Tests for the acute-phase response

Inflammatory response to tissue injury includes alteration in serum protein concentration, especially increases in fibrinogen, haptoglobin, caeruloplasmin, immunoglobulins (Ig) and C-reactive protein (CRP), and decrease in albumin. The changes occur in acute infection, during active phases of chronic inflammation, with malignancy, in acute tissue damage (e.g. following acute myocardial infarction) with physical injury. Measurement of the acute-phase response is a helpful indicator of the presence and extent of inflammation or tissue damage and response to treatment. The usual tests are estimation of CRP and measurement of the erythrocyte sedimentation rate (ESR); some studies have suggested that plasma viscosity is also a useful indicator, but there is debate on the relative value of theses tests.^1,2

Kits that are sensitive and precise are available for CRP assay; small increases in serum levels of CRP can often be detected before any clinical features become apparent, whereas as a tissue-damaging process resolves, the serum level rapidly decreases to within the normal range (<5 mg/l). The ESR is slower to respond to acute disease activity and it is insensitive to small changes in disease activity. It is less specific than CRP because it is also influenced by immunoglobulins (which are not acute-phase reactants) and by anaemia. Moreover, because the rate of change of ESR is slower than that of CRP, it rarely reflects the current disease activity and clinical state of the patient as closely as the CRP. The ESR is a useful screening test, and the conventional manual ESR method is simple, cheap and not dependent on power supply, thus making it suitable for point-of-care (near-patient) testing. It is recommended that, in clinical practice, both tests should be carried out in tandem.³ Because CRP assay is a biochemical test usually performed in the clinical chemistry laboratory, it will not be discussed further here.

Erythrocyte Sedimentation Rate

The method for measuring the ESR recommended by the International Council for Standardization in Haematology (ICSH)⁴ and also by various national authorities⁵ is based on that of Westergren, who developed the test in 1921 for studying patients with pulmonary tuberculosis. ESR is the measurement of the sedimentation of red cells in diluted blood after standing for 1 h in an open-ended glass tube of 30 cm length mounted vertically on a stand.

Conventional Westergren Method

The recommended tube is a straight glass or rigid transparent plastic tube 30 cm in length and not less than 2.55 mm in diameter. The bore must be uniform to within 5% throughout. A scale graduated in mm extends over the lower 20 cm. The tube must be clean and dry and kept free from dust. If reusable, before being reused it should be thoroughly washed in tap water, then rinsed with deionized or distilled water and allowed to dry. Specially made racks with adjustable levelling screws are available for holding the sedimentation tubes firmly in an exactly vertical position. The rack must be constructed so that there will be no leakage of the blood from the tube. It is conventional to set up sedimentation-rate tests at room temperature (18–25°C). Sedimentation is normally accelerated as the temperature increases, and if the test is to be carried out at a higher ambient temperature, a normal range should be established for that temperature. Exceptionally, when high thermal amplitude cold agglutinins are present, sedimentation becomes noticeably less rapid as the temperature is increased toward 37°C.

For the diluent, prepare a solution of 109 mmol/l trisodium citrate (32 g/l Na₃C₆H₅O₇.2H₂O). Filter through a micropore filter (0.22 mm) into a sterile bottle. It can be stored for several months at 4°C but must be discarded if it becomes turbid through the growth of moulds.

Method

The method described below, originally described by the International Council for Standardization in Haematology (ICSH)⁴ and now adopted by the Clinical and Laboratory Standards Institute (CLSI) as its approved method,⁵ is intended to provide a reference method for verifying the reliability of any modification of the test.

Either collect venous blood in ethylenediaminetetra-acetic acid (EDTA) and dilute a sample accurately in the proportion of 1 volume of citrate to 4 volumes of blood, or collect the blood directly into the citrate solution. The test should then be carried out on the diluted sample within 4 h of collecting the blood, although a delay of up to 6 h is permissible provided that the blood is kept at 4°C. EDTA blood can be used within 24 h if the specimen is kept at 4°C, provided that 1 volume of 109 mmol/l (32 g/l) trisodium citrate is added to 4 volumes of blood immediately before the test is performed.

Mix the blood sample thoroughly and then draw it up into the Westergren tube to the 200 mm mark by means of a teat or a mechanical device; mouth suction should never be used. Place the tube exactly vertical and leave undisturbed for exactly 60 min, free from vibrations and draughts and not exposed to direct sunlight. Then read to the nearest 1 mm the height of the clear plasma above the upper limit of the column of sedimenting cells. The result is expressed as ESR = X mm in 1 h. A poor delineation of the upper layer of red cells may sometimes occur, especially when there is a high reticulocyte count.

Range in health

The mean values and the upper limit for 95% of normal adults are given in Table 6.1. There is a progressive increase with age, but it is difficult to define a strictly healthy population for determining normal values in individuals older than 70 years.⁶ In the newborn, the ESR is usually low. In childhood and adolescence, it is the same as for normal men with no differences between boys and girls. It is increased in pregnancy, especially in the later stages, and independent of anaemia;⁷ this is due to the physiological effect of haemodilution (an increase in the plasma volume)

Table 6.1 Erythrocyte sedimentation rate ranges in health

Age range (years)	ESR mean MM IN 1 H
10–19	8
20–29	10.8
30–39	10.4
40–49	13.6
50–59	14.2
60–69	16
70–79	16.5
80–91	15.8
Pregnancy
Early gestation	48 (62 if anaemic)
Later gestation	70 (95 if anaemic)

In the newborn, the ESR has been reported to be 0–2 mm in 1 h, increasing to 4 mm in 1 h at 1 week, up to 17 mm in 1 h by day 14, and then 10–20 mm in 1 h for both girls and boys, until puberty.⁸ However, as the studies in infants were obtained by the capillary method, they are not strictly comparable to the Westergren method.

