Haemoglobinometry

The haemoglobin concentration (Hb) of a solution may be estimated by measurement of its colour, by its power of combining with oxygen or carbon monoxide or by its iron content. The methods to be described are all colour or light-intensity matching techniques, which also measure, to a varying extent, any methaemoglobin (Hi) or sulphaemoglobin (SHb) that may be present. The oxygen-combining capacity of blood is 1.34 ml O₂ per g haemoglobin. Ideally, for assessing clinical anaemia, a functional estimation of Hb should be carried out by measurement of oxygen capacity, but this is hardly practical in the routine haematology laboratory. It gives results that are at least 2% lower than those given by the other methods, probably because a small proportion of inert pigment is always present. The iron content of haemoglobin can be estimated accurately,¹ but again the method is impractical for routine purposes. Estimations based on iron content are generally taken as authentic, but iron bound to inactive pigment is included. Iron content is converted into haemoglobin by assuming the following relationship: 0.347 g iron = 100 g haemoglobin.²

Measurement of haemoglobin concentration using a spectrometer (spectrophotometer) or photoelectric colorimeter

Two methods are in common use: (1) haemiglobincyanide (HiCN; cyanmethaemoglobin) method and (2) oxyhaemoglobin (HbO₂) method. There is little to choose in accuracy between these methods, although a major advantage of the HiCN method is the availability of a stable and reliable reference preparation.

Although the HiCN reagent contains cyanide, there is only 50 mg of potassium cyanide per litre and 600–1000 ml would have to be swallowed to produce serious effects. However, the use of potassium cyanide has been viewed as a potential hazard; alternative nonhazardous reagents that have been proposed are sodium azide³ and sodium lauryl sulphate,^4,5 which convert haemoglobin to haemiglobinazide and haemiglobinsulphate, respectively. They are used in some automated systems, but no stable standards are available and they, too, are toxic substances that must be handled with care.

Other methods that have been used include Sahli’s acid-haematin method, which is less accurate because the colour develops slowly, is unstable and begins to fade almost immediately after it reaches its peak. The alkaline-haematin method gives a true estimate of total Hb even if carboxyhaemoglobin (HbCO), Hi or SHb is present; plasma proteins and lipids have little effect on the development of colour, although they cause turbidity. The original method was more cumbersome and less accurate than the HiCN or HbO₂ methods, but a modified method has been developed in which blood is diluted in an alkaline solution with non-ionic detergent and read in a spectrometer at an absorbance of 575 nm against a standard solution of chlorohaemin.^6,7 One evaluation has given encouraging results,⁸ although another study has shown a bias of 2.6% when compared with the reference method, with non-linearity in the relationship between haemoglobin concentration and absorbance at high and low haemoglobins.⁹

Haemiglobincyanide (cyanmethaemoglobin) method

The haemiglobincyanide (cyanmethaemoglobin) method is the internationally recommended method² for determining the haemoglobin concentration of blood. In some countries cyanide reagents are no longer available. The basis of the method is dilution of blood in a solution containing potassium cyanide and potassium ferricyanide. Haemoglobin, Hi and HbCO, but not SHb, are converted to HiCN. The absorbance of the solution is then measured in a spectrometer at a wavelength of 540 nm or a photoelectric colorimeter with a yellow-green filter (e.g. Ilford 625, Wratten 74, Chance 0 Gr1).

Oxyhaemoglobin method

The HbO₂ method is the simplest and quickest method for general use with a photometer. Its disadvantage is that it is not possible to prepare a stable HbO₂ standard, so the calibration of these instruments should be checked regularly using HiCN reference solutions or a secondary standard of preserved blood or lysate (see p. 25). The reliability of the method is not affected by a moderate increase in plasma bilirubin, but it is not satisfactory in the presence of HbCO, Hi or SHb.

Direct spectrometry

The haemoglobin concentration of a diluted blood sample can be determined by spectrometry without the need for a standard, provided that the spectrometer has been correctly calibrated. The blood is diluted 1:201 (or 1:251) with cyanide-ferricyanide reagent (see p. 25) and the absorbance is measured at 540 nm. Haemoglobin concentration is calculated as follows:

where A⁵⁴⁰ = absorbance of solution at 540 nm; 16 114 = monomeric molecular weight of haemoglobin; dilution = 201 when 20 ml of blood are diluted in 4 ml of reagent; 11.0 = millimolar coefficient extinction; d = layer thickness in cm; and 1000 = conversion of mg to g.

When assigning a value to a haemoglobin solution that may be used as a reference preparation, it is necessary first to calibrate the spectrometer. This requires checking wavelength with a holmium oxide filter, absorbance with a set of calibrated neutral density filters and stray light with a neutral density filter at 220 nm (National Physical Laboratory, Teddington, UK). Matched optical or quartz glass cuvettes with a transmission difference of <1% at 200 nm should be used. Subsequently, the calibration of the spectrophotometer can be checked by verifying that it gives an accurate reading of the HiCN standard. Slight deviations from the expected A⁵⁴⁰ HiCN value for the standard may be used to correct the results of test samples for a bias in measurement.²

Direct reading portable haemoglobinometers

Range of Haemoglobin Concentration in Health

See Chapter 2, Tables 2.1, 2.2 and 2.3. It should be noted that there are sex differences, diurnal variations and environmental and physiological factors that must also be taken into account.

Packed cell volume or haematocrit

The packed cell volume (PCV) can be used as a simple screening test for anaemia, as a reference method for calibrating automated blood count systems and as a rough guide to the accuracy of haemoglobin measurements. The PCV × 1000 is about three times the Hb expressed in g/l. In conjunction with estimations of Hb and RBC, it can be used in the calculation of red cell indices. However, its use in under-resourced laboratories may be limited by the need for a specialized centrifuge and a reliable supply of capillary tubes.

Manual cell counts and red cell indices

The principles of manual cell counts, the use of the haemocytometer counting chamber for manually counting white cells and platelets and the limitations of these measurements are described in Chapter 26.

An accurate RBC enables the MCV and MCH to be calculated. In well-equipped laboratories, where these indices are provided by an automated system (see p. 41), they are of considerable clinical importance and are widely used in the classification of anaemia. Where automated analysers are not used, manual RBCs (and consequently, calculations of these red cell indices) are so inaccurate and time-consuming that they have become obsolete.

The only measurement that can be obtained with reasonable accuracy by manual methods is MCHC because this is derived from Hb and PCV from the following formula:

Manual differential leucocyte count

Differential leucocyte counts are usually performed by visual examination of blood films that are prepared on slides by the spread or ‘wedge’ technique. Unfortunately, even in well-spread films, the distribution of the various cell types is not totally random (see below).

For a reliable differential count on films spread on slides, the film must not be too thin and the tail of the film should be smooth. To achieve this, the film should be made with a rapid movement using a smooth glass spreader. This should result in a film in which there is some overlap of the red cells, diminishing to separation near the tail, and in which the white cells in the body of the film are not too badly shrunken. If the film is too thin or if a rough-edged spreader is used, many of the white cells, perhaps even 50% of them, accumulate at the edges and in the tail (Fig. 3.1). Moreover, a gross qualitative irregularity in distribution is the rule: polymorphonuclear neutrophils and monocytes predominate at the margins and the tail; lymphocytes predominate in the middle of the film (Fig. 3.2). This separation probably depends on differences in stickiness, size and specific gravity of the different types of cells.

Figure 3.1 Badly spread film. Two areas of a badly spread film from a patient with a white blood cell count of 20 × 10⁹/l showing (A) many leucocytes in the tail and (B) very few leucocytes in body of film.

Figure 3.2 Schematic drawing of a blood film made on a slide. The film has been spread from left to right. An indication is given of the way the white blood cells are distributed.

Differences in distribution of the various types of cells are probably always present to a small extent even in well-made films. Various systems for performing the differential count have been advocated, but none can compensate for the gross irregularities in distribution in a badly made film. On well-made films, the following technique of counting is recommended.

Method

Count the cells using a ×40 objective in a strip running the whole length of the film. Avoid the lateral edges of the film. Inspect the film from the head to the tail and if fewer than 100 cells are encountered in a single narrow strip, examine one or more additional strips until at least 100 cells have been counted. Each longitudinal strip represents the blood drawn out from a small part of the original drop of blood when it has spread out between the slide and spreader (Fig. 3.3). If all the cells are counted in such a strip, the differential totals will closely approximate the true differential count. This technique is liable to error if cells in the thick part of the film cannot be identified; also, it does not allow for any excess of neutrophils and monocytes at the edges of the film, but this preponderance is slight in a well-made film and in practice makes little difference to the result.

