Automated complete blood count (CBC)

INTRODUCTION — This topic reviews the automated complete blood count (CBC), including definitions of the parameters, their clinical significance and relevance, and the methods used to determine these values.

Separate topics discuss evaluations of abnormal findings on the CBC and the peripheral blood smear.

●Counts too low

•Red blood cells (RBCs) – (See "Approach to the child with anemia" and "Diagnostic approach to anemia in adults".)

•Platelets – (See "Approach to the child with unexplained thrombocytopenia" and "Diagnostic approach to thrombocytopenia in adults".)

•White blood cells (WBCs) – (See "Overview of neutropenia in children and adolescents" and "Approach to the adult with unexplained neutropenia".)

•All three – (See "Approach to the adult with pancytopenia".)

●Counts too high

•RBCs – (See "Diagnostic approach to the patient with erythrocytosis/polycythemia".)

•Platelets – (See "Approach to the patient with thrombocytosis".)

•WBCs – (See "Approach to the patient with neutrophilia".)

●Blood smear abnormalities – (See "Evaluation of the peripheral blood smear".)

DEVELOPMENT OF CELL COUNTING METHODS — Modern automated hematology instruments use the following methods to count cells, alone or in combination [1]:

●Impedance-based methods based on the Coulter principle (changes in electrical current induced by blood cells flowing through an electrically charged opening)

●Optical methods such as light scatter

●Fluorescence-based methods

This allows enumeration and evaluation of blood cells with great accuracy, precision, and speed.

Manual counting — During the first half of the 20th century, the CBC was performed using manual techniques that were laborious and imprecise:

●Cell counting – Blood cell counts (red blood cells [RBCs], white blood cells [WBC], platelets) were generated manually by spreading blood from a diluted sample onto a slide and performing a Wright-Giemsa stain, on which cells were counted using a ruled counting chamber (hemocytometer). Reticulocytes can be counted as a percentage of total RBCs when stained with a special stain. (See "Evaluation of the peripheral blood smear", section on 'Reticulocytes'.)

●Hemoglobin concentration – Hemoglobin was analyzed colorimetrically by the cyanmethemoglobin method.

●Hematocrit – The hematocrit (HCT; also called packed cell volume) was measured by high-speed centrifugation of a column of blood, either in a specially designed tube (the Wintrobe tube) (picture 1), or in a sealed microcapillary tube ("spun" hematocrit, often obtained by fingerstick blood collection) (picture 2).

●WBC differential – The WBC differential was determined from a stained blood smear by counting 100 to 200 WBCs and enumerating neutrophils, eosinophils, basophils, lymphocytes, monocytes, and immature cells.

●RBC indices – In 1932, Wintrobe developed a set of indices that estimated RBC size and hemoglobin content. All were calculated manually from the RBC count, hemoglobin concentration, and HCT; the table (table 1) summarizes the calculations for MCV (mean corpuscular volume), MCH (mean corpuscular hemoglobin), and MCHC (mean corpuscular hemoglobin concentration).

Electrical impedance (Coulter aperture) — Wallace Coulter patented a device in 1953 that used electrical impedance to count RBCs and WBCs as they passed through an aperture [2].

The method for counting works as follows:

●Blood is diluted in a current-conducting solution such as isotonic saline.

●The cell suspension is drawn by a vacuum through a small aperture positioned between two sensing electrodes connected across a direct current potential.

●As each cell passes through the aperture, the resistance between the electrodes increases.

●The cell produces a momentary increase in impedance in electrical conduction (because it is non-conductive), resulting in an electrical pulse.

●The number of pulses indicate the cell count.

●The amplitude of each pulse is proportional to each cell's volume.

A cell count on an appropriately diluted whole blood sample yields the RBC count; the WBCs do not appreciably affect the count because they are three orders of magnitude less abundant than RBCs. RBC lysing reagents are then added to the sample to produce an accurate WBC count.

Upper and lower pulse height thresholds can be selected such that only particles within a specific volume range are counted. The lower pulse height threshold can be set to eliminate platelets (platelet volume is approximately 9 fL, one-tenth the volume of RBCs).

Multi-channel instruments — In the 1960s, the first multi-channel automated hematology instruments appeared. These instruments revolutionized the CBC as a laboratory test, enabling high throughput, low cost, and fast turnaround time for RBC and WBC parameters. Platelet counts (PLTs) were not a part of the first automated CBC since the aperture resolution on the early instruments could not adequately separate platelets from RBCs.

Anticoagulated whole blood samples were aspirated into the apparatus and automatically aliquoted and diluted into RBC and WBC counting chambers (baths) for cell counting and sizing with impedance apertures, and into a spectrophotometric cuvette for hemoglobin determination using the cyanmethemoglobin method. Coincidence correction (to correct for two or more cells passing through the aperture that are counted as one cell) and calculations of RBC indices were performed automatically.

Since these machines measure the RBC count and MCV directly, they can calculate the HCT using the Wintrobe formula, which calculates HCT based on the RBC count and the MCV (table 1).

Light scattering — Light scattering technology for cell counting was introduced in the 1970s.

In this technique, cells are hydrodynamically focused in a flow cell and illuminated by a narrow beam of laser light. The light scattered by each cell is captured by a photodetector and converted to an electrical pulse. The total number of pulses is proportional to the cell count; pulse amplitude is proportional to cell volume. The number of pulses within a predetermined size range (30 to 180 fL for RBCs, 0 to 20 or 0 to 30 fL for platelets) yields the corresponding RBC and platelet counts. WBCs are counted in the same way after the RBCs are lysed.

