Sherrie L. Perkins
Careful assessment of the blood elements is often the first step in assessment of hematologic function and diagnosis. Many hematologic disorders are defined by specific results of blood tests. Examination of blood smears and hematologic parameters often yields important diagnostic information and allows broad differential diagnostic impressions to be formed, directing additional specific testing. Cellular morphology, in concert with quantification of the blood cellular components and evaluation of a variety of parameters relating to cellular size and shape, is required. This chapter introduces the fundamental concepts and limitations 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.
Blood elements include erythrocytes, or red cells; leukocytes, or white 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 analyses are covered in this chapter. Red blood cells (RBCs) are the most numerous blood cells in the blood and are required for tissue respiration. RBCs lack nuclei and contain hemoglobin, an iron-containing protein that acts in the transport of oxygen and carbon dioxide. White blood cells (WBCs)serve in immune function and include a variety of cell types that have specific functions and characteristic morphologic appearances. In contrast to red cells, WBCs are nucleated and include neutrophils, lymphocytes, monocytes, eosinophils, and basophils. Platelets are cytoplasmic fragments derived from marrow megakaryocytes that function in coagulation and hemostasis.
Evaluation of the blood requires quantification of each of the cellular elements by either manual or automated methods. Automated methods, using properly calibrated equipment (1), are usually more precise than manual procedures. In addition, automated methods may provide additional data describing cellular characteristics such as cell volume. However, the automated measurements describe average cellular characteristics but do not adequately describe the scatter of individual values around the average. Hence, a bimodal population of small (microcytic) and large (macrocytic) RBCs might be reported as normal cell size. Therefore, a thorough blood examination also requires microscopic evaluation of a stained blood film to complement hematology analyzer data (2,3).
Specimen Collection
Proper specimen collection is required for acquisition of reliable and accurate laboratory data for any hematologic specimen. Before a specimen is obtained, careful thought as to what studies are needed will aid in collection of the material and prevent inadequate or improper 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 significantly affect hematologic parameters (4,5,6). Similarly, the age of the specimen may affect the quality of the data collected (7,8). 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 collection tubes containing anticoagulant. The three most commonly used anticoagulants are tripotassium or trisodium 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 thrombin formation. EDTA is the preferred anticoagulant for blood counts because it produces complete anticoagulation with minimal morphologic and physical effects on cells (9). Heparin causes a bluish coloration of the background when a blood smear is stained with Wright-Giemsa stains, but does not affect cell size or shape. Heparin is often used for red cell testing, osmotic fragility testing, and functional or 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; Becton Dickinson, Franklin Lakes, NJ), which contain the correct concentration of anticoagulant when filled appropriately, thereby minimizing error (10). Anticoagulated blood may be stored at 4°C for a 24-hour period without significantly altering cell counts or cellular morphology (7). However, it is preferable to perform hematologic analysis as soon as possible after the blood is obtained.
Reliability of Tests
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 both 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 blood smear examination (10) and automated blood cell analyzers (11). Automated instrumentation requires regular quality assurance evaluations and careful calibration to reach expected performance goals and ability to collect accurate and reproducible data (1,12). In addition, the International Consensus Group for Hematology Review has suggested criteria that should lead to manual review of a specimen after automated analysis and differential counting (2).
Cell Counts
Cell counts are important parameters in evaluating the blood. Cell counts may be determined either manually or by automated hematology analyzers. Whether performed by manual or automated methodologies, the accuracy and precision of the counts depend on proper dilution of the blood sample, even distribution of cells and precise sample measurement. As blood contains large numbers of cells, sample dilution is usually required for accurate analysis. The type of diluent is dependent on the cell type to be enumerated. 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 to simplify counting. 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, or enumeration of cells. The highest degree of precision occurs when a large number of cells can be evaluated. Clearly, automated methods are superior to manual methods for counting large numbers of cells and minimizing statistical error. Table 1.1 lists the comparable values of reproducibility for automated and manual (hemocytometer) counting methods.
Manual counts are done using a microscope after appropriate dilution of the sample in a hemocytometer, a specially constructed counting chamber that contains a specific volume. Red cells, leukocytes, and platelets may be counted. Due to the inherent imprecision of manual counting and the amount of technical time required, most cell counting is now performed by automated instruments that increase the accuracy and speed of analysis by the clinical laboratory, minimizing levels of human manipulation for test entry, sampling, sample dilution, and analysis (13). With increasing automation, some hematology analyzers can be coupled with instruments performing other laboratory tests using the same tube of blood (14). There are a variety of different automated hematology analyzers available, dependent on the volume of samples to be tested and the needs of the physician ordering testing. The analyzers range in price and workload capacity from those that would be appropriate for an individual physician’s office or point-of-care facility to those needed in a busy reference laboratory with capacity for over 100 samples to be analyzed per hour (13).
Most automated hematology analyzers perform a variety of hematologic measurements, in addition to cell counting, such as hemoglobin concentration (Hb), red cell size, and leukocyte differentials. Many instruments also perform more specialized testing, such as reticulocyte counts (15). The ability of analyzers to perform accurate WBC differential counts, particularly those that can perform a five-part differential (enumerating neutrophils, lymphocytes, monocytes, eosinophils, and basophils), has been a significant technologic advance over the past 15 years. Automated methods for white cell counts and differentials use several distinct technical approaches, including measurement of electrical impedance, differential light scatter, optical properties, or antigen staining either alone or in combination (16,17).
Most of the newer-generation hematology analyzers utilize optical flow cytometric technologies with or without additional cytochemical staining to detect specific cell types such as red cells, white cells, and platelets (Fig. 1.1) (13). The newer analyzers have additional ability to detect reticulocytes as a part of the normal complete blood count (CBC) differential using a fluorescent RNA dye and many will also enumerate nucleated red blood cell numbers based on their optical properties (18). In addition, many of the current analyzers do auto sampling directly from tubes and use a very small sample ranging from 35 to 150 μl for a full CBC analysis. Using flow cytometric technologies, some analyzers also have the ability to detect specific blood cell populations by specific antigen expression, such as detection of CD34 peripheral blood stem cells (19). Integration of data from cytochemical or antigenic staining and light scatter properties has improved the accuracy of the five-part differential and decreased the numbers of unidentifiable cells requiring technician review for identification.
