraffin-embedded sections requires decalcification of the bone biopsy, which destroys many of the enzymes found in blood cells. Because plastic-embedded sections do not require decalcification, they are more likely to retain enzymatic activities, thereby allowing cytochemical staining to be performed on the section (154). The sections are stained with either hematoxylin and eosin or Giemsa stains for examination.

Bone marrow biopsies are useful in evaluation of the cellularity of the bone marrow sampled. Several caveats must be kept in mind when assessing cellularity. Studies show variations in cellularity even within the same biopsy site (142) as well as between different anatomic sites. However, comparisons of the relative proportions of myeloid, erythroid, and megakaryocytic cells appear to be constant even in widely separated biopsy sites (155 and 156). In older patients, the subcortical area is often hypocellular, and care must be taken to obtain a large enough biopsy to allow evaluation of the marrow away from this area (142). The bone marrow biopsy section provides the best representation of the bone marrow and its anatomic relationships. The clot section, which is prepared from the bone marrow aspirate material, has a degree of inherent artifact because the bone marrow is removed from its normal relationship with bone, blood vessels, and other stromal elements. In particular, cellularity estimations may be falsely elevated by collapse of the normal stromal network (156).

In addition to providing information about the anatomic distribution and relationships of hematopoietic cells, the bone marrow biopsy is useful for evaluation of focal infiltrative processes such as carcinoma, lymphoma, other tumors, granulomatous inflammation, and fibrosis (156, 157, 158 and 159). Occasionally the marrow is so involved with an infiltrative process that no aspiration can be obtained (dry tap) and the biopsy provides the only diagnostic material (138). In addition, evaluation of other elements, such as bony trabeculae, blood vessels, and stroma, requires a biopsy specimen.


Several special stains may be performed on peripheral blood smears, bone marrow aspirate smears, bone marrow touch preparations, and bone marrow biopsy materials, which provide additional information about the cell lineage beyond what is obtained by standard staining with Romanowsky or hematoxylin and eosin stains. Special stains generally fall into two categories: cytochemical stains that use enzymatic reactions by the cell to impart staining, and immunocytochemical stains that stain cell-specific antigen epitopes. These stains are particularly useful in characterization of primary hematologic or metastatic malignancies.

Table 2.6. Changes in Differential Counts of Bone Marrow with Age

Cytochemical Stains

Cytochemical stains are extremely useful in the diagnosis and classification of acute leukemias. They allow correct identification of myeloid and lymphoid acute leukemias (160), as well as providing the basis for subclassification of the acute myeloid leukemias by the French–American–British (FAB) criteria (161 and 162). Cytochemical stains are usually performed on peripheral blood films, bone marrow aspirates, or touch preparations made from bone marrow, lymph node, or other tissue biopsies. Best results are obtained by using freshly obtained materials; however, some reactions may be carried out on materials that are several years old. In some cases, nondecalcified, plastic-embedded bone marrow biopsies may also be used (154).

Myeloperoxidase Stain

Primary granules of neutrophils and secondary granules of eosinophils contain myeloperoxidase (163). Monocytic lysosomal granules are faintly positive (164). Lymphocytes and nucleated red blood cells lack the enzyme. Originally the stain depended on oxidation of benzidine by hydrogen peroxide (165); however, because benzidine is a potential carcinogen, alternative substrates have been used. Suitable alternatives are 3-amino-9-ethylcarbazole (166) or 4-chloro-1-naphthol (167), which are oxidized by the myeloperoxidase to form a colored precipitate in myeloperoxidase-containing cells.

The myeloperoxidase enzyme is sensitive to light, and smears should be stained immediately or sheltered from light. Enzymatic activity may fade over time, so the stain should not be performed in blood smears older than 3 weeks. Permount coverslip mounting medium (Fisher Scientific, Pittsburgh, PA) may cause fading of the stain, and its use should be avoided. Myeloperoxidase is also sensitive to heat and methanol treatment. Erythroid cells may stain for peroxidase after methanol treatment due to a nonenzymatic interaction between the staining reagents and hemoglobin. This is called the pseudoperoxidase or Lepehne reaction.

Sudan Black B

Sudan black B stains intracellular phospholipids and other lipids. The pattern of staining closely parallels the myeloperoxidase reaction, with positive staining of granulocytic cells and eosinophils, weak monocytic staining, and no staining of lymphocytes, although some positivity may be seen in azurophilic granules of lymphoblasts (168). Sudan black B has an advantage over myeloperoxidase in that it may be used to stain older blood or bone marrow smears, and there is little fading of the stain over time (169).