Figure 3.3 Schematic drawing illustrating the longitudinal method of performing differential leucocyte counts. The original drop of blood spreads out between spreader and slide (D–D₁). The film is made in such a way that representative strips of films, such as A–A₁ and B–B₁, are formed from blood originally at A and B, respectively. To perform an accurate differential count, all the leucocytes in one or more strips, such as A–A₁ and B–B₁, should be inspected and classified.

This technique is easy to carry out; with high counts (10–30 × 10⁹/l) a short, 2–3 cm, film is desirable. In patients with very high counts (as in leukaemia), the method has to be abandoned and the cells should be counted in any well-spread area where the cell types are easy to identify. Other systems of counting, such as the ‘battlement’ count, are more elaborate but may minimize error owing to variation of distribution of cells between the centre and the edge of the film. The results of the differential count can be recorded using a multiple manual register or they can be directly entered onto a computer.

The variance of the differential count depends not only on artefactual differences in distribution owing to the process of spreading but also on ‘random’ distribution; together they are by far the most important cause of unreliable differential counts. The random distribution means that, if a total of 100 cells are counted, with a true neutrophil proportion of 50%, the range (± 2SD) within which 95% of the counts will fall is of the order of ± 14% (i.e. 36–64%) neutrophils. A 200-cell count can provide a more accurate estimate; in the previous example, the ± 2SD range will be about 40–60%. In a 500-cell count, the range would be reduced to 44–56% neutrophils. In practice, a 100- or 200-cell count is recommended as a routine procedure. However, if abnormal cells are present in small numbers, they are more likely to be detected when 200–500-cell counts are performed than with a 100-cell count.

Basophil and eosinophil counts

A manual basophil or eosinophil count may be necessary to validate an automated count or when abnormal characteristics of the cells render an automated count unreliable, e.g. with degranulated eosinophils. Count the percentage of eosinophils or basophils in a differential count of all the leucocytes on a stained blood film. If the cells of interest are infrequent, a 500-cell differential count should be performed. If fewer than 500 cells are seen in the film, continue the count on a second film. However, if the eosinophil count is markedly elevated a conventional 100-cell count will suffice for most purposes. Calculate the eosinophil or basophil count per litre from the total leucocyte count. It is essential to have thin, preferably short, films with the leucocytes evenly distributed throughout the film and readily identified (see p. 31).

Reporting the Differential Leucocyte Count

The differential count, expressed as the percentage of each type of cell, should be related to the total leucocyte count and the results should be reported in absolute numbers (× 10⁹/l). Myelocytes and metamyelocytes, if present, are recorded separately from neutrophils. Band (stab) cells are generally counted as neutrophils, but it may be useful to record them separately. They normally constitute <6% of the neutrophils; an increase may point to an inflammatory process even in the absence of an absolute leucocytosis.³² However, the band cell count is imprecise and, although it is sometimes recommended in infants, it has been found to be unhelpful in predicting occult bacteraemia in this group.³³

Platelet count

The method for manual counting of platelets using a counting chamber is described on p. 610. If an RBC by a semiautomated counter is available, it is possible to obtain an approximation of the platelet count by counting the proportion of platelets to red cells in a thin part of a film made from an EDTA-anticoagulated blood sample, using the ×100 oil-immersion objective and, if possible, eyepieces provided with an adjustable diaphragm, as for a reticulocyte count.

Reticulocyte count

Reticulocytes are juvenile red cells; they contain remnants of the ribosomal ribonucleic acid (RNA) that was present in larger amounts in the cytoplasm of the nucleated precursors from which they were derived. Ribosomes have the property of reacting with certain basic dyes such as azure B, brilliant cresyl blue or New methylene blue (see below) to form a blue or purple precipitate of granules or filaments.

This reaction takes place only in vitally stained unfixed preparations. Stages of maturation can be identified by their morphological features. The most immature reticulocytes are those with the largest amount of precipitable material; in the least immature, only a few dots or short strands are seen. Reticulocytes can be classified into four groups, ranging from the most immature reticulocytes, with a large clump of reticulin (group I), to the most mature, with a few granules of reticulin (group IV) (Fig. 3.4).

Figure 3.4 Photomicrographs of reticulocytes showing stages of maturation. (A,B) Most immature (group I); (C,D) intermediate (group II); (E,F) later-stage intermediate (group III); (G) most mature (group IV); and (H) haemolytic anaemia, stained supravitally by New methylene blue.

If a blood film is allowed to dry and is afterwards fixed with methanol, reticulocytes appear as polychromatic red cells staining diffusely basophilic if the film is stained with one of the basic dyes.

Complete loss of basophilic material probably occurs in the bloodstream and, particularly, in the spleen after the cells have left the bone marrow.³⁹ This maturation is thought to take 2–3 days, of which about 24 h are spent in the circulation.

The number of reticulocytes in the peripheral blood is a fairly accurate reflection of erythropoietic activity, assuming that the reticulocytes are released normally from the bone marrow and that they remain in circulation for the normal time period. These assumptions are not always valid because an increased erythropoietic stimulus leads to premature release into the circulation. The average maturation time of these so-called ‘stress’ or stimulated reticulocytes may be as long as 3 days. In such cases, a higher than normal proportion of immature reticulocytes will be found in circulation. A more precise assessment of reticulocyte maturation is possible by quantitative flow cytometry of their RNA content. Nevertheless, adequate information is usually obtained from a simple reticulocyte count recorded either as a percentage of the red cells or, preferably, when the RBC is known, as an absolute number per litre. When there is severe anaemia, the reticulocyte count should be corrected for the anaemia and expressed as a reticulocyte index.⁴⁰

Reticulocyte Stains and Count

Better and more reliable results are obtained with New methylene blue than with brilliant cresyl blue. New methylene blue is chemically different from methylene blue, which is a poor reticulocyte stain. New methylene blue stains the reticulofilamentous material in reticulocytes more deeply and more uniformly than does brilliant cresyl blue, which varies from sample to sample in its staining ability. Azure B is a satisfactory substitute for New methylene blue; it has the advantage that the dye does not precipitate and it is available in pure form.⁴¹ It is used in the same concentration and the staining procedure is the same as with New methylene blue.

Staining Solution

Dissolve 1.0 g of brilliant cresyl blue (CI 51010) or 1.0 g of New methylene blue (CI 52030) or azure B (CI 52010) in 100 ml of 3% trisodium citrate-saline solution (30 g sodium citrate in 1 l saline). Filter once the dye has been dissolved.

Method

Deliver 2 or 3 drops of the dye solution into a 75 × 10 mm plastic tube by means of a plastic Pasteur pipette. Add 2–4 volumes of the patient’s EDTA-anticoagulated blood to the dye solution and mix. Keep the mixture at 37°C for 15–20 min. Resuspend the red cells by gentle mixing and make films on glass slides in the usual way. When dry, examine the films without fixing or counterstaining.

The exact volume of blood to be added to the dye solution for optimal staining depends on the RBC. A larger proportion of anaemic blood, and a smaller proportion of polycythaemic blood, should be added than of normal blood. In a successful preparation, the reticulofilamentous material should be stained deep blue and the non-reticulated cells should be stained diffuse shades of pale greenish blue. Films should not be counterstained. The reticulofilamentous material is not better defined after counterstaining and precipitated stain overlying cells may cause confusion. Moreover, Heinz bodies will not be visible in fixed and counterstained preparations. If the stained preparation is examined under phase contrast, both the mature red cells and reticulocytes are well defined. By this technique, late reticulocytes characterized by the presence of remnants of filaments or threads are readily distinguished from cells containing inclusion bodies. Satisfactory counts may be made on blood that has been allowed to stand (unstained) for as long as 24 h, although the count will tend to decrease after 6–8 h unless the blood is kept at 4°C.

Counting Reticulocytes

An area of film should be chosen for the count where the cells are undistorted and where the staining is good. A common fault is to make the film too thin; however, the cells should not overlap. To count the cells, use the ×100 oil-immersion objective and, if possible, eyepieces provided with an adjustable diaphragm. If eyepieces with an adjustable diaphragm are not available, a paper or cardboard diaphragm, in the centre of which has been cut a small square with sides about 4 mm in length, can be inserted into an eyepiece and used as a less convenient substitute.