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Figure 1.1. Optical flow cytometric 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. Santa Clara, CA: Abbott Diagnostics, 1993.) |
Instruments from Abbott Laboratories (Cell-Dyn Sapphire System) (13,20), ABX Diagnostics (Pentra XL), and Sysmex (XT-2000i and XE-series) (13,21,22) primarily utilize fluorescent-based flow cytometry as the modality for analysis. Each system has slightly different fluorochrome staining combinations that aid in the identification of white cells, red cells, and platelets in combination with light scatter characteristics. All provide integrated reticulocyte counts and five-part differentials. Workload capacities range from 70 to 106 samples analyzed per hour. When reticulocytes are ordered as a part of the differential, the capacity falls to between 40 and 60 samples per hour (allowing for the staining and detection of the RNA dye fluorescence). Instruments by Bayer (Advia 2120) use a combination of flow cytometric techniques and a cytochemical peroxidase stain for the five-part differential. This instrument integrates electrical impedance data, flow cytometric light scatter, characteristic fluorescent staining, and cytochemical staining to generate an accurate white blood cell differential. The Bayer technology also calculates hemoglobin levels, claiming that this causes less interference by high white blood cell counts or lipemia in the specimen (13,23,24). Instruments from Beckman/ Coulter (Coulter LH 755 series) also utilize electrical impedance or conductivity in combination with light scatter approaches, integrating these technologies to provide full analysis and five-part differentials (Fig. 1.2). The Beckman/Coulter series includes nucleated red blood cells and reticulocyte counts in every differential. Its capacity is 45 samples per hour when reticulocytes are included and 100 samples per hour for a CBC without reticulocyte counts (13).
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Figure 1.2. Histograms and printout generated by the Coulter automated hematology analyzer utilizing light scatter and electrical impedance. BA, basophil; EO, eosinophil; HCT, hematocrit; HGB, hemoglobin; LY, lymphocyte; MCH, mean corpuscular hemoglobin; MCHC, mean corpuscular hemoglobin concentration; MCV, mean corpuscular volume; MO, monocyte; MPV, mean platelet volume; NE, neutrophil; PLT, platelet; RBC, red blood cell; RDW, red cell distribution width; WBC, white blood cell. |
Red Blood Cell Analytic Parameters
Red blood cells are defined by three quantitative values: The volume of packed red cells or hematocrit (Hct), the amount of hemoglobin, and the red cell concentration per unit volume. Three additional indices describing average qualitative characteristics of the red cell population are also collected. These are mean corpuscular volume (MCV), mean corpuscular hemoglobin (MCH), and mean corpuscular hemoglobin concentration (MCHC). All of these values are routinely collected and calculated by automated hematology analyzers, largely replacing many of the previously used manual or semi-automated methods of RBC characterization, with certain exceptions as noted below. The use of hematology analyzers imparts a high degree of precision compared to manual measurements and calculations (Tables 1.1 and 1.2).
Volume of Packed Red Cells (Hematocrit)
The Hct is the proportion of the volume of a blood sample that is occupied by red cells. Hct 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 (25). The height of the column of red cells after centrifugation compared with total blood sample volume yields the Hct. Macromethods (using 3-mm test tubes) with low-speed centrifugation or micromethods using capillary tubes and high-speed centrifugation may be used.
Manual methods of measuring Hct are simple and accurate means of assessing red cell status. They are easily performed with little specialized equipment, allowing adaptation for situations in which automated cell analysis is not readily available or for office use. However, several sources of error are inherent in the technique. The spun Hct measures the red cell concentration, not red cell mass. Therefore, patients in shock or with volume depletion may have normal or high Hct measurements due to hemoconcentration despite a decreased red cell mass. Technical sources of error in manual Hct determinations usually arise from inappropriate concentrations of anticoagulants (26), poor mixing of samples, or insufficient centrifugation (25). Another inherent error in manual Hct 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 relatively more plasma (27,28). It should be noted that abnormal red cells (e.g., sickle cells, microcytic cells, macrocytic cells, or spherocytes) often trap higher volumes of plasma due to increased cellular rigidity, possibly accounting for up to 6% of the red cell volume (28). Very high Hcts, as in polycythemia, may also have excess plasma trapping. Manual Hct methods typically have a precision coefficient of variation (CV) of approximately 2% (27).
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Table 1.2 Reproducibility of Red Cell Indices |
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Automated analyzers do not depend on centrifugation techniques to determine Hct, but instead calculate Hct by direct measurements of red cell number and red cell volume (Hct = red cell number/red cell volume). Automated Hct values closely parallel manually obtained measurements, and the manual Hct is used as the reference method for hematology analyzers (with correction for the error induced by plasma trapping). Errors of automated Hct calculation are more common in patients with polycythemia (29) or abnormal plasma osmotic pressures (30). Manual methods of Hct determination may be preferable in these cases. The precision of most automated Hcts is <1% (CV) (13).
Hemoglobin Concentration
Hemoglobin is an intensely colored protein, allowing its measurement by 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 (containing potassium ferricyanide and potassium cyanide) (31,32). Sulfhemoglobin is not converted but is rarely present in significant amounts. The absorbance of the cyanhemoglobin is measured in a spectrophotometer at 540 nm to determine Hb. This technique is used both in manual determinations and automated hematology analyzers. Hb 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 (33,34,35). With automated methods the precision for hemoglobin determinations is <1% (CV) (13).
Red Cell Count
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 (36). In hematology analysis both erythrocytes and leukocytes are counted after whole blood dilution in an isotonic solution. 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, when marked leukocytosis is present, red cell counts and volume determinations may be erroneous unless corrected for white cells. The observed precision for RBC counts using automated hematology analyzers is <1% (CV) (13) compared with a minimum estimated value of 11% with manual methods (36).
Mean Corpuscular Volume
The average volume of the red blood cell is a useful parameter that is used in classification of anemias and may provide insights into pathophysiology of red cell disorders (37,38,39). The MCV is usually measured directly with automated instruments but may also be calculated from the erythrocyte count and the Hct by means of the following formula (25):
MCV = Hct (L/L) ÷ 1,000/red cell count (1012/L)
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% (13), compared to ∼10% for manual methods (27).
Agglutination of cells, as in cold agglutinin disease or paraproteinemia, may result in a falsely elevated MCV (40). Most automated analyzers gate out MCV values above 360 fl, thereby excluding most red cell clumps, although this may falsely lower Hct determinations. In addition, severe hyperglycemia (glucose >600 mg/dl) may cause osmotic swelling of the red cells, leading to a falsely elevated MCV (30,41).