Specific (Naphthol AS-D Chloroacetate) Esterase

This stain, also called the Leder stain, is used to identify cells of the granulocytic series (170 and 171). It fails to stain lymphocytes, and monocytes usually do not stain. Because of enzymatic stability in formalin-fixed paraffin-embedded tissues, this stain is extremely useful for identifying granulocytes and mast cells in tissue sections (172). Thus, it is particularly helpful in diagnosis of extramedullary myeloid tumors (granulocytic sarcoma, chloroma) composed of myeloid blasts found in tissues (173). The esterase enzyme within the cell hydrolyzes the naphthol AS-D chloroacetate substrate (170). This reaction product is then coupled to a diazo salt to form a bright red-pink reaction product at the site of enzymatic activity. The enzyme activity is inhibited by the presence of mercury, acid solutions, heat, and iodine. These may give rise to false-negative staining results.

Nonspecific (a-naphthyl Butyrate or a-naphthyl Acetate) Esterases

These stains are used to identify monocytic cells, but do not stain granulocytes or eosinophils (170 and 174). Mature T lymphocytes stain with a characteristic focal, dotlike pattern. In addition to monocytes, the stain reacts with macrophages, histiocytes, megakaryocytes, and some carcinomas. The a-naphthyl butyrate stain is considered to be more specific, although slightly less sensitive than the a-naphthyl acetate stain (174). Differential staining with the different esterases is seen in megakaryoblasts, which do not stain with the a-naphthyl butyrate but stain with the a-naphthyl acetate substrate (175).

Terminal Deoxynucleotidyl Transferase

Terminal deoxynucleotidyl transferase (TdT) is an intranuclear enzyme that catalyzes the addition of deoxynucleotides triphosphates to the 3'-hydroxyl ends of oligonucleotides or polydeoxynucleotides without need for a template strand (176). TdT is found normally in thymocytes and immature lymphoid cells within the bone marrow, but it is not found in mature lymphocytes. Hence, it is a useful marker in identifying acute lymphoblastic leukemias and lymphoblastic lymphomas (176 and 177). TdT activity is found in approximately 90% of acute lymphoblastic leukemias as well as in a small subset of acute myelogenous leukemias (178 and 179). TdT levels may be measured biochemically (180) and by cytochemical staining with an immunofluorescent technique (181) or immunohistochemical methods (182). Indirect immunofluorescent staining is very sensitive and may be applied to air-dried samples several weeks after collection (183). Immunofluorescent and immunohistochemical methods of TdT detection have also been applied to frozen (184) and paraffin-embedded tissue sections (185), and the enzyme can also be detected in cell suspensions by flow cytometric methods (186 and 187).

A panel of myeloperoxidase (or Sudan black B), a-naphthyl butyrate esterase (or double esterase stain), and TdT staining is often used in characterizing acute leukemias. The more precise lineage assignment provided by flow cytometric analysis is applicable to fresh bone marrow or peripheral blood specimens for differentiating between lymphoid and myeloid leukemias (188). However, because the FAB subclassification of acute myelogenous leukemias is based on the cytochemical staining pattern of the blasts (161 and 162), these stains are usually performed even when flow cytometric analysis is performed. In addition, cytochemical stains may be performed on bone marrow or peripheral blood smears collected routinely at the time of bone marrow examination if a diagnosis of acute leukemia was not suspected and fresh material was not collected for flow cytometric analysis.

Leukocyte Alkaline Phosphatase

Alkaline phosphatase activity is found in the cytoplasm of neutrophils, osteoblasts, vascular endothelial cells, and some lymphocytes. The alkaline phosphatase level of peripheral blood neutrophils is quantitated by the leukocyte alkaline phosphatase (LAP) score, and is useful in differentiation of chronic myelogenous leukemia (CML) from leukemoid reactions and other myeloproliferative disorders (189). The LAP score is usually performed using the Kaplow procedure (190). This method uses a naphthol AS-BI phosphate as the substrate, which is coupled to fast violet B salt by the enzyme to produce a bright red reaction product that is visualized over neutrophils. The LAP score is determined by evaluation of the staining intensity (ranging from 0 to 4+) of 100 counted neutrophils or bands. Normal LAP scores range from 15 to 130, but there may be variation in these ranges between laboratories. Many different disease states may cause elevation or depression of the LAP score (Table 2.7). Patients with CML have low LAP scores (usually between 0 and 13). Other conditions, including paroxysmal nocturnal hemoglobinuria and some myelodysplastic syndromes, may be characterized by low LAP scores. Leukemoid reactions, in response to infection, and other myeloproliferative disorders (myelofibrosis with myeloid metaplasia and polycythemia vera) often have an elevated LAP score (191). There is rapid loss of alkaline phosphatase activity in samples drawn in EDTA anticoagulant (192). The test is optimally performed on fresh capillary blood or on samples anticoagulated with heparin, and should be performed within 48 hours. The blood smears may be held in the freezer for 2 to 3 weeks with little loss of activity.