The counting procedure should be appropriate to the number of reticulocytes present. Very large numbers of cells have to be surveyed if a reasonably precise count is to be obtained when only small numbers of reticulocytes are present. When the count is <10%, a convenient method is to survey successive fields until at least 100 reticulocytes have been counted and to count the total red cells in at least 10 fields to determine the average number of red cells per field.

Calculation

Number of reticulocytes in n fields = x

Average number of red cells per field = y

Total number of red cells in n fields = n × y

Reticulocyte percentage = [x ÷ (n × y)] × 100%

Absolute reticulocyte count = % × RBC.

Thus, when the reticulocyte percentage is 3.3 and the RBC is 5 × 10¹²/l, the absolute reticulocyte count per litre is as follows: [3.3/100] × 5 × 10¹² = 165 × 10⁹

It is essential that the reticulocyte preparation be well spread to ensure an even distribution of cells in successive fields.

When the reticulocyte count exceeds 10%, only a relatively small number of cells will have to be surveyed to obtain a standard error of 10%.

An alternative method is based on the principle of balanced sampling, using a Miller ocular (Graticules Ltd, Morley Road, Tonbridge, UK). This is an eyepiece giving a square field, in the corner of which is a smaller ruled square, one-ninth the area of the total square (Fig. 3.5). Reticulocytes are counted in the large square and the total number of red cells is counted in the small square.

Figure 3.5 Miller ocular.

The number of fields that should be surveyed to obtain a desired degree of precision depends on the proportion of reticulocytes (Table 3.3).

Table 3.3 Accuracy of reticulocyte counts with Miller ocular

It is essential that the reticulocyte preparation be well spread and well stained. Other important factors that affect the accuracy of the count are the visual acuity and patience of the observer and the quality and resolving power of the microscope. The most accurate counts are carried out by a conscientious observer who has no knowledge of the supposed reticulocyte level, thus eliminating the effect of conscious or unconscious bias.

Differentiating between Reticulocytes and Other Red Cell Inclusions

The decision as to what is and what is not a reticulocyte may be difficult because the most mature reticulocytes contain only a few dots or threads of reticulofilamentous material. Fortunately, in well-stained preparations viewed under the light microscope, the Pappenheimer (iron-containing) type of granular material – usually present as a single small dot, less commonly as multiple dots – stains a darker shade of blue than does the reticulofilamentous material of the reticulocyte. As described earlier, phase contrast will help to distinguish them. If there is any doubt, Pappenheimer bodies can be identified by overstaining the film for iron by Perls’ reaction.

Haemoglobin H undergoes denaturation in the presence of brilliant cresyl blue or New methylene blue, resulting in round inclusion bodies that stain greenish-blue (see Chapter 15, Figs 15.6, 15.7). These can be easily differentiated from reticulofilamentous material (Fig. 15.8).

Heinz bodies are also stained by New methylene blue, but they stain a lighter shade of blue than the reticulofilamentous material of reticulocytes and stain well with methyl violet (Figs 15.5, 15.6).

Automated blood count techniques

A variety of automated instruments for performing blood counts are in widespread use. Semiautomated instruments require some steps (e.g. dilution of a blood sample) to be carried out by the operator. Fully automated instruments require only that an appropriate blood sample is presented to the instrument. Semiautomated instruments often measure a small number of components (e.g. WBC and Hb). Fully automated multichannel instruments usually measure from 8 to 20 components for the basic FBC and white blood cell differential, including some variables that have no equivalent in manual techniques. Automated instruments usually have a high level of precision, which, for cell counting and cell-sizing techniques, is greatly superior to that achievable with manual techniques. If instruments are carefully calibrated and their correct operation is ensured by quality control procedures, they produce test results that are generally accurate. When blood has abnormal characteristics, the results for one or more parameters may be aberrant; instruments are designed so that such inconsistent results are ‘flagged’ for subsequent review. The abnormal characteristics that lead to inaccurate counts vary between instruments, so it is important for instrument operators to be familiar with the types of factitious results to which their instruments are prone.

Blood cell counters may have automated procedures for sample recognition (e.g. by bar-coding), for ensuring that adequate sample mixing occurs, for taking up the test sample automatically and for detection of clots or inadequately sized samples. Ideally, blood sampling is carried out by piercing the cap of a closed tube so that samples that carry an infection hazard can be handled with maximum safety.

Laboratories performing large numbers of blood counts each day require fully automated blood counters capable of the rapid production of accurate and precise blood counts, including platelet counts and differential counts, either three-part or five- to seven-part. The sample throughput required varies with the workload and the timing of arrival of blood specimens in the laboratory, but for most large laboratories, a throughput of 100 or more samples per hour is required. Sample size and the availability of a ‘predilute’ mode are particularly relevant if the laboratory receives many paediatric specimens.

Choice of an instrument for an individual laboratory, as well as for point-of-care sites outside the laboratory (see p. 574), should take account of capital expenditure and running costs, including maintenance and reagents; size of instrument; requirements of services such as water, compressed air, drainage and an electricity supply with stable voltage; environmental disturbance by generation of heat, vibration and noise; any influence on performance by the ambient temperature and humidity; storage requirements for the often bulky reagents; ease of operation; and the likely level of support that can be expected from the manufacturer.

A practical guide on the principles of the various systems has been published,⁴⁵ and there are guidelines to help in the choice of an instrument suitable for the needs of an individual laboratory and also to assess its performance, as compared with the claims of the manufacturer, when it has been installed and is being used in routine practice.⁴⁶ Choice of instrument may be aided by reference to published reports of instrument evaluations and related monographs.^45,47–49 Some semiautomated instruments aspirate a sample of accurately determined volume and so can perform absolute cell counts and accurate estimations of Hb. Most automated instruments, however, count for a specified period of time rather than measure an exact volume of blood; they therefore require calibration by means of the direct counts derived from instruments counting cells in a defined volume of diluted blood. For some variables, instruments are calibrated by the manufacturer, but others require calibration in the laboratory. Performance characteristics of an instrument vary over time, so periodic recalibration is needed: both when quality control procedures indicate the necessity and when certain components are replaced.

Haemoglobin concentration

Many automated counters still measure haemoglobin concentration by a modification of the manual HiCN method with cyanide reagent; however manufacturers are changing their methods to allow the use of a non-hazardous chemical, such as sodium lauryl sulphate, imidazole, sodium dodecyl sulphate or dimethyl laurylamine oxide, which avoids possible environmental hazards from disposal of large volumes of cyanide-containing waste. Modifications include alterations in the concentration of reagents and in the temperature and pH of the reaction. A non-ionic detergent is included to ensure rapid cell lysis and to reduce turbidity caused by cell membranes and plasma lipids. Measurements of absorbance are made for haemoglobin measurement at various wavelengths depending on the final stable haemochromogen, cyanmethaemoglobin, oxyhaemoglobin, methaemoglobin or monohydroxyferriporphyrin and at a set time interval after mixing of blood and the active reagents but before the reaction is completed.

Red blood cell count

Red cells and other blood cells can be counted in systems based on either aperture impedance or light-scattering technology. Because large numbers of cells can be counted rapidly, there is a high level of precision. Consequently, electronic counts have rendered the RBC and the red cell indices derived from it (the MCV and the MCH) of much greater clinical relevance than was possible when only a slow and imprecise manual RBC was available.

Counting systems

Impedance Counting

Impedance counting, first described by Wallace Coulter in 1956,⁵⁰ depends on the fact that red cells are poor conductors of electricity, whereas certain diluents are good conductors; this difference forms the basis of the counting systems used in Beckman Coulter, Sysmex, Abbott, Horiba Medical and a number of other instruments.

For a cell count, blood is highly diluted in a buffered electrolyte solution. The flow rate of this diluted sample is controlled by a mercury siphon (as in the original Coulter system) or by displacement of a tightly fitting piston. This results in a measured volume of the sample passing through an aperture tube of specific dimensions (e.g. 100 mm in diameter and 70 mm in length). By means of a constant source of electricity, a direct current is maintained between two electrodes, one in the sample beaker or the chamber surrounding the aperture tube and another inside the aperture tube. As a blood cell is carried through the aperture, it displaces some of the conducting fluid and increases the electrical resistance. This produces a corresponding change in potential between the electrodes, which lasts as long as the red cell takes to pass though the aperture; the height of the pulses produced indicates the volume of the cells passing through. The pulses can be displayed on an oscillograph screen. The pulses are led to a threshold circuit provided with an amplitude discriminator for selecting the minimal pulse height, which will be counted (Fig. 3.6). The height of the pulses is used to determine the volume of the red cells.