Analysis of light scatter from multiple angles provides better separation of WBCs from nucleated RBCs and RBCs that fail to lyse (a problem with neonatal samples, which have difficult-to-lyse RBCs). Some instruments determine both optical and impedance WBC counts and compare them as a means of better eliminating interference from nucleated or unlysed RBCs.

Attempts to automate the WBC differential started with image analysis instruments in the 1970s and progressed to a flow-through automated differential [3].

RBC evaluation with light scattering technology requires that the RBCs be put into a hypotonic solution so that liquid enters the cells and they become spherical; then they are fixed. Scattered light is measured at low forward angle (0 to 3 degrees) and high forward angle (5 to 15 degrees), yielding pulses proportional to RBC size and refractive index, respectively. The refractive index is proportional to the hemoglobin content of the cell, allowing hemoglobin content to be directly determined for individual RBCs. This generates an RBC "cytogram" that produces statistics for RBC volume (MCV) and hemoglobin concentration.

RBC volume and hemoglobin concentration histograms and smoothed curves can be constructed, and the mean, standard deviation (SD), and coefficients of variation (CV) electronically calculated, yielding the various RBC indices including the MCV and red cell distribution width (RDW). These are calculated differently on different CBC instruments (table 2).

General features of modern instruments — Modern analyzers include an automated sampling mode to minimize contact between the operator and the blood. Blood is aspirated from closed tubes with automated bar code readability for positive sample identification.

A built-in mixing system ensures sampling of a well-mixed specimen, and the instruments usually include a sample aspiration monitor to aid in detecting clots and determining that the correct sample volume has been aspirated.

●Tests performed on all samples – Some tests are routinely performed on all samples:

•Hemoglobin, red blood cell (RBC) count, and RBC indices

•White blood cell (WBC) count

•Platelet count

●Tests that can be requested – Other tests such as the reticulocyte count and WBC differential are performed only when requested, saving reagent costs.

•Ideally, if a reticulocyte count is requested, it should be ordered at the same time as the CBC, but it can be added on later if the sample is not too old (no more than 48 hours after collection). (See 'Reticulocytes' below.)

•Advances in various methods for the WBC differential allow distinction among different types of WBCs and identification of abnormal WBCs. (See 'WBC differential' below.)

Automated slide makers also are available that can create and stain blood films as directed by laboratory-programmed algorithms, which consider both the results of specific parameters as well as the presence of instrument flags. The combination of automated hematology analyzers coupled with automated slide makers and stainers have resulted in automated hematology work cells with high throughput and increased efficiency [4]. Image analysis has further increased efficiency, with identification of different cell types via machine learning algorithms [5,6].

Modern instruments perform curve fitting, multi-dimensional population cluster discrimination, and moving averages for quality control. Computer memory can be used to store and sort samples and to maintain sample scatterplots in "list" mode for reanalysis. Certain parameters that are similar are named differently depending on each instrument manufacturer. The table summarizes which parameters are available on which instrument (table 2).

Because each instrument determines some parameters differently, it is possible to get different results if the same sample is analyzed on different instruments. Some of these differences could potentially be significant, depending on the patient's condition. In healthy patients, there will be no clinically significant differences.

SAMPLE COLLECTION AND PROCESSING

Collection

●Phlebotomy – Collection of the blood samples should be performed by a trained phlebotomist or experienced member of the health care team. The table summarizes sources of spurious results on the CBC from phlebotomy-related causes (table 3).

It is very common to observe hemolyzed blood samples due to small gauge needles or tourniquets that are too tight, leading to shearing of red blood cells (RBCs). Hemolysis during phlebotomy spuriously affects the RBC count and hemoglobin (Hb). Collection of a blood sample proximal to an intravenous line will result in a dilute blood sample with spurious results.

●Collection tube – Samples to be used for a CBC should be collected into a tube with an anticoagulant solution such as ethylenediaminetetraacetic acid (EDTA) or citrate. EDTA may cause platelet clumping or satellitism in some patients, leading to pseudothrombocytopenia [7]. Platelet aggregates can also falsely increase the white blood cell (WBC) count [8]. This can be identified by reviewing the edge of the blood smear and/or repeating the test using a tube with citrate. (See "Diagnostic approach to thrombocytopenia in adults", section on 'Pseudothrombocytopenia'.)

The ratio of anticoagulant in a tube to blood is carefully calibrated; underfilling and overfilling of the tube should be avoided.

•Underfilling of the tube can lead to overexposure to the anticoagulant (eg, EDTA).

•Overfilling of the tube can reduce exposure to the anticoagulant with possible clotting, producing inaccurate results.

Storage

●Storage temperature – Blood samples should be kept at room temperature if the specimen will be analyzed within 24 hours of collection. Samples should be refrigerated if the analysis is to occur up to 72 hours after collection [9].

Freezing of blood samples must be avoided; freezing will cause cells to lyse, leading to inaccurate values. Blood samples that are exposed to heat will cause RBCs to fragment, producing significant anisocytosis and bizarre microcytes resembling blood samples seen in burn victims.

●Storage duration prior to CBC testing – We recommend that samples >72 hours old not be used for CBC testing. Samples >72 hours old can show spurious results such as an elevated mean corpuscular volume (MCV) and elevated mean platelet volume (MPV) due to prolonged EDTA exposure with swelling of RBCs and platelets (algorithm 1). Alterations in the percentages of different WBC subsets can also be seen (table 3).

Blood films should be prepared within eight hours of sample collection. With longer storage, cells become pyknotic with altered morphology that can lead to spurious identification dysplasia in WBCs and platelets that is not actually present, as well as overcalling of RBC artifacts due to crenated RBCs.