Table 2.7. Conditions Associated with Abnormal LAP scores (191, 271)

Acid Phosphatase

Acid phosphatase is found in all hematopoietic cells, but the highest levels are found in macrophages and osteoclasts (193). A localized dotlike pattern is seen in many T lymphoblasts, but this staining pattern is not reliable. The tartrate-resistant acid phosphatase (TRAP) is an isoenzyme of acid phosphatase that is found in high levels in the cells of hairy cell leukemia (194). Several methods of measuring TRAP activity have been described, but one using naphthol-ASBI What does ASBI stand for? phosphoric acid coupled to fast garnet GBC What does GBC stand for? is reliable and reproducible (195). Not all cases of hairy cell leukemia stain for TRAP, and staining intensity may be variable. Positive TRAP staining may also be seen in some activated T lymphocytes, macrophages, and some histiocytes (such as Gaucher cells) (196).

Periodic Acid–Schiff

The periodic acid–Schiff (PAS) stain detects intracellular glycogen and neutral mucosubstances, which are found in variable quantities in most hematopoietic cells (197). PAS staining is seen in blasts of both acute lymphoblastic and acute myelogenous leukemias, although there is great variability between cases (160 and 198). Erythroleukemias demonstrate an intense diffuse cytoplasmic positivity with PAS, which may be helpful in diagnosis (199). In addition, PAS staining is very useful in demonstrating the abnormal glucocerebrosidase accumulation in Gaucher disease.


Cellular iron is found as either ferritin or hemosiderin. It is identified in cells by the Perls or Prussian blue reaction, in which ionic iron reacts with acid ferrocyanide to impart a blue color (200). The stain is used to identify iron in nucleated red blood cells (sideroblastic iron) and histiocytes (reticuloendothelial iron) and to identify Pappenheimer bodies in erythrocytes. Normally, red cell precursors contain one or more small (less than 1 µm in diameter), blue granules in 20 to 50% of the cells. When increased numbers of these granules surround at least two-thirds of the nucleus of the red cell precursor, the cell is called a ringed sideroblast. The stain is best used on bone marrow aspirate smears but can also be used on blood films and bone tissue sections. Decalcification of the bone marrow biopsy may lead to loss of iron from the cells, leading to a false impression of low iron.

Toluidine Blue

Toludine blue specifically marks basophils and mast cells by reacting with the acid mucopolysaccharides in the cell granules to form metachromatic complexes (170). Malignant mast cells or basophils may have low levels of acid mucopolysaccharides and may not react with this stain (201).

Methyl Green Pyronin

This stain is used to demonstrate high levels of RNA (pyroninophilia) in the cytoplasm of plasma cells, immunoblasts, and the cells of Burkitt lymphoma (202). It may be helpful in distinguishing Burkitt lymphoma from lymphoblastic lymphoma in children (203).

Immunocytochemical Stains

Immunocytochemical staining is based on the use of an antibody that recognizes a specific antigenic epitope on a cell. Due to the high level of specificity that may be obtained with immunologically based methods, more accurate diagnoses may be made. In general, these types of stains may be applied to blood smears or bone marrow aspirates, cell suspensions, or tissue sections, although not all antibody preparations are equally effective on all types of specimens. A host of antibodies specific to hematopoietic cell types are available commercially.

Immunocytochemical staining of fresh blood or bone marrow cell suspensions and analysis by flow cytometry are becoming increasingly common in clinical laboratories (204). The flow cytometer detects both light scatter data and the presence of specific fluorochrome-labeled antibodies that have bound to the cell surface. Use of different fluorochromes can allow more than one antibody to be studied simultaneously on the same cell by means of different excitation wavelengths. The study of these cell surface markers allows rapid and accurate analysis of lymphomas and leukemias, enumeration of T-cell subsets, and identification of tumor cells. In addition, recent advances have allowed detection of intracytoplasmic or nuclear antigens, such as myeloperoxidase and TdT, by flow cytometric analysis (186). In many cases, particularly in the acute leukemias, the flow cytometric analysis of an acute leukemia provides important prognostic information that is not available though cytochemical staining (205, 206 and 207). Clinical and technical aspects of flow cytometric analysis of hematologic tumors are covered in detail in Chapter 4.