Mean Corpuscular Hemoglobin
MCH is a measure of the average hemoglobin content per red cell. It may be calculated manually or by automated methods using the following formula (25):
MCH = hemoglobin (g/L)/red cell count (1012/L)
MCH is expressed in picograms (pg, or 10-12 g). Thus, the MCH is a reflection of hemoglobin mass. In anemias secondary to impaired hemoglobin synthesis, such as iron deficiency anemia, hemoglobin mass per red cell decreases, resulting in a lower MCH value. MCH measurements may be falsely elevated by hyperlipidemia (35), as increased plasma turbidity will erroneously elevate hemoglobin measurement. Centrifugation of the blood sample to eliminate the turbidity followed by manual hemoglobin determination allows correction of the MCH value. Leukocytosis may also spuriously elevate MCV values (33). The CV for automated analysis of MCH is <1% in most modern analyzers, compared with approximately 10% for manual methods (27).
Mean Corpuscular Hemoglobin Concentration
The average concentration of hemoglobin in a given red cell volume or MCHC may be calculated by the following formula (25):
MCHC = hemoglobin (g/dl)/Hct (L/L)
The MCHC is expressed in grams of hemoglobin per deciliter of packed red blood cells, representing the ratio of hemoglobin mass to the volume of red cells. 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 Hb may lead to crystallization. The accuracy of the MCHC determination is affected by factors that affect measurement of either Hct (plasma trapping or presence of abnormal red cells) or hemoglobin (hyperlipidemia, leukocytosis) (33). The CV for MCHC for automated methods ranges between 1.0 and 1.5% (13).
As noted above, the MCV, MCH, and MCHC reflect average values and may not adequately describe blood samples when mixed populations of red 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 and normocytic. It is important to examine the blood smear as well as red cell histograms to detect such dimorphic populations (2). The MCV is an extremely useful value in classification of anemias (13,39,42), but the MCH and MCHC often do not add significant, clinically relevant information. However, the MCH and MCHC play an important role in laboratory quality control because these values will remain stable for a given specimen over time (12).
Red Cell Distribution Width
The red cell distribution width (RDW) is a red cell measurement that quantitates cellular volume heterogeneity reflecting the range of red cell sizes within a sample (37,43,44). RDW has been proposed to be useful in early classification of anemia as it becomes abnormal earlier in nutritional deficiency anemias than other red cell parameters, especially in cases of iron deficiency anemia (37,45). RDW is particularly useful in characterizing microcytic anemia, allowing to distinguish between uncomplicated iron deficiency anemia (high RDW, normal to low MCV) and uncomplicated heterozygous thalassemia (normal RDW, low MCV) (37,45,46,47), although other tests are usually required to confirm the diagnosis (48). RDW is also useful in identifying red cell fragmentation, agglutination, or dimorphic cell populations (including patients who have had transfusions, have sideroblastic anemias, or have been recently treated for a nutritional deficiency) (45,49).
Reticulocyte Counts
Determination of the numbers of reticulocytes or immature, nonnucleated red blood cells that still retain RNA provides useful information about the bone marrow’s capacity to synthesize and release red cells in response to a physiologic challenge, such as anemia. In the past, reticulocyte counts were performed manually using supravital staining with methylene blue that will stain precipitated RNA as a dark blue meshwork or granules (at least two per cell), allowing reticulocytes to be identified and enumerated manually (50). Normal values for reticulocytes in adults are 0.5 to 1.5%, although they may be 2.5 to 6.5% in newborns (falling to adult levels by the second week of life). Because there are relatively low numbers of reticulocytes, the CV for reticulocyte counting is relatively large (10 to 20%) (51).
To increase accuracy of reticulocyte counting, automated detection methods to detect staining allow for many more cells to be analyzed, thereby increasing accuracy and precision of counts (15,51,52). Most of the newest automated hematology analyzers have automated reticulocyte counting as part of the testing capabilities and allow reticulocyte counts to be included with routine complete blood count parameters. Reticulocytes are detected by a fluorescent dye that binds to RNA. Comparisons of stand-alone instruments and integrated hematology analyzers demonstrate superior accuracy when compared to manual counting methods, with CVs of 5 to 8% (13,53).
Leukocyte Analysis
White Blood Cell Counts
Leukocytes may be enumerated by either manual methods or automated hematology analyzers. Leukocytes are counted after 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 (usually a 1:20 dilution, although it may be less in cases of leukocytopenia or more with leukocytosis). Manual counts are done using a hemocytometer or counting chamber. As with red cell counts, manual leukocyte counts have more inherent error, with CVs 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 CVs in the 1 to 3% range (13). Automated leukocyte counts may be falsely elevated in the presence of cryoglobulins or cryofibrinogen (54), aggregated platelets (55), and nucleated red blood cells or when there is incomplete lysis of red cells, requiring manual counting. Falsely low neutrophil counts have also been reported due to granulocyte agglutination secondary to surface immunoglobulin interactions (56,57).
Leukocyte Differentials
White cells are analyzed to find the relative percentage of each cell type by a differential leukocyte count. Uniform standards for performing manual differential leukocyte counts on blood smears have been proposed by the NCCLS (58) to ensure reproducibility of results between laboratories. It is important to scan the entire blood smear at low power to ensure that all atypical cells and cellular distribution patterns are recognized. 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-pushed 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 (58).
In manual leukocyte counts, three main sources of error are encountered: Distribution of cells on the slide, cell recognition errors, and statistical sampling errors. Poor blood smear preparation and staining are major contributors to cell recognition and cell distribution errors (60). 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. The CV in manual counts is between 5 and 10% and is also highly dependent on the skill of the technician performing the differential. Accuracy may be improved by increasing the numbers of cells counted, but for practical purposes, most laboratories will do a differential on 100 white cells (61,62).
Automated leukocyte differentials markedly decrease the time and cost of performing routine examinations as well as increasing accuracy to a CV of 3% to 5% (61,62,63). However, automated analysis is incapable of accurately identifying and classifying all types of cells and is particularly insensitive to abnormal or immature cells. Therefore, most analyzers will flag possible abnormal white cell populations, indicating the need for examination by a skilled morphologist for identification (63). The capacity for performing automated leukocyte differentials is incorporated into hematology analyzers, which identify cells on the basis of cellular size, cell complexity, or staining characteristics as a part of the complete blood count, allowing for generation of a five-part differential count that enumerates neutrophils, monocytes, lymphocytes, eosinophils, and basophils (13).