Figure 3.6 Effect of threshold discrimination (horizontal axis) in separating cell signals from background noise.

Reliability of electronic counters

Electronic counts are precise, but care needs to be taken so that they are also accurate. The recorded count on the same sample may vary from instrument to instrument and even between different models of the same instrument. Inaccuracy may be introduced by coincidence (i.e. by two cells passing through an orifice simultaneously and being counted as one cell or by a pulse being generated during the electronic dead time of the circuit); by recirculation of cells that have already been counted; by red cell agglutination (which causes a clump of cells to be counted as one cell); and by the counting of bubbles, lipid droplets, microorganisms or extraneous particles as cells. Faulty maintenance may lead to variation in the volume aspirated or the flow rate. Single-channel instruments may have their thresholds set incorrectly and multichannel instruments may be incorrectly calibrated.

A statistical correction may be applied for coincidence (coincidence correction); in some instruments, this is done automatically by electronic editing. Errors of coincidence can be detected by carrying out a series of measurements at various dilutions of the same specimen, plotting the data on graph paper and then extrapolating the graph to the baseline for the true value. Alternatively, the need for coincidence correction can be avoided by having the dimensions and flow characteristics of the aperture through which the cells pass such that cells can only pass in single file; this may be achieved by sheathed flow or hydrodynamic focusing in which diluted blood is injected into a sheath of fluid as it flows into the sensing zone. This induces the cells to pass through the centre of the sensing zone in single file and free of distortion. Coincidence can be more effectively reduced with sheathed flow and precisely focused light in an electro-optical detector than in an impedance counter so that less dilution of the blood sample is needed.⁴⁶ Electrical impulses generated by recirculation of cells can be eliminated by electronic editing; alternatively, recirculation of cells in the region of the aperture can be prevented by ‘sweep flow’ in which a directed stream of diluent sweeps cells and debris away from the aperture, thus preventing cells from being recounted and debris from being counted as cells.

Inaccurate counts consequent on red cell agglutination are usually the result of cold agglutinins. They are recognized as erroneous because of an associated marked factitious elevation of the MCV. A correct count can be achieved by prewarming the blood sample and, if necessary, also prewarming the diluent.

A correct RBC and, particularly, a correct measurement of the MCV is dependent on the use of an appropriate diluent. For impedance counters, pH, temperature and rate of ionization have to be standardized and remain constant because changes alter the electrical field and may lead to artefactual alterations in the size, shape and stability of the blood cells in the diluent. Diluents must be free of particles and give a background count of <50 particles in the measured volume. The correct diluent for each individual instrument must be used; other diluents, even those made by the same manufacturer, may not be interchangeable. Any laboratories using diluents other than those recommended by the manufacturer of the instrument must satisfy themselves that no error is being introduced.

For red cell counting in simple single-channel counters a suitable diluent requires a pH of 7.0–7.5 and osmolality of 340 ± 10 mmol. Physiological saline (9 g/l NaCl) or phosphate-buffered saline, which have the advantages of simplicity and ready availability, can be used as a red cell diluent, provided that the counts are performed immediately after dilution to avoid errors owing to sphering. Commercial solutions of saline (for intravenous use) are usually particle-free. Other solutions may require filtration through a 0.22 or 0.45 mm micropore filter to remove dust.

Setting Discrimination Thresholds

An accurate RBC requires that thresholds be set so that all red cells, but a minimum of other cells, are included in the count. Some counters have a lower threshold but no upper threshold so that white cells are included in the ‘RBC’. Because the WBC is usually very low in relation to the RBC, this is not usually of practical importance; however, an appreciable error can be introduced if the WBC is greatly elevated, particularly if the patient is also anaemic. The setting of the lower threshold is of considerable importance because it is necessary to ensure that microcytic red cells are included in the count without also counting large platelets.

Current multichannel instruments, both impedance counters and counters using light-scattering technology, have thresholds that are either precalibrated by the manufacturer or are automatically adjusted, depending on the characteristics of individual blood samples. Single-channel impedance instruments capable of performing a direct RBC require setting of thresholds so as to separate pulses generated by red cells from background noise and from pulses generated by platelets. This is done by adjusting the aperture current and the pulse amplification. A simple method is to dilute a fresh blood sample and carry out successive counts on the suspension, while the lower threshold control is moved incrementally from its maximum to its minimum position. At the maximum position, the count should be zero or close to zero and the counts will increase as the amplitude is reduced. The counts at each setting are plotted on arithmetic graph paper (Fig. 3.7). The correct threshold setting is at the left of the horizontal part of the graph before the line begins to slope. It is important to check that the setting selected is valid for microcytic cells. The threshold can be defined more precisely for an individual sample by means of a pulse height analyser linked to the counting system. The lower threshold is correctly set if beyond this point there are <0.5% of the counts at the peak (mode) of the pulse size distribution curve (Fig. 3.6).

Figure 3.7 Method to establish working conditions of cell counters. The correct setting of the threshold (at arrow) is intended to exclude noise pulses without loss of the signal pulses produced by the blood cells.

Packed cell volume and mean cell volume

Modern automated blood cell counters estimate PCV/haematocrit by technology that has little connection with packing red cells by centrifugation. It is sometimes convenient to use different terms to distinguish the manual and automated tests and for this reason the International Council for Standardization in Haematology has suggested that the term ‘haematocrit’ (Hct) rather than PCV should be used for the automated measurement. However, it should be noted that, in the past, the terms ‘packed cell volume’ and ‘haematocrit’ have been used interchangeably for the manual procedure.

With automated instruments, the derivations of the RBC, PCV and MCV are closely interrelated. The passage of a cell through the aperture of an impedance counter or through the beam of light of a light-scattering instrument leads to the generation of an electrical pulse, the height of which is proportional to cell volume. The number of pulses generated allows the RBC to be determined, as discussed earlier. Pulse height analysis allows either the MCV or the Hct to be determined. If the average pulse height is computed, this is indicative of the MCV and the Hct can be derived by multiplying the estimated MCV by the RBC. Similarly, if the pulse heights are summated, this figure is indicative of the Hct and the MCV can, in turn, be derived by dividing the Hct by the RBC.

Automated instruments require calibration before the Hct or MCV can be determined. Calibration of the Hct can be based on manual Hct determinations. Alternatively, the MCV can be calibrated by means of the pulse heights generated by latex beads, stabilized cells or some other calibrant containing particles of known size; however, unfixed human red cells that are biconcave and flexible will not necessarily show the same characteristics in a cell counter as latex particles or some other artificial calibrant. BCR Certified preparations are available from the Institute for Reference Materials and Measurements (IRMM) (see p. 588). Aperture-impedance systems measure an apparent volume that is greater than the true volume, being influenced by a ‘shape factor’;⁵² this factor is less than 1.1 for young, flexible red cells; between 1.1 and 1.2 for fixed biconcave cells; and about 1.5 for spheres, whether they be fixed cells or latex spheres.^51,52

The MCV, and therefore the Hct, as determined by an automated counter, will vary with certain cell characteristics other than volume. As indicated earlier, such characteristics include shape, which in turn is partly determined by flexibility. With impedance counters, the normal disc-shaped red cell becomes elongated into a cigar shape as it passes through the aperture; this is caused by deformation in response to shear force, which occurs in cells of normal flexibility. Cells with a reduced haemoglobin concentration undergo more elongation than normal cells; this leads to a reduced ‘shape factor’, a reduced pulse height in relation to the true size of the cell and underestimation of the MCV. Conversely, cells with abnormally rigid membranes and cells such as spherocytes with a high haemoglobin concentration will undergo less deformation than normal and the MCV will be overestimated. Earlier light-scattering instruments also underestimated the volume of red cells with a reduced haemoglobin concentration because light scattering was affected by the haemoglobin concentration.⁵³ These artefacts are seen even with normal red cells of varying haemoglobin concentration but are more apparent with red cells from patients with defects in haemoglobin synthesis such as those from patients with iron deficiency. Light-scattering instruments have been developed to avoid artefacts of this type. Cells are isovolumetrically sphered; light-scattering characteristics of sphered red cells are predictable and permit the computation of both individual cell volume and intracellular haemoglobin concentration using a calibrated Mie map that describes the scatter and refraction characteristics of spherical particles in a monochromatic light source.⁵⁴ Light scattering by each individual cell is measured at two angles: low angle at 2–3° and high angle scatter at 5–15°, which permits computation of both cell volume and haemoglobin concentration.⁵³ The measure of cellular haemoglobin is designated as the cellular haemoglobin concentration mean (CHCM) to distinguish it from the traditional MCHC derived from the Hb and the PCV. If all measurements are accurate, the CHCM and the MCHC should give the same results, thus providing an internal quality control mechanism.