Quality control

●Rule of threes – In a healthy individual, the rule of threes should be met (three times the RBC count equals the hemoglobin, and three times the hemoglobin equals the hematocrit). If the rule is violated, this suggests either:

•The results may be spurious (in an otherwise healthy individual).

or

•A true hematologic condition may be present.

●Blood smear artifacts – (See "Evaluation of the peripheral blood smear".)

RBC PARAMETERS

RBC count, Hb, and HCT — Red blood cell (RBC) parameters are summarized in the table (table 4) and include:

●RBC count – RBC count is the number of RBCs per microL of blood (number of RBCs x 10¹²/L). An elevated RBC count indicates polycythemia (reactive or neoplastic) or thalassemia. A decreased RBC typically indicates anemia. (See "Diagnostic approach to the patient with erythrocytosis/polycythemia" and "Diagnostic approach to anemia in adults" and "Approach to the child with anemia".)

●Hemoglobin – Hemoglobin (Hb) is the concentration of Hb in whole blood, in grams/deciliter (g/dL). Increased Hb may be due to polycythemia (reactive or neoplastic) or may occur with dehydration. A decreased Hb typically indicates anemia, although it may be caused by increased intravascular volume.

●Hematocrit – The hematocrit (HCT) is the packed spun volume of blood made up of RBCs, expressed as a percentage of total blood volume. It can be measured or calculated; the table provides the formula (table 1).

An increased HCT can indicate polycythemia (reactive or neoplastic) or dehydration; if calculated, an increased HCT may reflect a normal number of RBCs of increased size.

RBC indices — The RBC indices describe the volume and Hb content of the cells. These values generally do not vary in the same individual over time when performed in the same laboratory using the same instrument, and changes may warrant investigation, especially in individuals with anemia.

Different laboratories may use different hematology instruments, so it is best practice to use the same laboratory for following a patient's hematologic results over time.

MCV and RDW — MCV and RDW reflect the average RBC volume and the distribution of volumes, respectively.

●MCV – Mean corpuscular volume (MCV) is the average volume (size) of the RBCs. It can be measured or calculated. Anemia can be classified based on whether the MCV is low, normal, or elevated. (See "Diagnostic approach to anemia in adults", section on 'RBC indices'.)

●RDW – Red cell distribution width (RDW) is a measure of the variation in MCV, which is reflected in the degree of anisocytosis on the peripheral blood smear.

RDW is calculated; the RDW is the coefficient of variation (CV) or the standard deviation (SD) of the MCV distribution curve, depending on the instrument.

•Beckman Coulter instruments set the RDW as the CV of the MCV distribution curve.

•Some Sysmex instruments set the RDW as the SD of the curve.

Modern cell counting and sizing apertures have sufficient precision to function as "channelizers." As an example, Coulter instruments channel cells within the RBC and platelet aperture into 256 bins in the size range from 0 to 360 fL. RBCs are counted as particles with a volume within the range of 36 to 360 fL and classified into appropriate "channels" by size (related to the pulse height).

For instruments with pulse editing circuitry, aberrant pulses are excluded from these channels. A plot of frequency versus channel size allows the development of an RBC volume histogram, which can be smoothed into an RBC size distribution curve.

Interpretation of abnormal MCV and RDW — The RBC size frequency distribution curve typically has a symmetrical or Gaussian shape. The MCV and RDW are both directly derived from this curve.

●MCV – The MCV should not significantly vary in a healthy adult. Changes outside of a laboratory's reference interval or a change from a person's baseline that is >5 fL in the same laboratory using the same instrument should prompt an investigation, especially in a patient with anemia. The interpretation and evaluation of an abnormal MCV is discussed separately:

•Low MCV – (See "Microcytosis/Microcytic anemia".)

•High MCV – (See "Macrocytosis/Macrocytic anemia".)

●RDW – If all parameters of the CBC are normal, an abnormal RDW should not prompt a significant work-up; instead, a repeat blood draw and CBC can be performed.

The RDW is a calculated value, which inherently implies more variation. Some instruments use the SD and others use the CV of the MCV distribution curve; this will lead to dramatically different values depending on the instrument. (See 'MCV and RDW' above.)

The RDW is most useful in patients with microcytic anemia, where an elevated RDW is consistent with iron deficiency and a normal RDW is consistent with anemia of chronic disease/anemia of inflammation (ACD/AI). However, the RDW cannot replace a serum ferritin or iron studies or other testing in making this distinction. (See "Causes and diagnosis of iron deficiency and iron deficiency anemia in adults", section on 'Diagnostic evaluation' and "Anemia of chronic disease/anemia of inflammation", section on 'Diagnostic evaluation'.)

Abnormalities of this curve may indicate certain types of anemia or platelet clumping:

•Left shoulder (small cells) – A left "shoulder" extension to the curve, or failure of the curve to reach baseline on the left side (population of RBCs with smaller volumes), can indicate microspherocytes or schistocytes. In some cases, unusually large platelets or platelet clumps may be counted as small RBCs.

A distinct population of small RBCs can indicate a condition such as X-linked sideroblastic anemia (figure 1). (See "Sideroblastic anemias: Diagnosis and management", section on 'X-linked sideroblastic anemias'.)

•Right shoulder (large cells) – A right-sided shoulder usually corresponds to a population of extremely large RBCs or reticulocytes. A trailing population to the extreme right (MCV >200 fL) can indicate RBC agglutination (picture 3). (See "Cold agglutinin disease", section on 'When to suspect CAD'.)

•Abnormal shapes – All pulses in the RBC aperture are enumerated for the RBC count, even if they have an aberrant shape. However, RBCs with an aberrant shape generate a pulse height that is also aberrant, and this is edited out of the MCV computation. In some samples, up to 20 percent of counted RBCs may be excluded from the MCV calculation.