Most systems 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 an optical flow cell in a continuous stream so that single cells are analyzed for cell size (dark-field light scatter) and complexity (forward scatter) (Fig. 1.1) or cytochemical characteristics of myeloperoxidase staining (bright-field detector). The data are plotted as a scattergram (Fig. 1.2), which allows white cells to be divided into a five-part differential (neutrophils, lymphocytes, monocytes, eosinophils, and basophils) and also indicates large unstained or unclassified cells. Lymphocytes are characterized as small (low-scatter) unstained cells. Larger atypical lymphocytes, blasts, or circulating plasma cells fall into the larger cell with a low-complexity channel. Neutrophils have higher complexity and appear as larger cells. Eosinophils appear smaller than neutrophils because they tend to absorb some of their own light scatter. Monocytes have lower levels of complexity and are usually found between neutrophils and lymphocytes. To enumerate basophils, which lack specific staining characteristics and are difficult to enumerate with automated flow-through techniques, a basophil-nuclear lobularity channel may be utilized. 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 relatively large 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. Abnormal cell populations will generate a flag, indicating a need for morphologic review of the peripheral smear (2). Analysis using this technique examines thousands of cells per sample, increasing statistical accuracy (13).
Most of the current hematology analyzers have settings that will allow for evaluation of very hypocellular specimens, such as body fluids. They may be used for analysis of these fluids for enumeration of red cells and white cells, as well as providing a five-part differential count of the white cells. Because of the sampling of higher numbers of cells in these relatively hypocellular specimens, accuracy of cell counts and differential counting is improved (23,24,64).
A few instruments, such as the Advia 2120 and the Coulter LH755, also have integrated automated blood smear preparation technology allowing smear preparation directly from the tube that the CBC analysis is performed upon. Thus, the tube is loaded once into a single machine to allow for CBC analysis as well as peripheral blood smear preparation (13). Abbott Laboratories’ Cell-Dyn and Sysmex also have automated slide makers and stainers, which provide wedge smears from up to 80 slides per hour directly from CBC tubes; however, these are free-standing instruments separated from the hematology analyzer. The automated push smear technology helps to provide technical uniformity in blood smear preparation as well as staining. However, there is less flexibility in adjusting stain characteristics. These instruments sample directly from the tube, also minimizing handling of samples by technical staff.
In addition to technology that has the ability to make and stain slides, some automated differential technology is now available. Sysmex (Cellavision) has an automated image analyzer that by pattern recognition will capture digital images of 100 to 500 cells in a smear and classify them into morphologic categories to provide a five-part differential. Depending on the model utilized, these technologies have the abilities to perform between 20 and 60 automated digital differentials per hour. The systems have the capacity to store images and are useful in training technologists in the recognition of the specific cell types as well as providing an easily accessible means whereby smears obtained at different times from a single patient may be compared morphologically. These systems have limitations in ability to identify morphologically abnormal cells, so specimens with dysplastic changes, unusual morphologic variants, or significant artifacts may not be evaluable or may provide false data. Often these systems will place a certain percentage of cells in an unclassifiable area, requiring review by a technologist for definitive identification of the cell type and completion of the differential. As microscopy is automated, there is a uniform scanning of each slide and images are presented on a computer screen, decreasing technician microscope time and scanning pattern variability, and also allowing for the ability to greatly enlarge digitally captured images (65).
Platelet Analysis
Platelets are anucleate cytoplasmic fragments that are 2 to 4 microns in diameter. As with the other blood components, they may be counted by either manual or automated methods. Manual methods involve dilution of blood samples and enumeration in a counting chamber or hemocytometer using phase contrast microscopy. Sources of error are similar to other manual counting techniques and include dilution errors and low numbers of events counted. The CV, especially in patients with thrombocytopenia, may be >15% (66,67). Platelets are counted in automated hematology analyzers after removal of red cells by sedimentation or centrifugation, or using whole blood. Platelets are identified by light scatter, impedance characteristics, or platelet antigen staining (13,68). These give highly reliable platelet counts with a CV of <2%. Falsely low platelet counts may be caused by the presence of platelet clumps or platelet agglutinins (55) or adsorption of platelets to leukocytes (69,70). 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 (71,72).
Automated hematology analyzers also determine mean platelet volume (MPV), which has been correlated with several disease states (73,74). In general, MPV has an inverse relationship with platelet number, with larger platelet volumes (secondary to new platelet production) seen in thrombocytopenic patients in whom platelets are decreased due to peripheral destruction (as in idiopathic thrombocytopenic purpura) (75). MPV is also increased in myeloproliferative disorders (76). However, it should be noted that platelets tend to swell during the first 2 hours in EDTA anticoagulant, shrinking again with longer storage (77,78). Decreased MPV has been associated with megakaryocytic hypoplasia and cytotoxic drug therapy (79).
Reticulated platelets are newly released platelets that retain residual RNA, analogous to red cell reticulocytes. Reticulated platelet counts give an estimate of thrombopoiesis and may be useful in distinguishing platelet destruction syndromes from hypoplastic platelet production (80). Reticulated platelets are usually detected by flow cytometric methods using thiazole orange dyes that bind to RNA (81). Normal values vary between 3 and 20%, and 2.5- to 4.5-fold increases in reticulated platelet counts are seen in the clinical setting of idiopathic thrombocytopenic purpura (82). Increased reticulated platelets may herald the return of platelet production after chemotherapy (83). Although automated hematology analyzers offering reticulated platelet counts are not yet available, it is anticipated that this test may be incorporated in newer models, similar to the reticulocyte count.
Advantages and Sources of Error with Automated Hematology Analyzers
Clearly, the use of automated hematology analyzers has reduced laboratory costs and turnaround time coincident with improving the accuracy and reproducibility of blood counts. The CV for most of the parameters measured is in the range of 1 to 2%. This level of reproducibility is not achievable with the use of most manual techniques (Tables 1.1 and 1.2).
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, negatively affect data accuracy. Electrical or mechanical failures as well as minor voltage fluctuations may induce marked errors in data collection. 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 (84). Reference methods for instrument calibration have been developed by both the NCCLS and the ICSH and are widely used by hospital and clinical laboratories to ensure regulatory compliance (12,58,85).
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 1.3). 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 (54). High levels of paraprotein may lead to falsely elevated hemoglobin levels, therefore affecting MCH and MCHC calculations (34). Older analyzers reported spurious increases in hemoglobin levels when white cell counts exceed 30 ÷ 109/L due to increased turbidity, but this is decreased with newer flow systems so that hemoglobin levels remain extremely accurate in the face of white blood cell counts as high as 100 ÷ 109/L (13). Extremely high white cell counts may also falsely raise the red cell count and Hct as the white cell count is incorporated into the red cell count. High glucose levels (>400 to 600 mg/dl) and the associated hyperosmolarity cause red cell swelling and generate a high MCV and Hct with a falsely low MCHC (30,86). Increased turbidity associated with hyperlipidemia may also cause falsely elevated hemoglobin determinations, MCH, and MCHC (35).