The automated MCV and Hct are prone to certain errors that do not occur or are less of a problem with manual methods. These include those resulting from microclots or partial clotting of the specimen, extreme microcytosis and the presence of cryoglobulins or cold agglutinins; the last is a relatively common cause of factitious elevation of the MCV because clumps of cells are sized as if they were single cells. Because the RBC is underestimated, the Hct is less affected, although it is also inaccurate. It is rare for warm agglutinins to cause a similar problem. Sickling may cause a factitious increase in MCV and Hct, whereas alterations in plasma osmolarity occurring, for example, in severe hyperglycaemia, also cause factitious elevation of the MCV and Hct.^49,55,56

Red cell indices

Red cell indices traditionally have been the derived parameters of MCV, MCH and MCHC; more recently, red cell distribution width (RDW) has also been included and, for some instruments, haemoglobin distribution width (HDW). These indices can provide a basis for classifying anaemias and in various combinations they have been used to aid in the distinction between iron deficiency and thalassaemias.^57–59 It is important to note, however, that these formulae may not be consistent between different instruments and their use provides only a guide to the most likely diagnosis. When diagnosis is important, as in preconceptual or antenatal screening for thalassaemia, definitive tests are required, even in patients whose red cell indices are more suggestive of iron deficiency.

Variations in red cell volumes: red cell distribution width

Automated instruments produce volume distribution histograms that allow the presence of more than one population of cells to be identified. Instruments may also assess the percentage of cells falling above and below given MCV thresholds and ‘flag’ the presence of an increased number of microcytes or macrocytes. Such measurements may indicate the presence of a small but significant increase in the percentage of either microcytes or macrocytes before there has been any change in the MCV.

Most instruments also produce a quantitative measurement of the variation in cell volume, an equivalent of the microscopic assessment of the degree of anisocytosis. This parameter has been named the ‘red cell distribution width’. The RDW is derived from pulse height analysis and can be expressed either as the standard deviation (SD) in fl or as the coefficient of variation (CV) (%) of the measurements of the red cell volume. The RDW SD is measured by calculating the width in fl at the 20% height level of the red cell size distribution histogram and the RDW CV is calculated mathematically as the coefficient of variation, i.e. RDW (CV)% = 1SD/MCV × 100%.

Most instruments express the RDW as the SD, but Sysmex instruments and the Beckman Coulter instrument the DxH express it as both SD and CV. The normal reference range is in the order of 12.8 ± 1.2% as CV and 42.5 ± 3.5 fl as SD. However, widely different ranges have been reported; therefore it is important for laboratories to determine their own reference ranges. The RDW expressed as the CV has been found of some value in distinguishing between iron deficiency (RDW usually increased) and thalassaemia trait (RDW usually normal) and between megaloblastic anaemia (RDW often increased) and other causes of macrocytosis (RDW more often normal).

Percentage hypochromic red cells and variation in red cell haemoglobinization: haemoglobin distribution width

Instruments that determine the haemoglobin concentration of individual red cells provide the percentage of hypochromic red cells, with distribution curves of the haemoglobin concentration, and are able to ‘flag’ the presence of increased numbers of hypochromic or hyperchromic cells. The percentage of hypochromic red cells depends on the concentration of haemoglobin in individual cells rather than being a mean, such as MCH or MCHC. It is a more sensitive marker of the availability of iron for erythropoiesis because small changes in the number of red cells with inadequate haemoglobin can be measured before there is any change in the MCHC. Hypochromic red cells are defined as cells with a haemoglobin concentration of less than 28 g/dl (280 g/l).⁶¹ In the healthy population the percentage of hypochromic red cells does not exceed 2.5% and values greater than this are indicative of iron deficient erythropoiesis.⁶² It has been reported to be a useful indicator of functional iron deficiency (where reticuloendothelial iron stores are normal or even high, but the iron is not delivered to erythroblasts and is therefore unavailable for erythropoiesis) in haemodialysis patients. Other manufacturers’ instruments have other parameters reported to be equivalent to percentage hypochromic red cells, such as low haemoglobin density (LHD%) on some Beckman Coulter instruments.⁶³ LHD% is derived from a sigmoid transformation of the MCHC and has been proposed as a parameter to assess the available iron stores for erythropoiesis.

The degree of variation in red cell haemoglobinization is quantified as the haemoglobin distribution width or HDW; this is the CV of the measurements of haemoglobin concentration of individual cells. The normal 95% range is 1.82–2.64. Because the volume of individual red cells is determined, it is possible to distinguish between hypochromic microcytes, which are indicative of a defect in haemoglobin synthesis and hypochromic macrocytes, which often represent reticulocytes.⁶⁴ The identification of an increased percentage of hyperchromic cells may be caused by the presence of spherocytes, irregularly contracted cells or sickled cells.

Total white blood cell count

The WBC is determined in whole blood in which red cells have been lysed. The lytic agent is required to destroy the red cells and reduce the red cell stroma to a residue that causes no detectable response in the counting system without affecting leucocytes in such a manner that the ability of the system to count them is altered. Various manufacturers recommend specific reagents and for multichannel instruments that also perform an automated differential count use of the recommended reagent is essential. For a simple single-channel impedance counter, the following fluid is satisfactory:

Relatively simple instruments are also available that determine the Hb and the WBC by consecutive measurements on a single blood sample. The diluent contains a reagent to lyse the red cells and another to convert haemoglobin to haemiglobincyanide. Hb is measured by a modified HiCN method and white cells are counted by impedance technology. Apart from the reagents specified by the manufacturers, a diluent containing potassium cyanide and potassium ferricyanide together with ethylhexadecyldimethyl-ammonium bromide can be used.^65,66

Fully automated multichannel instruments perform WBCs by impedance or light-scattering technology or both. Residual particles in a diluted blood sample are counted after red cell lysis or, in the case of some light-scattering instruments, after the red cells have been rendered transparent. Thresholds are set to exclude normal platelets from the count, although giant platelets are included. Some or all of any nucleated red cells present are usually included, so that when nucleated red cells are present the count approximates more to the TNCC than to the WBC.

Factitiously low automated WBCs occasionally occur as a consequence of leucocyte agglutination, prolonged sample storage or abnormally fragile cells (e.g. in leukaemia). Factitiously high counts are more common and usually result from failure of lysis of red cells. With certain instruments this may occur with the cells of neonates or be consequent on uraemia or on the presence of an abnormal haemoglobin such as haemoglobin S or haemoglobin C; high counts may also be the result of microclots, platelet clumping or the presence of a cryoglobulin.

Automated differential count

Most automated differential counters that are now available use flow cytometry incorporated into a full blood counter rather than being stand-alone differential counters. Increasingly, automated blood cell counters have a differential counting capacity, providing either a three-part or a five- to seven-part differential count. Counts are performed on diluted whole blood in which red cells are either lysed or are rendered transparent. A three-part differential count assigns cells to categories usually designated: (1) ‘granulocytes’ or ‘large cells’; (2) ‘lymphocytes’ or ‘small cells’; and (3) ‘monocytes’, ‘mononuclear cells’, or ‘middle cells’. In theory, the granulocyte category includes eosinophils and basophils, but in practice it is common for an appreciable proportion of cells of these types to be excluded from the granulocyte category and to be counted instead in the monocyte category.⁶⁷ Some other three-part differentials categorize leucocytes as WBC-small cell ratio (equivalent to lymphocytes), WBC-middle cell ratio (equivalent to monocytes, eosinophils and basophils) and WBC-large cell ratio (equivalent to neutrophils).⁶⁸

Five- to seven-part differential counts classify cells as neutrophils, eosinophils, basophils, lymphocytes and monocytes and in an extended differential count may also include immature granulocytes or large immature cells (composed of blasts and immature granulocytes) and atypical lymphocytes (including small blasts). Automated instruments performing differential counts (that do not enumerate immature granulocytes or nucleated red cells separately) are able to ‘flag’ or reject counts from the majority of samples with nucleated red cells, myelocytes, promyelocytes, blasts or atypical lymphocytes. To a lesser extent, instruments incorporating a three-part differential count, although not capable of enumerating eosinophils or basophils as individual categories of cells, are able to flag a significant proportion of samples that have an increased number of one of these cell types.