•High RDW – A high RDW implies a large variation in RBC sizes. A very elevated RDW can be seen in iron deficiency anemia, transfused anemia, myelodysplastic syndromes, and hemoglobinopathies, whereas a normal to slightly elevated RDW can be seen in thalassemia trait and ACD/AI. (See "Evaluation of the peripheral blood smear" and "Diagnostic approach to anemia in adults", section on 'RBC indices'.)

RDW has been proposed as a tool to distinguish iron deficiency (elevated RDW) from thalassemia trait (normal RDW) in samples with low MCV. However, thalassemia trait can also cause an elevated RDW. (See "Microcytosis/Microcytic anemia", section on 'RDW (size variability)'.)

MCH and MCHC — These parameters reflect RBC hemoglobin content and hemoglobin concentration. In the absence of anemia, a slightly abnormal MCH or MCHC should not prompt further evaluation.

●MCH – Mean corpuscular hemoglobin (MCH) is the average hemoglobin content in the population of RBCs. It is calculated from the hemoglobin value and RBC count, as shown in the table (table 1). A low MCH indicates decreased hemoglobin content per cell and is typically reflected in hypochromia on the peripheral blood smear. This may be seen in iron deficiency and disorders of globin synthesis.

●MCHC – Mean corpuscular hemoglobin concentration (MCHC) is the average hemoglobin concentration per RBC, in grams/dL. It is calculated from the hemoglobin and the hematocrit (table 1). MCHC does not significantly vary in healthy individuals and can be used to "match" samples if there is a question of a sample mix-up.

Low and high MCHC values are helpful in classifying anemias. Very low MCHC values are typical of iron deficiency anemia, and very high MCHC values typically reflect spherocytosis or RBC agglutination. Examination of the peripheral blood smear is helpful in distinguishing these findings. (See "Evaluation of the peripheral blood smear", section on 'Red blood cells' and "Diagnostic approach to anemia in adults", section on 'Anemia definitions'.)

•Low MCHC – A low MCHC can be seen with:

-Iron deficiency.

-Hyperglycemia, which causes an osmotic imbalance and increase in cell water.

•High MCHC – An increased MCHC can be seen with spherocytosis or as a spurious result; the table summarizes sources of interference (table 3):

-Spuriously low RBC count due to a cold agglutinin, which causes several agglutinated RBCs to be counted as a single RBC.

-Spuriously increased hemoglobin, which can result from a very high WBC count, lipemia, a monoclonal protein, or a hemoglobin-based oxygen carrier (HBOC), all of which produce turbidity in the colorimetric method used to determine hemoglobin. (See "Oxygen carriers as alternatives to red blood cell transfusion".)

RETICULOCYTES

Reticulocyte count — Automated counting of reticulocytes became available in the early 1990s, significantly improving the precision of reticulocyte counts over manual counting. All of the major automated instruments are capable of generating a reticulocyte count, but often this must be requested specifically, because it requires additional reagents and generates additional costs. (See 'General features of modern instruments' above.)

Reticulocytes are identified by the absence of a nucleus and the presence of measurable amounts of RNA. RNA is often detected with a fluorescent dye that binds nucleic acids, such as thiazole orange. This can yield a reticulocyte count as well as an immature reticulocyte fraction (IRF; sum of the reticulocytes with medium and high fluorescence) [10,11]. Reference intervals for the IRF are dependent on each method used.

Automation provides an absolute reticulocyte count (the number of reticulocytes per microL of blood), as well as the reticulocyte percentage, as a percent of all red blood cells (RBCs). (See "Diagnostic approach to anemia in adults", section on 'Reticulocyte count'.)

Reticulocyte indices — Similar to RBCs, reticulocytes can be analyzed for mean corpuscular volume (MCVr), hemoglobin content (CHr), and other parameters. Different instruments use various proprietary reticulocyte indices.

The main use proposed for reticulocyte indices is as an early indicator or screening test for iron deficiency. (See "Causes and diagnosis of iron deficiency and iron deficiency anemia in adults", section on 'Findings on CBC' and "Anemia of chronic disease/anemia of inflammation", section on 'Tests in development'.)

The specific parameters measured depend on the instrument, as summarized in the table (table 2):

●Siemens Advia – These instruments stain reticulocytes with Oxazine 750 and measure light adsorption [12]. Other reticulocyte parameters are reported in sphered reticulocytes (eg, MCVr, CHCMr, HDWr, CHr, CHDWr) similar to those for the sphered mature red blood cells (RBCs) [13]. (See 'RBC indices' above.)

High, medium, and low fluorescent fractions are also calculated.

The hemoglobin content of reticulocytes (CHr), measured on the Siemens Advia instruments, appears to be a sensitive and specific indicator in detecting iron deficiency, monitoring iron therapy in patients with kidney failure on chronic dialysis, and monitoring for the advent of iron deficiency following treatment with erythropoietin [14,15].

Another reticulocyte index, the mean reticulocyte volume (MCVr), has been proposed to be helpful in diagnosing iron deficiency, monitoring response to treatment of nutritional anemias, an early indicator of erythropoiesis after bone marrow transplantation, and as a tool in the evaluation of erythropoietin abuse in athletes [12,16-18].

●Abbott – Some Abbott instruments use a proprietary fluorescent RNA dye called CD4K540 [19]. This dye is 10-fold as bright as Auramine O and fluoresces at 530 nm (green). A plot of seven-degree light scatter versus fluorescence separates platelets from RBCs and separates very highly fluorescent non-viable WBCs and nucleated RBCs from reticulocytes. The combined RBCs and reticulocytes are gated; a histogram of the mature RBC plus reticulocyte fluorescence is analyzed with a valley-finding algorithm to separate and enumerate the reticulocytes. The IRF is identified arbitrarily by setting a threshold 30 channels above the RBC/reticulocyte threshold.