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Table 1.3 Disorders and Conditions That May Reduce the Accuracy of Blood Cell Countinga |
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Despite the high level of accuracy and precision, the automated hematology analyzers usually have data that create a warning flag in 10 to 25% of samples, requiring manual examination of the blood smear (2,13). Blood smear examination still plays an important role in characterizing these samples or showing findings outside the preset parameters for the laboratory. In addition, some cells require morphologic examination to identify, such as Sézary cells (87), and red cell morphology is best analyzed by direct smear examination (39,42).
Morphologic Analysis of Blood Cells
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 which are associated with specific disease states are found in Table 1.4. 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 and staining techniques and familiarity with the morphologic appearances of normal and pathologic cells.
Preparation of Blood Smears
Blood films may be prepared on either glass slides or coverslips. Each method has specific advantages and disadvantages (88,89). Blood smears are often prepared from samples of anticoagulated blood remaining from automated hematologic analysis or may be prepared automatically by the hematology analyzer at the time of analysis (3,13). However, artifacts in cell appearance and staining may be induced by anticoagulant. Optimal morphology and staining are obtained from nonanticoagulated blood, most often from a fingerstick procedure. Mechanical dragging of the cells across the glass of the slide or coverslip and uneven distribution of blood may also distort the cells; however, these artifacts may be minimized with proper technique (90).
Coverslip smears (Fig. 1.3A) are prepared using a good grade of flat, no. 1, 0.5-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, 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 excessively thick areas (90,91).
Blood smears may also be prepared on clean glass slides by the wedge method (Fig. 1.3B). This often leads to irregular distribution of cells on the slide, a distinct disadvantage over the coverslip procedure. However, glass slides are less fragile, are 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 approximately 1 to 2 cm from one end. A second spreader slide is placed at a 30- to 45-degree angle and moved backward to make contact with the blood drop. The blood drop will spread along the slide edge, and 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 (90,91,92).
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 (92,93). Mechanical spreaders mimic the manual technique and are useful when large numbers of blood smears are prepared (59). In general, smears made by automated techniques are usually inferior to those made by an experienced technician.
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Table 1.4 Pathologic Red Cells in Blood Smears |
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Routine Staining of Blood Smears
Blood smears are usually stained with either Wright or May-Grünwald-Giemsa stains. Both stains are modifications of the Romanowsky procedure (94). The stain may be purchased commercially or may be made in the laboratory. The basic stain is formulated from methylene blue and eosin. Giemsa stains use known quantities of acid bichromate to form the converted azure compounds. The Wright stain formulation uses sodium bicarbonate to convert methylene blue to methylene azure, which stains the cell. All types of Romanowsky stains are water insoluble but can be 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 when binding 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 will 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.
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Figure 1.3. 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 J. Clinical laboratory methods, 9th ed. St. Louis: C.V. Mosby, 1982.) |
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 of the slide 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 pH of the solutions, and new buffers often correct the problem.
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 100÷) or high-power dry lens (40÷) 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 (88,89,90).
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 blood cells, termed rouleaux. This finding may be mimicked in normal patients in areas of the smear where the red cells are too closely packed. However, if rouleaux 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. Areas of the blood smear that are too thin will have loss of red cell central pallor, mimicking spherocytes (95).
Red cells should be uniform in size and shape with an average diameter of 7.2 to 7.9 μm. This may be evaluated by use of a micrometer or by comparison with the diameter of a small lymphocyte nucleus, which is approximately the same size or slightly smaller. 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 of <6 μm.
Normal erythroid cells are round. Variations in red cell 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) (95). In addition, red cell fragments or schistocytes suggestive of red cell mechanical destruction are more easily detected by blood smear examination (96,97).
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 blood is not anticoagulated or a fingerstick preparation is used, and this may cause the spurious impression of a low platelet count (88,89,95,98).
Leukocyte morphology and distribution are analyzed last. The number of leukocytes may be estimated by scanning the blood film at an intermediate power. Mechanical effects leading to abnormal distribution of larger cells should be excluded by examination of the edges of the blood film in particular (3,58). White cells 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 and excessive spreading). Care must be taken when making the smear because cells, particularly neoplastic cells, may be more easily disrupted by excessive mechanical pressure than normal leukocytes. 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 blood smear include neutrophils, eosinophils, basophils, lymphocytes, and monocytes. The presence of immature myeloid cells (myelocytes, metamyelocytes, promyelocytes, and blasts) is distinctly abnormal (2,3,95). At least 100 cells should be identified and counted to yield a manual white blood cell differential (2,3,58). 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 (99,100). In contrast, many myelodysplastic disorders are characterized by hypogranularity of neutrophils in addition to abnormal nuclear segmentation (95). Cytoplasmic inclusions may be seen in some storage disorders, lysosomal disorders, or infections (95,101).
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 (91). Wet mounts are used to detect sickling of red cells, spherocytes, and parasites within erythrocytes (102). Some organisms, such as spirochetes and trypanosomes, may be detected by movement.
Supravital staining is performed on living, motile cells and helps avoid artifacts induced by smear preparation, fixation, and staining (103). 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 or 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 (104).
The most commonly used supravital stain is new methylene blue or brilliant cresyl blue, used for manual reticulocyte determinations. These stains allow visualization of the reticulin network of erythrocyte ribosomes in newly formed red blood cells. 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 (52). Reticulocytes are not identified positively on Wright-stained blood smears, although their presence is suggested by polychromatophilia of red blood cells. As noted previously, many newer 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. The degree of ribosomal staining may also be quantitated to allow assessment of reticulocyte age (13,52,105). Automated reticulocyte counts may have increased errors in the presence of Heinz bodies (106) or Howell-Jolly bodies (52,107) in the red cells. Normal reference values for reticulocytes are influenced by patient age, sex, and physical activity level (108).
Bone Marrow Examination
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 cytologic 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 (109). 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 (109).
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 that 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 (109,110,111). 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 (111). 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 approximately 1 cm thick at the second intercostal space, so care must be taken to avoid penetrating the chest cavity, although sternal bone marrow needles have guards to prevent penetration of the needle beyond the sternal plate. 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 (95). The anterior iliac crest may be used if previous radiation, surgery, or patient discomfort do not allow a posterior approach (111,112).
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. 1.4). Some models, primarily used for sternal bone marrow aspiration procedures, have adjustable guards that limit the extent of needle penetration and reduce morbidity (95).