Immunohistochemical staining is the use of specific antibody probes on tissue sections or smears of blood and bone marrow. This allows the localization of a specific antigenic epitope to the cell surface, cytoplasm, or nucleus. The antigen binding may then be detected by immunofluorescence, which requires a special fluorescence microscope, or by enzymatic formation of a colored reaction product linked to the antigen–antibody complex. Immunoenzymatic staining techniques include immunoperoxidase, immunoalkaline phosphatase, and avidin–biotin techniques (208 and 209). These procedures allow study of the specimen with standard light microscopy and provide a permanent record of staining that may be re-examined. In the past, the repertoire of antibodies available for use on paraffin-embedded tissues was limited, and many antibodies required frozen sections of fresh tissues to be used. Over time, however, there has been a large increase in the number of antibodies that can be used on fixed and processed tissues, so frozen section analysis has limited usefulness in light of the severe drawbacks of frozen section morphology (210). Recently several automated immunostaining instruments have become available that allow highly reproducible results and require less technician time and expertise (211 and 212).

Immunohistochemical staining is of great value in establishing a diagnosis for a large-cell undifferentiated tumor (usually involving a differential of lymphoma, carcinoma, or melanoma) (213 and 214). In addition, antibodies are routinely available that allow immunophenotyping of malignant lymphomas and leukemias in paraffin-embedded tissues and aid in recognition of malignant lymphoid infiltrates that may be confused for reactive processes (210, 215 and 216). Many commonly used cytochemical stains, including myeloperoxidase (217 and 218), TdT (185), and TRAP (219), are being converted into immunohistochemical stains that may then be applied to tissue sections when no bone marrow aspirates, touch imprints, or blood smears are available.


Cytogenetic Analysis

Many hematologic malignancies and premalignant conditions are associated with specific cytogenetic changes (220, 221, 222, 223 and 224). These include distinctive changes in chromosome number, translocations, and inversions of genetic material. These chromosomal changes are often associated with activation or increased transcription of oncogenes, and may contribute to acquisition of a malignant phenotype (220 and 225). Thus, cytogenetic analysis has become important in diagnosing hematologic disorders, identifying specific prognostic subgroups, and monitoring for progression of disease or residual disease following therapy. Both standard chromosomal preparations and fluorescent-labeled, in-situ hybridization techniques may be used for cytogenetic analysis of chromosomal changes (226 and 227). Further details about cytogenetic techniques and analysis are provided in Chapter 6.

Molecular Genetics

In addition to standard morphologic analysis and cytogenetics, technology has been developed that allows analysis of molecular changes in tumor cells in the clinical laboratory (228 and 229). By use of Southern blot and polymerase chain reaction (PCR) techniques, hematopoietic proliferations may be studied for alterations in genes that are associated with development of hematopoietic malignancies (230 and 231). Molecular genetic analysis was initially used to detect monoclonality of lymphoid neoplasms by identifying either immunoglobulin (B-cell) or T-cell receptor gene rearrangements (230, 232, 233 and 234). This finding is extremely useful in classification of lymphoproliferative disorders that may be difficult to diagnose on morphologic grounds alone or that lack specific phenotypic markers (235). In the past few years, there has been an explosion in the use of molecular techniques to detect translocations that previously had been detected only by conventional cytogenetics, including bcr-abl translocations seen in chronic myelogenous leukemias and acute leukemias (236, 237) and bcl-2 translocations seen in follicular lymphomas (238 and 239). These molecular studies have an advantage over conventional morphologic and cytogenetic analyses in that they may detect very small populations of malignant cells (as few as 1 to 5% of the cells in a sample) and more rapid test completion (especially with PCR-based testing) (230, 234 and 235).

This degree of sensitivity makes this technology very attractive for the purpose of monitoring for tumor persistence or recurrence after therapy. Previously, molecular genetic studies required collection of fresh or frozen diagnostic material; however, many of the newer PCR assays can make use of formalin-fixed materials with sensitivity similar to that of fresh or frozen materials (240, 241, 242, 243 and 244). This allows analysis to be performed on a wider range of cases, including archival materials. The topic of molecular genetics is covered in further detail in Chapter 7.

Electron Microscopy

The electron microscope allows examination of ultrastructural details of a cell. In the past, electron microscopy was used as a research tool and occasionally as a diagnostic tool for difficult hematologic diagnoses. However, with the advent of increasing numbers of specific immunocytochemical stains, the use of the electron microscope as a diagnostic tool for hematopathologic processes has been largely discontinued.