●Beckman-Coulter – These instruments stain blood with new methylene blue, followed by clearing with a hypotonic solution [20]. RBCs are measured for volume, conductivity, and laser light scatter; the latter signal is proportional to the residual RNA within the red cell. Reticulocyte volume is measured, and reticulocytes with the most RNA (corresponding to the highest light scatter regions) are counted. The DxH800 captures five laser light scatter measurements for each cell, in contrast to one light scatter measurement for the LH750/780. The IRF is then the ratio of those reticulocytes with the most RNA over the total number of reticulocytes.

●Sysmex – Reticulocytes and the immature reticulocyte fraction (IRF) are determined on Sysmex instruments using a polymethine dye that stains reticulocyte RNA. Graphing fluorescence versus forward light scatter separates mature RBCs from reticulocytes. The IRF is the sum of the moderately and highly fluorescent reticulocytes (MFR+HFR). A combination of RBC and reticulocyte parameters on the Sysmex Xn has been proposed as a screening method for iron deficiency anemia and hereditary RBC diseases with a sensitivity and specificity of 95.2 and 99.9 percent, respectively, using a classification and regression tree analysis [21].

PLATELET PARAMETERS

Platelet count — The platelet count is the number of platelets per microL of blood (or number of platelets x 10⁹/L).

●Evaluation – Evaluation of thrombocytopenia and thrombocytosis is discussed in separate topic reviews:

•Low – A decreased platelet count (thrombocytopenia) may reflect platelet destruction, sequestration, or ineffective thrombopoiesis. If platelet clumping occurs (eg, in association with an ethylenediaminetetraacetic acid [EDTA]-dependent platelet agglutinin), the clumps may be so large (picture 4) that they exceed the threshold for platelet size and lead to spuriously low platelet counts (table 3). This problem can be circumvented by using a blood sample collected in a tube using citrate or heparin as the anticoagulant. (See "Neonatal thrombocytopenia: Etiology" and "Causes of thrombocytopenia in children" and "Diagnostic approach to thrombocytopenia in adults".)

•High – An elevated platelet count (thrombocytosis, also called thrombocythemia) may be seen in reactive and neoplastic conditions. (See "Approach to the patient with thrombocytosis".)

●Methodologies – Platelets may be counted using manual counting, impedance, light scatter, fluorescence, immunologic methods (flow cytometry), and digital image analysis [22]. Automated platelet counting primarily uses impedance, optical scatter, and/or fluorescence, although immunologic methods are available on some instruments. Image analysis is a developing technology.

When instrument platelet flags occur, it is important to verify the platelet count by estimation from a peripheral blood smear.

•Manual counting – In an area of the blood smear where red blood cells (RBCs) barely touch each other, the number of platelets per 100X field multiplied by 20 x 10⁹/L gives an estimate of the platelet count. If platelet clumps are seen, it is likely that the platelet count is spuriously low and that the white blood cell (WBC) count is spuriously high. (See "Evaluation of the peripheral blood smear", section on 'Platelets'.)

•Impedance – On instruments that use impedance-based platelet counting, platelets are enumerated as particles in a given range (eg, from 2 fL to 30 fL). A raw data frequency curve is generated from a channelized histogram of platelet volumes and is extrapolated to 70 fL. In contrast to the Gaussian distribution typical of RBCs, platelets show a lognormal or skewed distribution. A mean platelet volume (MPV) is calculated from a fitted curve of the platelet volume distribution curve to yield a geometric mean.

Inspection of the platelet histogram can aid in determining inaccuracies in the platelet count; these are typically "flagged" by the instrument. A peak at volumes <2 fL suggests cytoplasmic fragments or interference by electronic noise, while a failure to return to baseline at volumes >20 to 30 fL indicates interference by microcytic RBCs or failure to include giant platelets in the count. These limitations are often cited; however, some automated hematology analyzers have shown better correlation of their impedance platelet counts with the reference method compared with optical methods [23].

Different instruments use different techniques to detect interferences that can alter the platelet count. Some instruments enumerate particles between 2 to 3 fL (on the low end) and 20 to 30 fL (on the high end) as platelets and monitor the percent of total particles near the lower counting threshold (1 to 2 fL) and the upper counting threshold (20 to 30 fL). When the number of particles in these regions exceeds certain limits, "flags" are generated.

•Optical scatter – An optical platelet count and volume can be determined from two-dimensional analysis of low and high angle light scatter measurements. These two light scatter measurements are related to platelet volume (size) and refractive index (density) [24]. Platelet volume histograms and smoothed curves can be constructed from this plot and used to determine the platelet count, mean platelet volume (MPV) and standard deviation (SD) or coefficient of variation (CV) of the volume (platelet distribution width [PDW]). Interference by microcytic RBCs, RBC fragments, and WBC cytoplasmic fragments is eliminated since these particles have a different refractive index than platelets and are not counted.

In some Abbott instruments, platelets are first plotted using scattering angles of 90 degrees versus 7 degrees. Thresholds are set to isolate the platelets from large and small cells and fragments on either side of the histogram. A second histogram is constructed along the main body of this refined platelet population to determine the upper threshold separating platelets from microcytic RBCs.