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Figure 1.4. 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). |
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Figure 1.5. Bone marrow aspirate smear stained with Wright Giemsa stain. The bone marrow aspirate shows a central spicule with dispersion of hematopoietic precursor cells around the spicule. The preparation allows for optimal evaluation of cytologic features of the bone marrow precursor cells. Panel A (low power) demonstrating distribution of hematopoietic cells near the darkly staining bone marrow spicule in a bone marrow aspirate. Panel B (high power) demonstrating cytologic features of bone marrow aspirate hematopoietic cells. |
In most cases, marrow aspiration and biopsy may be carried out with little risk of patient discomfort, provided adequate local anesthesia is used. Apprehensive patients may be sedated before the procedure, but this is usually not necessary (113,114). 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 tissue, 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 2.0 ml of bloody fluid is obtained (109). 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, relocation to another sampling site may be required (109,113).
The aspirated material is given to a technical assistant, who makes smears of the material (Fig. 1.5) 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 using either coverslips or slides to spread the marrow (Fig. 1.3). After smears are made, the aspirate may be allowed to clot to form a histologic clot section for processing. In some cases, where immediate slide preparation is not available, the bone marrow may be aspirated into a tube containing a small amount of anticoagulant to impede clotting. The aspirate may later be filtered and submitted for histologic processing into a particle clot section. EDTA is the best anticoagulant to use because it introduces the least amount of morphologic artifact to the specimen (109). 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 and drawing marrow into tubes containing anticoagulant. Occasionally, a portion of an anticoagulated marrow aspirate is spun down to obtain a buffy coat, thereby concentrating the cellular elements. 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 also to make touch preparations from the biopsy, as outlined below, to allow cytologic examination of the bone marrow elements (109,115).
The bone marrow biopsy (Fig. 1.6) may be performed using the same skin incision 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 biopsy site away from the area where the aspiration was performed to avoid collection of a specimen with extensive artifact induced by the aspiration procedure (116). Use of a 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 farther. 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, decalcified, and processed for histologic examination (109,111).
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 approximately 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. Unstained smears should be retained for possible special stains if indicated (109,111).
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Figure 1.6. Bone marrow core biopsy. Histologic preparation of the bone marrow core biopsy following fixation and decalcification. The biopsy is stained with hematoxylin and eosin. This preparation allows for optimal evaluation of bone marrow cellularity and interaction of bone marrow cells with bony trabeculae, and is helpful in evaluating extrinsic features such as metastatic tumor or fibrosis in the marrow. Panel A (low power) shows bony spicules and marrow in section of bone marrow core biopsy. Panel B (high power) shows morphologic detail of hematopoietic tissue within the section. |
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 and preparation artifacts. An overall impression of the cellularity may be given (i.e., cellular or paucicellular). More accurate evaluation of bone marrow cellularity requires examination of a bone marrow biopsy or particle clot section, although the biopsy represents a tiny fraction of the total marrow and may also be subject to sampling error (109,111,117). The stained aspirate smear will have a central zone of dark marrow particles and stroma surrounded by a thinner area of dispersed bone marrow cells and red cells (Fig. 1.5). 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 (109).
The aspirate smear allows cytologic examination of the bone marrow cells. A minimum of 500 nucleated cells should be evaluated under oil immersion magnification in most marrows. 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 due to spreading artifact. This is usually in the dispersed cell zone adjacent to the spicule. It should be noted that spicules may be absent in pediatric marrows, and marrow will appear as a relatively uniform dispersion of cells. 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 (111). Results of differential counts from sternal bone marrow aspirate smears obtained from 12 healthy men at the University of Utah are presented in Table 1.5 as an example of bone marrow differential count reference ranges. The proportions of each cell type and progression of the maturational sequence for myeloid and erythroid elements 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 1.6) (109,111,118). 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 (119,120,121). Plasma cells are rare in the marrow of infants and children. Lymphocytes are much less numerous in adult bone marrows, usually making up <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 (77,122).
During the first month of life, erythroid cells are prominent because of high levels of erythropoietin (123); 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 element within the bone marrow, and more mature cells are also 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 or allergies may increase the bone marrow eosinophil count. Megakaryocytes constitute the least abundant cell type seen in the bone marrow, usually making up <1% of the cells (111).
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, and fat cells. In children, osteoblasts and osteoclasts may be seen, although these cells are rare in adults and their presence may indicate metabolic bone disease (109,111). Normally, these other cells make up <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 (124,125) or storage disorders (109,111).
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 or zinc formalin. The bony core will require decalcification before histologic processing. The fixed materials are processed and embedded in paraffin or plastic, and sections are made for examination. The bone marrow biopsy and clot sections are stained with either hematoxylin and eosin or Giemsa stains for morphologic examination (109,111) (Fig. 1.6).
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Table 1.5 Differential Counts of Bone Marrow Aspirates from 12 Healthy Men |
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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 (117) 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 (109,111,117). 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 (117). The bone marrow biopsy section provides the best representation of the bone marrow and its anatomic relationships, such as normal localization of immature myeloid cells adjacent to bony trabeculae. 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 relationships with bone, blood vessels, and other stromal elements. In particular, cellularity estimations may be falsely elevated secondary to collapse of the normal stromal network in a clot section (111,113).
In addition to providing information about the anatomic distribution and relationships of hematopoietic cells, the bone marrow biopsy is useful for evaluation of infiltrative processes such as carcinoma, lymphoma, other tumors, granulomatous inflammation, and fibrosis (109,111). 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 (115). In addition, evaluation of other elements, such as bony trabeculae, blood vessels, and stroma, requires a biopsy specimen.
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Table 1.6 Changes in Differential Counts of Bone Marrow with Age |
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Special Stains
Several special stains may be performed on peripheral blood smears, bone marrow aspirate smears, bone marrow touch preparations, and bone marrow biopsy materials and will provide additional information about the cell lineage beyond what is obtained by standard staining with Giemsa or hematoxylin and eosin stains. Special stains generally fall into two categories: Cytochemical stains that use cellular enzymatic reactions to impart staining and immunocytochemical stains that identify cell-specific antigen epitopes. These stains are particularly useful in characterization of primary hematologic or metastatic malignancies.
Cytochemical Stains
Cytochemical stains are extremely useful in the diagnosis and classification of acute leukemias. They allow identification of myeloid and lymphoid acute leukemias, as well as providing the basis for subclassification of the acute myeloid leukemias by the French-American-British (FAB) criteria and the World Health Organization (WHO) classification (126,127,128). 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.