Both impedance counters and light-scattering instruments are capable of producing three-part differential counts from a single channel; the categorization is based on the different volume of various types of cell following partial lysis and cytoplasmic shrinkage. Most five- to seven-part differential counts require two or more channels in which cell volume and other characteristics are analysed by various modalities (Table 3.4). Analysis may be dependent only on volume and other physical characteristics of the cell or also on binding of certain dyes to granules or activity of cellular enzymes such as peroxidase. Technologies used to study cell characteristics include light scattering and absorbance and impedance measurements with low- and high-frequency electromagnetic current or radiofrequency current. Cells may have been exposed to lytic agents or a cytochemical reaction may have occurred before cell characteristics are studied. Two-parameter analysis or more complex discriminant functions divide cells into clusters that can be matched with the position of the various white cell clusters in normal blood. Thresholds, some fixed and some variable, divide clusters from one another, permitting cells in each cluster to be counted.

Table 3.4 Automated full blood counters with a five-part or more differential counting capacity^a

a In addition to the blood counters listed here, there are an increasing number of instruments, some designed for point-of-care testing, on the market that are capable of providing full differential or partial differential counts using various technologies.

Automated differential counters using flow cytometry count a far greater number of cells than is possible with a manual differential count. Automated counts are consequently much more precise than manual counts. The accuracy of automated counters is less impressive than their precision. With all types of counters, unusual cell characteristics or ageing of a blood specimen can lead to misclassification of cells. Although the majority of samples containing abnormal cells are ‘flagged’, this is not invariably so; the presence of nucleated red cells, immature granulocytes, atypical lymphocytes and blasts (even occasionally quite large numbers of blasts) may not give rise to a ‘flag’. However, human observers performing a 100-cell manual differential count also miss significant abnormalities. In general, automated counts have compared favourably with routine manual counts, especially if the instruments are assigned only two functions, performing differential counts on normal samples and ‘flagging’ abnormal samples. If morphological abnormalities are flagged, microscopic examination of a stained blood film should always be undertaken.

The automated immature granulocyte count

Most fully automated analysers now report an immature granulocyte count. Promyelocytes, myelocytes and metamyelocytes are all included in the automated immature granulocyte count and are not identified as separate classes of cells. The presence of low numbers of immature granulocytes is more reliably detected on automated haematology analysers than by manual microscopy, due to the higher number of cells counted. Often low numbers of immature granulocytes, particularly in leucopenic samples or when small percentages are present, are missed in a 100-cell differential count or film review. Immature granulocytes may be identified either by a combination of light absorbance (after staining of the cells) and impedance or by flow cytometry to detect side-scattered light and fluorescence of cells stained with a fluorescent dye. Measurement of immature granulocytes may be clinically relevant. The percentage of immature granulocytes as measured by the Sysmex XE-2100 has, for example, been found to be predictive of infection, although it should be noted that it is no more predictive than the absolute neutrophil count.⁶⁹

Some instruments do not quantitate immature granulocytes and still rely on an abnormal white cell flag generated by the analyser to indicate their possible presence in the blood sample.

For instruments that do not report a separate automated count for nucleated red blood cells (NRBC), automated differential counts often include some, but not all, NRBC in the total ‘WBC’; thus, in the presence of a significant number of NRBC, the total count is neither a true ‘WBC’ nor a true ‘TNCC’ and the absolute WBC counts calculated from the total will necessarily be somewhat erroneous. This differs from the situation with earlier instruments that included any NRBC in the ‘WBC’. It may be possible to make some assessment of the proportion of the NRBCs included in the total count by studying the graphic output of the instrument; otherwise, if accurate absolute counts of different leucocyte types are needed, it is necessary to revert to earlier instruments to provide the TNCC and to correct it to a WBC by means of a differential count.

The automated nucleated red blood cell count

The ability of haematology instruments to perform precise and accurate automated NRBC counts over the entire concentration range in peripheral blood offers advantages to the diagnostic laboratory. Enumeration of NRBC is important because their presence can have a direct effect on the accuracy of the WBC on some blood cell counters. The correct WBC was previously only obtained by examination of a peripheral blood film. The NRBC are reported as the number per 100 white blood cells and subtraction of the number of NRBC from the total nucleated count gives the correct WBC. The morphological correction of the WBC can be inaccurate since if the nuclear size of an NRBC falls below the white blood cell threshold of the instrument, these cells are not included in the automated WBC in the first place. Instruments currently in use that automatically count NRBC and correct the WBC for NRBC interference include the Abbott Sapphire, the Sysmex XE-2100 and XE-5000, the Beckman Coulter LH750 and DxH, the Horiba Medical Pentra DX120 and the Siemens Advia 2120.

Instruments determine NRBC by staining them with a nuclear dye and using either fluorescence laser light scatter or flow cytometry to separate them from WBC or a combination of impedance and cell volume. Beckman Coulter instruments use cell volume conductivity and scatter measurements. The Siemens Advia 2120 utilizes nuclear density and degree of peroxidase staining to identify NRBC. The white cell count and differential are corrected for the presence of nucleated red blood cells where necessary.

The NRBC counting method on some instruments is not a direct measure of the cells and there is the possibility of other interfering substances in blood occupying the NRBC signature position and producing false-positive results.

Automated digital imaging analysis of blood cells

Over the last 20 years automated imaging processes have started to be introduced where stained blood films are scanned by a computer-driven microscope and leucocytes classified; early methods were slow and had difficulty in classifying abnormal cells and, as only a small number of cells were counted in a reasonable time, the precision of the automated count was no better than that of a manual count.⁷⁰ However, with improved computing technology and with the use of artificial neural networks, such instruments, e.g. CellaVision DM96 AB, Lund, Sweden and HemoFAXS, Tissuegnostics GmbH, Tabarstasse 10/12/8, A-1020, Wein, Austria, are now capable of providing a useful differential count on blood samples, even those containing abnormal cells.⁷¹ The DM96 is composed of a slide scanning unit and a computer. The scanning unit consists of a motorized microscope and a digital charge-coupled device (CCD) camera. The instrument scans the stained blood film, identifies potential white cells, takes digital images of them and uses artificial neural network-based software to analyse the cells. Up to 30 films an hour can be processed. Digital images of pre-classified cells are presented to the operator on a computer screen for conformation or re-classification. The operator of the instrument needs to be skilled in blood cell morphology in order to accept or re-classify accurately the cells presented.⁷²

New White Cell Parameters

Many instruments are able to ‘flag’ the presence of abnormal leucocytes by features such as an alteration in cell size, nuclear size or cell granularity which causes changes in impedance or light-scattering characteristics. Automated white cell counters can also analyse cell characteristics by novel technologies and identify cell types by features that differ greatly from those used when a blood film is examined visually. It is possible, for example, to identify eosinophils by the ability of their granules to polarize light⁷³ or to detect a left shift or the presence of blasts by the reduced light scattering of the nuclei of more immature granulocytes. There is also the potential to produce information that is not directly analogous with that available from a manual differential count. Recently white blood cell differential parameters have been reported to demonstrate clinical utility in the diagnosis of some diseases. Abnormal cell populations that have previously only triggered an abnormal flag, on some instruments, can now be quantitated.

Numerical data (coordinates) generated by analysers that use volume, conductivity and scatter (VCS) are now available. These coordinates indicating the position and size of the cell cluster in the VCS differential plot are available for neutrophils, lymphocytes, monocytes and eosinophils and cells of each cell type have normal values. Any deviations away from normal are thought to reflect differences in cell size and complexity and may be indicative of a potential disease process or specific to a morphological characteristic. These parameters are still for laboratory use only but it may be possible to use them as advanced flags for specific diseases or conditions in certain clinical circumstances. Lymphocyte positional parameters can be used to distinguish between different lymphoproliferative disorders and viral infections.⁷⁴ Neutrophil conductivity and neutrophil scatter can be used in the detection of dysplastic neutrophils,⁷⁵ with low values for these parameters correlating with neutrophil hypogranulation.