Some Sysmex instruments provide a platelet count using impedance, optical scatter, and fluorescence, and they use a computer algorithm to give the best count. The fluorescence-based platelet count uses a patented fluorescent dye containing oxazine, which stains mitochondria and rough surface endoplasmic reticulum and is platelet-specific when used with a dedicated fluorescence channel, thus minimizing interference with RBC fragments, small RBCs, and cytoplasmic fragments of WBCs [25].

•Immunologic methods – Immunologic method of platelet counting using flow cytometry is the proposed International Reference Method for enumeration of platelets [26].

Some of the Abbott instruments use an immunologic platelet counting method in which the sample is treated with a fluorescent monoclonal antibody against CD61, and platelets are identified by cluster analysis on a two-dimensional plot of fluorescence versus high forward angle light scatter. This may be especially useful for very low platelet counts or when flagged interferences with the platelet count are noted. In one study, this method was found to be the most accurate of all analyzers [27]. In another study, the optical and immunologic methods were in good agreement for platelet counts in the range of 25,000 to 547,000/microL; for platelet counts <25,000/microL, the optical method tended to overestimate the platelet count (figure 2) [28].

Mean platelet volume (platelet size) — The mean platelet volume (MPV) is the average platelet size in femtoliters (fL). It is determined as of the volume of circulating platelets in fL, similar to the MCV for RBCs. The MPV is inversely proportional to the platelet count since the body regulates the total platelet mass rather than the number or size of the platelets. (See "Megakaryocyte biology and platelet production", section on 'Control of platelet mass'.)

●MPV determination and reporting – There is no international standard for the MPV; it cannot be calibrated. The MPV will vary with the method performed and the instrument used.

The MPV value can be measured directly using optical technology or determined from the geometric mean of the transformed lognormal platelet volume data in impedance technology systems. The result may differ depending on which method was used. As an example of the instrument-to-instrument variation, a study examining MPV ranges in adults with normal platelet counts reported that impedance methods had normal values ranging from 6.0 to 13.2 fL, whereas optical methods had normal values ranging from 5.6 to 12.1 fL [29].

Many laboratories do not report the MPV as a routine part of the CBC because of technical problems associated with the accurate measurement of MPV (eg, swelling of platelets after prolonged exposure to EDTA, lack of an international standard, reference range of MPV varies with the platelet count). Likewise, many clinicians do not use the MPV to evaluate platelet abnormalities since there are more specific tests for platelet disorders. (See "Diagnostic approach to thrombocytopenia in adults" and "Inherited platelet function disorders (IPFDs)".)

In hospital laboratories where the patient's sample is drawn, transported to the laboratory, and run on the instrument within two hours, the MPV is likely to be more accurate.

●Evaluation of abnormal MPV – The MPV is affected by many things including prolonged storage in the collection tube containing EDTA, which raises the MPV.

Evaluation of the MPV should be done in context of the platelet count (algorithm 1).

•Healthy individual, normal CBC – In an otherwise healthy patient without a history of bleeding and with an unremarkable CBC, the MPV is unlikely to be useful in diagnosing an underlying disorder or in predicting an increased bleeding risk.

•Thrombocytopenia – In the setting of unexplained thrombocytopenia, an abnormal MPV may suggest a possible diagnosis but cannot be used to confirm or exclude any of these possibilities. The evaluation of thrombocytopenia is discussed separately. (See "Approach to the child with unexplained thrombocytopenia" and "Diagnostic approach to thrombocytopenia in adults".)

The following associations may be seen:

-High MPV – Indicates active bone marrow production of platelets (as in immune thrombocytopenia [ITP]). A high MPV is also seen in some inherited platelet function disorders (Bernard-Soulier syndrome, Gray platelet syndrome, MYH9-related disease, Paris-Trousseau syndrome) and in some myelodysplastic syndromes (table 5). (See "Inherited platelet function disorders (IPFDs)", section on 'Clinical spectrum (thrombocytopenia, platelet size, syndromic features)'.)

A sequential increase in MPV over time can indicate megakaryocytic regeneration, such as recovery from a hypoplastic or aplastic state.

-Low MPV – Indicates bone marrow suppression, as in aplastic anemia [30]. A low MPV may also be seen with Wiskott-Aldrich syndrome. (See "Wiskott-Aldrich syndrome", section on 'Thrombocytopenia and platelet abnormalities'.)

Examination of the peripheral blood smear is a logical next step when one of these conditions is suspected. (See "Evaluation of the peripheral blood smear".)

•Thrombocytosis – (See "Approach to the patient with thrombocytosis".)

●Inverse relationship between platelet count and MPV – Under normal circumstances, there is an inverse relationship between platelet size and number, as the total platelet mass rather than the platelet count is regulated by thrombopoietin. Thus, the MPV is likely to be higher in destructive thrombocytopenias when megakaryocytes are being stimulated, and the MPV is likely to be lower in states of bone marrow hypoplasia or aplasia. (See "Megakaryocyte biology and platelet production", section on 'Control of platelet mass'.)

An exception to this relationship occurs with splenic sequestration, in which a low MPV is seen because the spleen sequesters large platelets. Similarly, a higher MPV is seen in hyposplenic states since there is no spleen to sequester the larger platelets.

Reticulated platelets and immature platelet fraction (IPF) — Reticulated platelets are the youngest circulating platelets, analogous to the relationship between reticulocytes and mature RBCs. Like reticulocytes, reticulated platelets have an increased RNA content. Measurement of the RNA content of platelets with thiazole orange dye can be useful in the diagnostic classification of thrombocytopenia and in monitoring recovering thrombopoiesis (figure 3) [31-33].