Erythrocyte Sedimentation Rate

The erythrocyte sedimentation rate (ESR) is a common but nonspecific test that is often used as an indicator of active disease. It reflects the tendency of red blood cells to settle more rapidly in the face of some disease states, usually because of increases in plasma fibrinogen, immunoglobulins, and other acute-phase reaction proteins. In addition, changes in red cell shape or numbers may affect the ESR. Sickle cells and polycythemia tend to decrease the ESR, whereas anemia may increase it. ESR also increases with age in otherwise healthy people (although it tends to fall in patients over the age of 75) (245) and tends to be higher in women (246). People with liver diseases or carcinomas or other serious diseases may have a normal to low ESR because of an inability to produce the acute-phase proteins.

A common cause of ESR elevation is infection, but monoclonal gammopathy must be ruled out in patients who have a persistent, unexplained elevation in ESR. Elevated ESRs are also seen with pregnancy, malignancies, collagen vascular diseases, rheumatic heart disease, and other chronic disease states, including human immunodeficiency virus infection (247). The ESR is a poor screening test in asymptomatic individuals, detecting elevations in 4 to 8% of them, and hence should not be used to screen asymptomatic people for disease (248). The test is probably best used in patients with vague complaints to aid in the decision to undergo further tests (249), or as a tool to follow the course of a disease state. ESR is most commonly used for monitoring of the clinical course of temporal arteritis, rheumatoid arthritis, or polymyalgia rheumatica (250 and 251). It may also be useful for monitoring for relapse of Hodgkin disease or non-Hodgkin lymphomas (252).

The ESR is measured by the Westergren or Wintrobe method, or variations of these methods (253). Both are measured in millimeters per hour, but the normal values for each method vary because of differences in tube length and shape. Both methods require correction for the degree of patient anemia. Several technical variations have been introduced to the method of ESR determination, including micromethods, sedimentation at a 45° angle, and the ? sedimentation rate. The ? sedimentation rate measures erythrocyte packing in four 45-second cycles of dispersion and compaction in capillary tubes. This requires a special instrument, the Zetafuge, but gives results on very small amounts of blood that are not affected by anemia (254).

Plasma and Blood Viscosity

Plasma viscosity measurements are advocated by some authors as being superior to ESR measurements for monitoring disease states, particularly in autoimmune diseases and diseases characterized by the secretion of large amounts of immunoglobulin into the plasma (255, 256 and 257). Plasma viscosity measurements have the advantage of no red cell influences on the value obtained and yield a narrower reference range of normal values than observed with ESR. However, plasma viscosity is used more rarely than ESR, probably reflecting clinicians' familiarity with the latter (257). Like ESR, plasma viscosity may increase with age (245). Direct measurement of acute-phase proteins, such as protein C, may also be used to monitor the course of inflammatory diseases (258). However, these tests are usually more expensive than ESR determinations and may not provide sufficient additional clinical information to justify the added expense (259).

Whole blood viscosity measurements are of limited clinical utility because the measured blood viscosity may have little bearing on the viscosity of the blood in the circulation. Increased blood viscosity may contribute to the morbidity and mortality of patients with sickle cell disease, polycythemia, and ischemic vascular disease.

Total Quantity of Blood

In most cases, the total number of erythrocytes is closely related to the red cell concentration of the blood or hematocrit. However, in some cases blood volume may not reflect erythrocyte concentration, including immediately following severe hemorrhage, severe dehydration, or overhydration. To accurately assess the blood volume in these patients, plasma volume or red cell volume must be determined. The plasma volume is measured by dilution methods. A substance that is confined to the intravascular plasma compartment, such as Evans blue dye (262), 131 I-labeled albumin, or radioactive indium-labeled transferrin (260 and 261), is injected and the volume of distribution calculated from the degree of dilution of the injected substance over 15 to 30 minutes. Radiolabeled albumin is the most commonly used plasma label, but corrections must be made because the label is gradually removed from the circulation into the extravascular space (261 and 263), leading to errors of 10% or more in plasma volume determinations (260).

Total red cell volume is calculated by the Ashby technique, which uses radiolabeled red blood cells. A number of radioisotopes may be used, but 51Cr and 99mTc are the most common (264 and 265). The red cell volume is then calculated by the dilution of the labeled cells over time using the following formula:

Usually the measurements are made after a 15-minute interval, although longer periods may be needed in patients with viscous blood due to high hematocrits to ensure complete labeled cell mixing. Total red cell volume measurements must be corrected in patients with enlarged spleens because of sequestration of the labeled cells within this organ. Red cell volume may also be calculated from the total plasma volume and measured hematocrit by means of the following equation:

Total plasma volume may be useful in monitoring fluid and blood replacement. Red cell volume measurements are used to document true polycythemia when an elevated hematocrit is present (266 and 267). Total blood volume may be calculated from the sum of total red cell volume and plasma volume measurements.


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