Sepsis-associated neutrophil left shift can be identified by a change from normal values of neutrophil volume and may be used as an indicator of acute bacterial infections.⁷⁶

It has been suggested that VCS and other instrument parameters used for defining leucocyte types might also allow detection of the presence of malaria pigment in white blood cells or larger than normal monocytes and may be used as a screening test for malaria.^77,78

In a similar way, using forward scattered light and fluorescence, new parameters have been developed on the basis of abnormal leucocyte positions in the differential scattergram. Plasma cells appear in a high fluorescence area of the lymphocyte cluster in the differential channel of the XE-2100 and are flagged as atypical lymphocytes; they can now be quantitated.⁷⁹ On the same instrument, NEUT-X is the mean value for side scatter diffraction of the neutrophil population; it represents the internal structure of the neutrophils. It correlates with hypogranularity of neutrophils and when taken into consideration with anaemia is suggestive of a myelodysplastic syndrome.⁸⁰ Instruments such as Siemens Advia analysers that incorporate a cytochemical reaction give information on enzyme activity expressed as the mean peroxidase activity index (MPXI). An increased MPXI has been observed in infections, in some myelodysplastic syndromes and leukaemias, in the acquired immune deficiency syndrome (AIDS) and in megaloblastic anaemia, whereas a reduced MPXI occurs in inherited and acquired neutrophil peroxidase deficiency.^60,81,82 These new parameters provide numerical values for the changes that can be seen in the instruments’ scatter plots by an experienced operator. Such measurements have the potential for clinical usefulness and may allow the development of specific disease flags and new indicators of abnormality.

Automated instrument graphics

Fully automated instruments produce a graphic display of much of the data produced. This is displayed on a colour monitor and can be printed, either in black and white or in colour. Inspection of the graphic display can give further information beyond that which is available from assessment of the numeric data. Displays usually include histograms of red cell, white cell and platelet size and sometimes histograms of red cell haemoglobin concentration and scatter plots of size versus haemoglobin concentration. Differential counts are graphically represented as scatter plots of two variables or scatter plots of discriminant functions derived from more than two variables.

Typical printouts of histograms or scatter plots of current automated instruments are shown in Figure 3.8.

Figure 3.8 Patterns of blood count printout of some automated systems. (A) Beckman Coulter DxH; (B) Sysmex XE-2100; (C) Siemens Advia; (D) Abbot Cell-Dyn 4000.

Platelet count

Platelets can be counted in whole blood using the same techniques of electrical or electro-optical detection as are used for counting red cells. An upper threshold is needed to separate platelets from red cells and a lower threshold is needed to separate platelets from debris and electronic noise. Recirculation of red cells near the aperture should be prevented, as pulses produced may simulate those generated by platelets. Three techniques for setting thresholds have been used: (1) platelets can be counted between two fixed thresholds (e.g. between 2 and 20 fl); (2) pulses between fixed thresholds can be counted with subsequent fitting of a curve and extrapolation so that platelets falling outside the fixed thresholds are included in the computed count; and (3) thresholds can vary automatically, depending on the characteristics of individual blood samples, to make allowance for microcytic or fragmented red cells or for giant platelets. Factitiously low impedance platelet counts may be the result of giant platelets being identified as red cells or of EDTA-induced platelet clumping or satellitism (see p. 98). Misleadingly, high platelet counts may be due to markedly microcytic or fragmented red cells, to white cell fragments in leukaemia⁸³ or to bacteria or fungi.

An optical fluorescence platelet count has been introduced on some Sysmex analysers, in addition to the traditional impedance count.⁸⁴ A dye is used to stain the RNA/DNA of reticulocytes and platelet membranes and granules. The fluorescent staining of the platelets allows the exclusion of non-platelet particles from the count and also allows the inclusion of large or giant platelets. However, for samples from patients undergoing cytotoxic chemotherapy, the impedance count is sometimes more accurate. This is probably due to the erroneous staining of white cell fragments following apoptosis.⁸³ A switching algorithm has been designed on the instrument to report the most accurate platelet count, either optical or impedance.

An immunofluorescent method for platelet counting by flow cytometry has also been developed.⁸⁵ Platelets in a blood sample are labelled fluorescently with a specific monoclonal antibody or combination of antibodies and by measuring the RBC:platelet ratio the platelet count can be calculated. Suitable antibodies to platelet antigens are CD41, CD42 and CD61. This method using CD41 and CD61 has been adopted by the International Council for Standardization in Haematology as the reference method.⁸⁶ The Abbott Cell-Dyn and Sapphire instruments provide an automated immunological platelet count for diagnostic use. Although instruments can count platelets down to levels of 10 × 10⁹/l or less, it should be noted that precision at these levels is often poor with CVs of 22–66% being observed⁸⁷ and with counts below 10 × 10⁹/l differing appreciably between instruments and from the International Council for Standardization in Haematology reference method.⁸⁸

Reticulocyte count

Automated reticulocyte counts have been developed by using the fact that various dyes and fluorochromes combine with the RNA of reticulocytes.^40,103 Following binding of the dye, fluorescent cells can be enumerated using a flow cytometer. Most fully automated blood counters now incorporate a reticulocyte counting capacity so that use of a stand-alone reticulocyte counter is no longer necessary and use of a general purpose flow cytometer is no longer appropriate. An international standard for this method has been published by the Clinical and Laboratory Standards Institute (CLSI) in collaboration with ICSH.⁴⁴ The dyes used in the different systems include auramine O or polymethine with oxazine (Sysmex), thiazole orange (ABX), CD4K 530 (Abbott), as well as non-fluorescent dyes such as oxazine 750 (Siemens) and the traditional New methylene blue (Beckman Coulter, Abbott).

After staining, it is necessary to separate the reticulocytes from unstained red cells and, because the dyes also combine with DNA of nucleated cells, these cells must also be excluded. The threshold for this exclusion is determined by the intensity of fluorescence and particle sizing. Although the separation of reticulocytes from mature red cells is not always clear cut, automated reticulocyte counts correlate well with manual reticulocyte counts; however, absolute counts may differ because automated counts are dependent on the conditions of incubation and the method of calibrating the instrument.¹⁰³ Precision is much superior to that of the manual count because many more cells are counted, which has allowed reliable flagging of reticulocytopenia and the subjective element inherent in recognizing late reticulocytes is eliminated. Potential sources of inaccuracy are the inclusion of some leucocytes and platelets and, less often, Howell-Jolly bodies or malarial parasites in the ‘reticulocyte’ count.

Automated reticulocyte counts are fairly stable in blood that has been stored for 1–2 days at room temperature or up to 3–5 days at 4°C.

Point-of-care instruments

There are two types of technology to support point-of-care testing (POCT) (see p. 574), small bench top analysers and hand-held devices. The bench top systems are often smaller versions of laboratory analysers providing an FBC with red cell indices and either a five-part white cell differential or a partial three-part differential. Bench top analysers are equipped with automated calibration and quality control; however, they are too large for use at the patient’s bedside and are designed for use in clinics or small laboratories. It is recommended that instruments that employ primary sampling are used, rather than instruments that involve dilution of whole blood in the pre-analytical phase.¹¹² The most widely used test with a hand-held device is the measurement of haemoglobin concentration, but another device, using a disposable cartridge, has recently been introduced that measures haemoglobin concentration, counts leucocytes and platelets and performs a three-part differential on capillary blood. The range of equipment available will inevitably expand as more POCT is implemented. POCT devices should generate results that are comparable to those of the local reference laboratory. Internal quality control (IQC) must be available for all POCT instruments to detect significant deviations from acceptable performance. The analysis of control material before analysing patient samples can provide reassurance that the system is working correctly. At regular intervals parallel testing of a patient sample may be carried out at the POCT site and the main laboratory to ensure comparable results.

Ideally, there should also be an objective external method of quality assurance, external quality control (EQA). EQA involves the analysis of samples received from an accredited external source with undisclosed values; this could be from the supervising laboratory itself, from a manufacturer or from accredited national schemes. Results are subject to peer group assessment and statistical analysis to compare results across different sites.

Local haematologists/pathologists should encourage POCT users to participate in the supervising laboratory’s EQA.¹¹²

Haematologists should be aware of the potential for error if point-of-care blood gas analysers are used for estimation of Hb or Hct. Use of instruments that are based on conductivity measurements is not recommended since discrepancies of 20 g/l in the Hb and 0.04 l/l in the Hct can occur when the plasma protein concentration is low (e.g. if crystalloid has been used for blood replacement) and there is also a downward biases when the haematocrit is less than 0.30 l/l.¹¹² Furthermore, the reproducibility of Hct measurements can be poor. As long as there is adequate quality control, instruments based on spectrophotometry/co-oximetry agree more closely with laboratory measurements of Hb.¹¹²

Calibration of automated blood cell counters

The following methods are recommended for calibrating an automated blood cell counter:^113,114

For reasons of convenience and economy, control materials are commonly used as calibrants; but this practice is not recommended. Such materials are not sufficiently stable to serve as calibrants and their stated values are often approximations that are not assigned by reference methods. They are designed to give test results within a stated range over a stated period rather than a specific result.