Myeloperoxidase
Primary granules of neutrophils and secondary granules of eosinophils contain myeloperoxidase. Monocytic lysosomal granules are faintly positive. Lymphocytes and nucleated red blood cells lack the enzyme (129). Staining is due to oxidation of 3-amino-9-ethylcarbazole (130) or 4-chloro-1-naphthol (131) substrates by myeloperoxidase in the cell to form a brown-colored precipitate.
The myeloperoxidase enzyme is sensitive to light, and smears should be stained immediately or sheltered from light. Enzymatic activity in cells may diminish 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. 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 (pseudoperoxidase or Lepehne reaction). An antibody to myeloperoxidase is available for both flow cytometric analysis and immunohistochemical staining in fixed tissue sections (132,133).
Sudan Black B
Sudan black B stains intracellular lipid and phospholipids. 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. 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 (129).
Specific (Naphthol AS-D Chloroacetate) Esterase
The specific (naphthol AS-D chloroacetate) esterase stain, also called the Leder stain, is used to identify cells of the granulocytic series (129). It does not stain lymphocytes and monocytes. Because of enzymatic stability in formalin-fixed, paraffin-embedded tissues, this stain is extremely useful for identifying granulocytes and mast cells in tissue sections and is particularly helpful in diagnosis of extramedullary myeloid tumors (granulocytic sarcoma, chloroma) of blasts found in tissues (134). The cellular esterase enzyme hydrolyzes the naphthol AS-D chloroacetate substrate (135). 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 that may give rise to false-negative staining results.
Nonspecific (α-Naphthyl Butyrate or α-Naphthyl Acetate) Esterases
Nonspecific (α-naphthyl butyrate or α-naphthyl acetate) esterase stains are used to identify monocytic cells but do not stain granulocytes or eosinophils (135,136). Mature T lymphocytes stain with a characteristic focal, dotlike pattern. The stain also reacts with macrophages, histiocytes, megakaryocytes, and some carcinomas. The α-naphthyl butyrate stain is considered to be more specific, although slightly less sensitive, than the α-naphthyl acetate stain (136). Differential staining with the different esterases is seen in megakaryoblasts, which do not stain with the α-naphthyl butyrate, but stain with the α-naphthyl acetate substrate (137).
Terminal Deoxynucleotidyl Transferase
Terminal deoxynucleotidyl transferase (TdT) is an intranuclear enzyme that catalyzes the addition of deoxynucleotide triphosphates to the 3′-hydroxyl ends of oligonucleotides or polydeoxynucleotides without need for a template strand (138). TdT is found normally in the nucleus of thymocytes and immature lymphoid cells within the bone marrow, but is not found in mature lymphocytes, and it is a useful marker in identifying acute lymphoblastic leukemias and lymphomas (129,139). TdT activity is found in approximately 90% of acute lymphoblastic leukemias as well as in a small subset of acute myelogenous leukemias (140). TdT levels may be measured biochemically, by cytochemical staining with an immunofluorescent technique, by flow cytometry after permeabilization of freshly collected cells, or by immunohistochemical methods (138,141). Indirect immunofluorescent staining is very sensitive and may be applied to air-dried samples several weeks after collection (142). Immunohistochemical methods of TdT detection are useful in paraffin-embedded tissue sections (143).
A panel of myeloperoxidase (or Sudan black B), α-naphthyl butyrate esterase (or double esterase stain), and TdT staining is often used to characterize acute leukemias. More precise lineage assignment may be provided by flow cytometric analysis if fresh bone marrow or peripheral blood is available (139). However, because the FAB and WHO classifications of the acute myelogenous leukemias are based on the cytochemical staining pattern of the blasts (126,127,128), these stains are usually performed even when immunophenotypic analysis is performed. In addition, cytochemical stains may be retrospectively performed when 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 as a screening test to differentiate chronic myelogenous leukemia from leukemoid reactions and other myeloproliferative disorders (144). The LAP score is usually performed using the Kaplow procedure (145). 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 1.7). Patients with chronic myelogenous leukemia have low LAP scores (usually between 0 and 13). Paroxysmal nocturnal hemoglobinuria and some myelodysplastic syndromes may also 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 (144). There is rapid loss of alkaline phosphatase activity in samples drawn in EDTA anticoagulant (145). The test is optimally performed on fresh capillary blood fingerstick smears or on blood anticoagulated with heparin and should be performed within 48 hours after collection of the sample. The blood smears may be held in the freezer for 2 to 3 weeks with little loss of activity.
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Table 1.7 Conditions Associated with Abnormal Leukocyte Alkaline Phosphatase (LAP) Scores |
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Acid Phosphatase
Acid phosphatase is found in all hematopoietic cells, but the highest levels are found in macrophages and osteoclasts. 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 (146) and osteoclasts. Several methods of measuring TRAP activity have been described, but one using naphthol AS-BI phosphoric acid coupled to fast garnet GBC is reliable and reproducible (147). Not all cases of hairy cell leukemia stain for TRAP, and staining intensity may be variable (148). Positive TRAP staining may also be seen in some activated T lymphocytes, macrophages, some histiocytes (such as Gaucher cells), mast cells, and some marginal zone lymphomas (144,149). TRAP staining may also be detected by immunohistochemical methods in fixed tissue sections (149).
Periodic Acid-Schiff
The periodic acid-Schiff (PAS) stain detects intracellular glycogen and neutral mucopolysaccharides, which are found in variable quantities in most hematopoietic cells (150). PAS staining is seen in blasts of both acute lymphoblastic and acute myelogenous leukemias, although there is great variability between cases (129,139). Erythroleukemias demonstrate an intense diffuse cytoplasmic positivity with PAS, which may be helpful in diagnosis (128). In addition, PAS staining is very useful in demonstrating the abnormal glucocerebrosidase accumulation in Gaucher disease (151).
Iron
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 (152). The stain is used to identify iron in nucleated red blood cells (sideroblastic iron) and histiocytes (reticuloendothelial iron) or to identify Pappenheimer bodies in erythrocytes. Normally, red cell precursors contain one or more small (<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 (128). The stain is best used on bone marrow aspirate smears but can also be used on blood films and aspirate clot tissue sections. Decalcification of the bone marrow core biopsy may lead to loss of iron from the cells, leading to a false impression of low iron.
Toluidine Blue
Toluidine blue specifically marks basophils and mast cells by reacting with the acid mucopolysaccharides in the cell granules to form metachromatic complexes. Malignant mast cells or basophils may have low levels of acid mucopolysaccharides and may not react with this stain (129).