Another method to quantify reticulated platelets, the immature platelet fraction (IPF), has been developed on some Sysmex instruments using a proprietary fluorescent dye containing oxazine and polymethine. This method has demonstrated increased IPF in patients with thrombocytopenia due to increased peripheral platelet destruction [25]. Comparison of the IPF with reticulated platelets as measured by flow cytometry has shown good correlation [34].

IPF may be increased in various conditions with increased platelet production, but lack of standardization has hindered clinical use [35-41].

WBC PARAMETERS — White blood cell (WBC) parameters include the absolute WBC count and the WBC differential.

Absolute WBC count — This is the number of WBCs per microL of blood (number of WBCs x 10⁹/L). Neutrophils make up the majority of WBCs, and abnormal WBC counts are often (but not always) due to abnormal neutrophil counts.

●Low WBC count – A decreased WBC count (leukopenia) may be seen in a number of conditions including infections, bone marrow disorders, splenomegaly, and autoimmune conditions. (See "Overview of neutropenia in children and adolescents" and "Approach to the adult with unexplained neutropenia" and "Approach to the adult with lymphocytosis or lymphocytopenia", section on 'Lymphocytopenia'.)

●High WBC count – An elevated WBC (leukocytosis) may be seen in non-neoplastic conditions (especially infections) and in certain cancers. If the WBC is elevated, enumeration of the WBC differential and review of the peripheral blood smear are used along with clinical evaluation to determine the cause. (See "Approach to the patient with neutrophilia" and "Approach to the adult with lymphocytosis or lymphocytopenia", section on 'Causes of lymphocytosis'.)

Conditions that can interfere with an accurate WBC count are summarized in the table (table 3); these include sample clotting, platelet clumping or giant platelets that can lead to spuriously low platelet counts, high WBC counts, and high percent lymphocytes. Most instruments flag significant particle interference in this region.

When the WBC count is abnormal, a differential should be obtained and the peripheral blood smear examined to enumerate which type of WBC is decreased or increased. (See 'WBC differential' below and "Evaluation of the peripheral blood smear", section on 'White blood cells' and "Evaluation of the peripheral blood smear", section on 'Worrisome findings'.)

WBC differential — Automated differentiation of WBC subsets in liquid suspension began in the 1980s. Flow-through automated differentials were first performed using standard electrical resistance or impedance technology. A weak lysing reagent is used to lyse the red blood cells (RBCs) and shrink the WBC membranes around the nucleus, resulting in three peaks of leukocyte subsets, resulting in a three-part differential that is used in many of the smaller modern hematology instruments:

●Lymphocytes and basophils – The smallest size group (35 to 90 fL).

●Granulocytes – The largest size group (>160 fL). Includes segmented and band neutrophils and eosinophils. Can be used to give an estimate of the absolute neutrophil count (ANC); however, the ANC may not be completely accurate, particularly if significant numbers of bands, immature granulocytes, or eosinophils were present.

●Other mononuclear cells – Smaller intermediate size peak between 90 and 160 fL. Includes monocytes, immature granulocytes, and a portion of the eosinophils.

Abnormal cells can obscure the valleys between these three peaks, resulting in error "flags" and inaccurate counts. Most instruments also flag the automated WBC differentials when significant interferences are identified. Increased numbers of bands and eosinophils and more extreme abnormalities (blasts, reactive lymphocytes, nucleated RBCs) all give flagged samples, resulting in a slide review rate >30 percent for a hospital inpatient population. Thus, the three-part differential functioned best in the situation of screening a largely healthy outpatient population.

Five-part differentials were first introduced in the early 1990s [3]. Different instrument manufacturers all report at a minimum the basic five leukocyte subsets (neutrophils, eosinophils, basophils, lymphocytes, and monocytes) and have flags to detect the presence of abnormal cells (blasts, reactive lymphocytes, nucleated RBCs, lymphoma cells, mononuclear granulocytes).

Many of the newer analyzers report a seven-part differential, including the quantification of immature granulocytes and nucleated RBCs.

Most instruments are >90 percent sensitive in detecting large numbers of abnormal cells (when the population is >5 percent), but they can lack sensitivity to low frequency abnormalities (<5 percent). Sensitivity and specificity are also poor for detecting increases in band neutrophils.

Modern instruments use a combination of technologies for the WBC differential, including:

●Direct current (DC)

●Radiofrequency (RF) conductivity

●Laser light scattering

●Peroxidase staining

●Propidium iodide fluorescence (for nucleated RBCs and non-viable cells)

●Cell-specific lysing reagents

●Polymethine RNA/DNA histone dye

●Digital imaging

Both DC and RF methods use impedance technology, in which electrical current generated across an aperture opening is reduced as cells pass through the aperture, producing a voltage change (see 'Electrical impedance (Coulter aperture)' above). When DC current is applied, pulse height is related to overall cell volume. In contrast, when high RF current is applied across the counting aperture, the pulse height or voltage change is proportional to nuclear size and density.

Flags for abnormal WBCs — Instruments performing automated WBC differentials have a variety of "suspect" flags used to indicate abnormal cell populations and possible inaccuracies in the automated differential.

Flagged samples should be reviewed, generally by examining a peripheral blood film. Typically, this occurs within the clinical laboratory by medical technologists, and a further subset of these samples is sent for pathologist review based on individual laboratory criteria. (See "Evaluation of the peripheral blood smear".)

In general, these "suspect" flags can be divided into the following types:

●Interfering particles – This includes cells or cell fragments at the lower WBC counting threshold or the lowest forward/side light scatter region for lymphocytes. Typical resulting flags include: nRBC (nucleated red blood cell), CLUMP (platelet), GIANT (platelet).

●Large mononuclear cells – These include cells at the monocyte/neutrophil interface or with high values for high angle (90 degree) light scattering. Typical instrument flag: BLAST.