The procedure for assigning values to fresh blood samples and indirectly to a stable calibrant is as follows:

To calibrate the automated counter directly from the three fresh blood samples, perform two counts with each sample and take the means. If the measured counts differ from those assigned, recalibrate the counter appropriately.

To calibrate a stable calibrant, perform two counts on the calibrant and on each fresh sample using the automated instrument, A, and take the means. From the ratio of the test results on fresh blood to those on the calibrator, assign corrected values to the calibrator by using the following calculations:

where:

Considerable care is required to ensure that the initial measurements on the fresh blood are as accurate as possible. Dilutions should be made with individually calibrated pipettes and grade A volumetric flasks. The cell counter should be calibrated as described on p. 39, with a signal-to-noise ratio of >100:1 and the count corrected for coincidence. Details of procedures to be used are described by the International Committee for Standardization in Haematology.¹¹⁵ Procedures for verification of the performance of multichannel analysers by the users have also been published by ICSH⁴⁶ and in the USA, by the National Committee for Clinical Laboratory Standards.¹¹⁶

Flagging of automated blood counts

‘Flagging’^117,118 refers to a signal that the specimen being analysed may have a significant abnormality because one or more of the blood count variables are outside specified limits (usually 2SD) or there is a qualitative abnormality that requires a quality control check and/or additional investigation. Abnormal cells have differing characteristics, such as nuclear size and granule content, from normal cells. The instrument detects a cell population as having an abnormal size or shape by cluster analysis. Under these circumstances, abnormal cell flags are generated to alert the user to the possibility of inaccurate results. Abnormal cells or interfering substances may render the automated differential inaccurate or unreportable. A blood film will need to be examined microscopically to verify the automated count and confirm the presence of abnormal cells. Most point-of-care bench top analysers have the ability to generate flags in the presence of abnormal cells or interfering substances; however, the range of alert flags available on these instruments is limited and their sensitivity and specificity may not be as good as those on more sophisticated laboratory haematology analysers. Although it is theoretically desirable for every blood count to include examination of a stained film, this has become impossible as a result of increasing workloads; time- and cost-effective rationalization has therefore been required. This has been helped by the availability of automated analysers that report differential leucocyte counts on every specimen. Consequently, significantly fewer blood films are now examined microscopically. Thus, a decision of when a blood film should be made, stained and examined should take account of flagging and the need to ensure analytic reliability. This includes a check of any significant changes from a recent previous count (delta check), as well as any specific clinical circumstances. There are previously published guidelines that describe blood film review criteria following FBC and differential analysis,¹¹⁹ which may be adapted to the individual laboratory’s needs. Box 3.1 is a guide to this selection.

Microscopy

Microscope Components

The main components of most routine microscopes are illustrated in Figure 3.9. The objectives are usually marked with their magnifying power, but older lenses may be marked by their focal length instead. The approximate equivalents are in Table 3.5.

Figure 3.9 Cross-section of microscope, showing its components. E, eyepiece; S, stand; O, objective; M, mechanical stage; C, condenser. The broken lines indicate the light path. Note that this shows an external light source being directed into the microscope. In most modern microscopes there is a built-in lamp in the base.

Table 3.5 Approximate equivalents of focal length to magnifying power

Focal length (mm)	Magnification
2	×100
4	×40
16	×10
40	×4

The working distance of the objective is the distance between the objective and the object to be visualized. The greater the magnifying power of the objective, the smaller the working distance (Table 3.6).

Table 3.6 Objective magnifying power and the associated working distance

Objective	Working distance
×10	5–6 mm
×40	0.5–1.5 mm
×100	0.15–0.20 mm

These specifications mean that when a coverslip is used, if it is too thick it will not be possible to focus at high magnification. Thus, the coverslip should be no more than 0.15 mm thick for examination of covered preparations by the ×100 oil-immersion objective. Furthermore, if the glass slide is too thick, this may prevent correct focus of the light path through the condenser to the object, as described later.

Setting Up the Microscope Illumination

1. If the microscope requires an external light source, using the mirror at the base, direct the light into the condenser. If the illumination is built in, make sure that the lamp voltage is turned down before switching on the microscope; then turn up the lamp until it is at c 70% of maximum power.

2. Place a slide of a blood film with a coverslip on the stage.

3. Lower the condenser, open the iris diaphragm fully and bring the preparation on the slide into focus with the ×10 objective.

4. Check that the eyepieces are adjusted to the operator’s interpupillary width and that the specimen is in focus for each eye by rotating the focusing mechanism on the adjustable eyepiece.

5. Close the diaphragm and raise the condenser slowly until the edge of the circle of light comes into sharp focus and there is a faint blue tinge at the edge of the diaphragm.

6. Using the condenser centering screws, adjust its position so that the circle of light is in the centre of the field.

7. Open the diaphragm completely so that light fills the whole field of view.

8. Remove the eyepieces, so that the upper lens of the objective is seen to be filled with a circle of light. Close the diaphragm slowly until the circle of light occupies about two-thirds of the surface.

9. Replace the eyepieces, refocus the specimen and if necessary readjust the condenser aperture and lamp brightness to obtain the sharpest possible image.

Examination of Slides

Low power (×10). Start with the objective just above the slide preparation. Then raise the objective with the coarse adjustment screw until a clear image is seen in the eyepiece. If there is insufficient illumination, rack up the condenser slightly.

High power (×40). Rack the condenser halfway down; lower the objective until it is just above the slide preparation. Use the coarse adjustment to raise the objective very slowly until a blurred image appears. Then bring into focus using the fine adjustment. If necessary, raise the condenser to obtain sufficient illumination.

Oil immersion (×100). Place a small drop of immersion oil on the part to be examined. Rack up the condenser as far as it will go. Lower the objective until it is in contact with the oil. Bring it as close as possible to the slide, but avoid pressing on the preparation. Look through the eyepiece and turn the fine adjustment very slowly until the image is in focus.

After using the oil-immersion objective, to avoid scratching the lens or coating the ×40 lens with oil, first swing the ×10 objective (or an empty lens space on the nosepiece) into place before removing the slide. As far as possible, use oil only when essential (e.g. for determining malaria species) and examine blood films for morphology or differential leucocyte count with the ×40 lens without oil.

If you cannot focus using the oil-immersion lens, consider that:

The coverslip may be too thick

Two coverslips may have been accidentally applied

You might have inverted the slide.

Routine Maintenance of the Microscope

The microscope is a delicate instrument that must be handled gently. It must be installed in a clean environment away from chemicals, direct sunlight, heating sources or moisture. If the stage is contaminated with saline, it must be cleaned immediately to avoid corrosion. Even in a temperate climate, humidity and high temperatures cause growth of fungus, which can damage optical surfaces. Because storage in a closed compartment encourages fungal growth, do not store the microscope in its wooden box, but keep it standing on the bench protected by a light plastic cover.

After use of the microscope, wipe the oil-immersion objective with lens tissue, absorbent paper, soft cloth or medical cotton wool. If other lenses are smeared with oil, wipe them with a little toluene or a solution of 40% petroleum ether, 40% ethanol and 20% ether.

Lenses must never be soaked in alcohol because this may dissolve the cement.

Clean non-optical parts with mild detergent and remove grease or oil with petroleum ether, followed by 45% ethanol in water. Remove dust from the inside and outside of the eyepieces with a blower or soft camel-hair brush.

Clean the condenser in the same way as the lenses with a soft cloth or tissue moistened with toluene and clean the mirror (if present) with a soft cloth moistened with 5% alcohol. The iris diaphragm is very delicate and if damaged or badly corroded it is usually beyond repair.

Never force the controls. If movement of the focusing screws or mechanical stage becomes difficult, lubricate them with a small drop of machine oil. All accessible moving parts should be cleaned occasionally and given a touch of oil to protect against corrosion. Do not use vegetable oils because they become dry and hard. Always keep the surface of the fixed stage dry because moving wet slides requires increased force, which may damage the mechanical stage.

For care of microscopes in hot humid and hot dry climates, see p. 606.