Immunocytochemical Stains
Immunocytochemical staining is based on the use of an antibody that recognizes a specific antigenic epitope on a cell. There is a high level of specificity. In general, these stains may be applied to blood smears, bone marrow aspirates, cellular suspensions, or tissue sections. Not all antibody preparations are equally effective on all types of specimens, and staining procedures may vary depending on the specimen type. A wide variety of antibodies specific to hematopoietic cellular antigens is available commercially. Some of the newer antibodies have replaced classical cytochemical stains and may be useful on older or fixed specimens.
Immunocytochemical staining of fresh blood or bone marrow cell suspensions or cell suspensions from tissues and analysis by flow cytometry are becoming increasingly common in clinical laboratories (153,154). 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 (153). In many cases, particularly in the acute leukemias, the flow cytometric analysis of an acute leukemia provides important prognostic information that is not available through cytochemical staining (154,155,156) and is useful in detection of minimal residual disease (157,158,159). Clinical and technical aspects of flow cytometric analysis of hematologic tumors are covered in detail in Chapter 2.
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 (160). 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 (161). Recently, several automated immunostaining instruments have become available that allow highly reproducible results and require less technician time and expertise for highly reproducible staining (162).
Other Laboratory Studies
Cytogenetic Analysis
Many hematologic malignancies and premalignant conditions are associated with specific cytogenetic changes (128,163,164,165). 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 (166). Cytogenetic analysis has become important in diagnosing hematologic disorders, identifying specific prognostic subgroups, and monitoring for progression of disease or residual disease after therapy, and is integral to the most current classification of hematologic malignancies, such as the WHO classification (128,167,168). Both standard chromosomal preparations and fluorescent-labeled in situ hybridization techniques may be used for cytogenetic analysis of chromosomal changes (110,169). Further details about cytogenetic techniques and analysis are provided in Chapter 3.
Molecular Genetics
In addition to standard morphologic analysis and cytogenetics, technology has been developed that allows analysis of molecular changes in hematologic malignancies (167,168,170). By use of Southern blot and polymerase chain reaction (PCR) techniques, hematopoietic proliferations may be studied for genetic alterations associated with development of malignancy (171). Molecular genetic analysis was initially used to identify monoclonality in lymphoid neoplasms by identifying either immunoglobulin (B-cell) or T-cell-receptor gene rearrangements (172,173). 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 (172). 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. Common tests include the BCR-ABL1 translocations seen in chronic myelogenous leukemia and acute leukemia and used to monitor efficacy of treatment (167,174), BCL2 translocations characteristic of follicular lymphomas (175), the t(15;17) translocation associated with promyelocytic leukemia (176), and JAK2 translocations associated with myeloproliferative disorders (177,178). As molecular characterization and genetic profiling of specific hematologic disorders expand, such as through microarray analysis (179), it may be anticipated that more PCR and molecular tests will be developed. 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), may allow for quantification of low levels of transcripts to allow monitoring of disease status, and can lead to more rapid test completion (especially with PCR-based testing) (174,176). Molecular tests are most useful when a known specific entity is being tested for or in monitoring residual disease as they do not provide effective screening capability for additional genetic alterations that may affect prognosis, as does conventional cytogenetic analysis (110,163,165).
The high degree of sensitivity makes molecular testing, particularly by PCR or in situ hybridization, 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 assays can make use of formalin-fixed materials with sensitivity similar to that of fresh or frozen materials (180,181). 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 4.
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 polycythemic disorders 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 adults older than age 75) (182) and tends to be higher in women. People with liver disease, carcinomas, or other serious diseases may have a normal to low ESR because of an inability to produce the acute-phase proteins (183).
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 (184,185,186). The ESR is a poor screening test in asymptomatic individuals, detecting elevations in 4 to 8% of normal adults and, hence, should not be used to screen asymptomatic people for disease (186). The test is probably best used in the clinical scenario of a patient with vague complaints to aid in the clinical decision to undergo further testing or as a tool to follow the clinical disease course in temporal arteritis, rheumatoid arthritis, polymyalgia rheumatica, or lymphomas (186).
The ESR is measured by the Westergren or Wintrobe method or by a modification of these tests (187). 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 patient anemia. Several technical variations to the method of ESR determination have been introduced, including micromethods, sedimentation at a 45-degree angle, and the zeta sedimentation rate. The zeta sedimentation rate measures erythrocyte packing in four 45-second cycles of dispersion and compaction in capillary tubes. This requires a special instrument, the Zetafuge (Coulter Electronics, Hialeah, FL), but gives reproducible results on very small amounts of blood and is not affected by patient anemia (188).
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 (such as plasma cell dyscrasias) (189). 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 (190). However, plasma viscosity is used more rarely than ESR, probably reflecting clinical familiarity with the latter test. Like ESR, plasma viscosity may increase with age (189). Direct measurement of acute-phase proteins, such as C-reactive protein, may also be used to monitor the course of inflammatory diseases and cardiac risk (191,192). However, these tests are usually more expensive than ESR determinations and may not provide sufficient additional clinical information to justify the added expense (193). Whole blood viscosity measurements are of limited clinical use 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.
Blood Volume Measurement
In most cases, the total number of erythrocytes is closely related to the red cell concentration of the blood or Hct. However, blood volume may not reflect erythrocyte concentration, including immediately after severe hemorrhage, severe dehydration, or overhydration. To accurately assess the blood volume in these patients, plasma volume or red cell mass or volume must be determined (38,194). The plasma volume is measured by dilution methods. A substance that is confined to the intravascular plasma compartment, such as Evans blue dye (195), 131I-labeled albumin, or radioactive indium-labeled transferrin, 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, but corrections must be made because the label is gradually removed from the circulation into the extravascular space, leading to errors of 10% or more in plasma volume determinations (196). Plasma volume may also be estimated from red cell volume (197).
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. Biotinylated cells may also be used (198). The red cell volume is then calculated by the dilution of the labeled cells over time using the following formula:
Red cell volume = cpm of isotope injected/cpm/ml red cell concentration per unit volume in sample
Usually, the measurements are made after a 15-minute interval, although longer periods may be needed with high blood viscosity due to high Hcts to ensure complete labeled cell mixing. Total red cell volume measurements must be corrected with splenic enlargement secondary to sequestration of the labeled cells within this organ. Red cell volume may also be calculated from the total plasma volume and measured Hct by means of the following equation:
Red cell volume = Hct × plasma volume/100 - Hct
Total plasma volume may be useful in monitoring fluid and blood replacement. Red cell volume measurements are used to document true polycythemia, although some authors advocate the use of erythropoietin levels and red cell colony growth as less invasive surrogate tests for red cell volume or red cell mass measurements (38). Total blood volume may be calculated from the sum of total red cell volume and plasma volume measurements.
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