●Large lymphoid cells – These include large cells in the lymphoid region or at the interface between lymphoid and monocyte regions. Typical instrument flags include: ATYPICAL LYMPH, BLAST.

●Neutrophil left shift – A flag can be generated if there is a shifted position in the neutrophil cluster, with a large amount of forward or side light scatter. Typical instrument flags include: IMMATURE NEUTROPHILS, BANDS.

Investigational WBC parameters — The combination of laser light scatter, fluorescence, DNA and RNA binding dyes, and monoclonal antibodies has expanded the scope of the automated differential to allow enumeration of abnormal cell types, such as blasts, immature granulocytes, and reactive lymphocytes. Hematology analyzers also provide additional data in the form of cell population data (CPD) for an individual type of leukocyte [42].

These applications are mostly investigational. Any concern about abnormal or immature WBCs should be addressed by review of the blood smear by an experienced pathologist or other expert.

●Lymphocytes – Some Abbott instruments use fluorescein isothiocyanate (FITC)- and R-phycoerythrin (PE)-conjugated monoclonal antibodies to enumerate CD3, CD4, and CD8 T cell counts [43,44]. CPD on lymphocytes has been used for discriminating viral infections from lymphoid malignancies [45].

●Hematopoietic progenitor cells – The Sysmex XE and XN analyzers use a polymethine dye that binds RNA, DNA, and histones and light scatter to enumerate hematopoietic progenitor cells, which some authors have suggested be used as a surrogate for CD34 stem cell quantification for peripheral blood stem cell harvesting [46,47].

●Neutrophils – CPD on neutrophils has been used to screen for bacterial infection and sepsis [48-50]. Some analyzers also enumerate immature granulocytes (IGs), which is being studied as a predictor of sepsis, although this parameter has low sensitivity [51]. Hematology analyzers are also being used to study neutrophil patterns that correlate with myocardial infarction and acute coronary syndromes [52,53].

SUMMARY

●Methods – During the first half of the 20th century, analysis of the complete blood count (CBC) advanced from manual cell counting on a Wright-Giemsa stained blood film to the use of electrical impedance (in which a cell passing through an aperture distorts the electrical field) to light scattering. Other advances have included automated sampling and bar code reading for sample identification, built-in mixing, sample monitor for clots and sample size, cell size gating, methods to measure or calculate red blood cell (RBC) indices, and methods to distinguish among different types of white blood cells (WBCs). (See 'Development of cell counting methods' above.)

●Modern instruments – Modern instruments are able to run multiple parameters; the table summarizes parameters for specific instruments (table 2). The RBC count, RBC indices, WBC count, and platelet count are routinely performed on all samples. Other tests such as the reticulocyte count and WBC differential are performed when requested, saving reagent costs. Ideally, if a reticulocyte count is requested, it should be done at the time the CBC is ordered, but it can be added on within 48 hours. (See 'General features of modern instruments' above.)

●Sample handling – A properly obtained sample analyzed within 72 hours of collection gives the most accurate results. Proper collection of blood (and avoidance of hemolysis from small needles) is important. Sources of error are summarized in the table (table 3) and described above. (See 'Sample collection and processing' above.)

●RBCs – An elevated RBC count indicates polycythemia (reactive or neoplastic) or thalassemia. A decreased RBC typically indicates anemia. Increased hemoglobin or hematocrit (HCT) may indicate polycythemia or dehydration. Decreased RBC count, hemoglobin, or HCT indicates anemia. The HCT can be measured or calculated. The mean corpuscular volume (MCV) indicates RBC size, and the red cell distribution width (RDW) the degree of size variation; these may help in suggesting the cause of anemia. The amount of hemoglobin in RBCs is reflected in the mean corpuscular hemoglobin (MCH) and mean corpuscular hemoglobin concentration (MCHC). The table summarizes the calculations (table 1). Causes of abnormal findings are discussed above. (See 'RBC parameters' above.)

●Reticulocytes – Reticulocytes are identified by the absence of a nucleus and the presence of measurable amounts of RNA, typically detected with a fluorescent nucleic acid-binding dye. Reticulocyte parameters are under investigation for evaluating iron deficiency. (See 'Reticulocytes' above and "Causes and diagnosis of iron deficiency and iron deficiency anemia in adults", section on 'Iron studies (list of available tests)'.)

●Platelets – Causes of thrombocytopenia and thrombocytosis are discussed separately. (See "Approach to the child with unexplained thrombocytopenia" and "Diagnostic approach to thrombocytopenia in adults" and "Approach to the patient with thrombocytosis".)

The mean platelet volume (MPV) is the average platelet size in femtoliters (fL), similar to the MCV for RBCs. MPV should be evaluated in the context of the platelet count and generally does not require further evaluation if the platelet count is normal (algorithm 1). In individuals with thrombocytopenia, an abnormal MPV may indicate a specific underlying condition. Often, more specific tests and review by a hematologist or other expert is indicated. (See 'Platelet parameters' above.)

●WBCs – Decreased WBC count typically reflects decreased neutrophils and may be seen in infections, bone marrow disorders, splenomegaly, and autoimmune conditions. Increased WBC count (leukocytosis) may be seen in non-neoplastic conditions (especially infections) and in certain cancers. If the WBC is abnormal, enumeration of the WBC differential and review of the peripheral blood smear are used along with clinical evaluation to determine the cause. (See 'WBC parameters' above.)

ACKNOWLEDGMENT — UpToDate gratefully acknowledges Stanley L Schrier, MD (deceased), who contributed as Section Editor on earlier versions of this topic review and was a founding Editor-in-Chief for UpToDate in Hematology.