Modified methods

Length of tube

The overall length of the tube is not a critical dimension for the test provided that it fits firmly in an appropriate holding device. The tube must, however, be long enough to ensure that packing of the cells does not start before the test has been completed.

Plastic glass tubes

A number of plastic materials (e.g. polypropylene and polycarbonate) are recommended as substitutes for glass in Westergren tubes. Nevertheless, not all plastics have similar properties, and it must be demonstrated that the ESR with the chosen tubes is reproducible and not affected by the plastic.

Disposable glass tubes

Disposable glass tubes should be supplied clean and dry and ready for use. It is necessary to show that neither the tube material nor the manufacturer’s cleaning process affect the ESR.

Capillary method

Short tubes of narrower bore than in the standard tube are available mainly for tests on infants. These are, however, no longer in general use, and it is necessary to establish normal ranges or a correction factor to convert results to an approximation of ESR by the Westergren method.

Time

Sedimentation is measured after aggregation has occurred and before the cells start to pack, usually at 18–24 min. From the rate during this time period the sedimentation that would have occurred at 60 min is derived and converted to the conventional ESR equivalent by an algorithm.⁹

Sloping tube

Red cells sediment more quickly when streaming down the wall of a sloped tube. This phenomenon has been incorporated into automated systems in which the end-point is read after 20 min with the tube held at an angle of 18° from the vertical.¹⁰ Incorporating a low-speed centrifugation step (approx. 800 rpm) in this automated method reduces the end-point time further.¹¹ These have been shown to give results comparable to the conventional method.

Anticoagulant

EDTA blood can be used without citrate dilution, at least if packed cell volume (PCV) is below 0.36 (haemoglobin <110 g/l); less precise results are obtained when the PCV is higher. Because of the biohazard risk of blood contamination inherent in using open-ended tubes, it is now recommended that, where possible, a closed system be used in routine practice. Manual methods are available that avoid transfer of the blood into the sedimentation tube. Automated closed systems use either blood collected in special evacuated tubes containing blood anticoagulated with citrate or EDTA. A sample is taken up through a pierceable cap and then automatically diluted in the system if this is required.

Whenever a different method or tube is planned, a preliminary test should be carried out to check precision and to compare results with those obtained by the CLSI standardized method described earlier.

Evaluation of a new routine method

For the evaluation of a new ESR method the test should be carried out on at least 60 samples. These samples could be collected separately from the same subjects in accordance with specified requirements (e.g. directly into tubes containing citrate or undiluted EDTA samples). The samples should come from patients with a wide variety of diseases (as well as from normal subjects) to cover the range of ESR from 15 to 105 mm, with approximately the same number of samples in each quartile. If any test fails to give a clear-cut plasma erythrocyte interface in either the test system or the standardized test, the pair of values should be eliminated from the data.

Any new method may be considered to be satisfactory if 95% of results differ consistently by no more than 5%. However, because the ESR may be affected by several uncontrolled variables, the reference method cannot be used to adjust the measurements that are obtained. Thus if the new method gives disparate readings, it will be necessary to establish a normal range specifically for the method.

Quality Control

The standardized method can be used as a quality-control procedure for routine tests or alternatively stabilized whole blood preparations which are now available are suitable as a daily control for use with automated systems (e.g. ESR-Chex).¹² Three or four specimens of EDTA blood kept at 4°C will also serve as a control on the following day.

Another control procedure is to calculate the daily cumulative mean, which is relatively stable when at least 100 specimens are tested each day in a consistent setting (see Chapter 25). A coefficient of variation of <15% between daily sets appears to be a satisfactory index for monitoring instrument performance.¹³

Semiquantitative Slide Method

Enhanced red cell adhesion/aggregation can be demonstrated by allowing a drop of citrated blood to dry on a slide. An estimate of the amount of cell aggregation on the film by image analysis provides a semiquantitative measure of the acute-phase response that appears to correlate with the ESR.¹⁴ Based on this principle, serial microscopic images of red cells aggregating on a glass slide taken every 30 s for 5 min can distinguish a normal ESR from a high value (Figs 6.1, 6.2). The images demonstrate greater spacing of cells in blood with higher ESR values compared with blood with lower ESR values.

Figure 6.1 ESR of 9 mm in 1 h after 2 min.

Figure 6.2 ESR of 69 mm in 1 h after 2 min.

Mechanism of Erythrocyte Sedimentation

The rate of fall of the red cells is influenced by a number of interacting factors. It depends on the difference in specific gravity between red cells and plasma, but it is influenced very greatly by the extent to which the red cells form rouleaux, which sediment more rapidly than single cells. Other factors that affect sedimentation include the ratio of red cells to plasma (i.e. the PCV), the plasma viscosity, the verticality or otherwise of the sedimentation tube, the bore of the tube and the dilution (if any) of the blood.

The all-important rouleaux formation and the red cell clumping that are associated with the increased ESR are mainly controlled by the concentrations of fibrinogen and other acute-phase proteins (e.g. haptoglobin, ceruloplasmin, α₁-acid-glycoprotein, α₁-antitrypsin, and CRP). Rouleaux formation is also enhanced by the immunoglobins and is retarded by albumin. Defibrinated blood normally sediments extremely slowly (i.e. not more than 1 mm in 1 h) unless the serum globulin concentration is increased or there is an unusually high globulin:albumin ratio.

Anaemia, by altering the ratio of red cells to plasma, encourages rouleaux formation and accelerates sedimentation. In anaemia, cellular factors may also affect sedimentation. Thus, in iron deficiency anaemia, a reduction in the intrinsic ability of red cells to sediment may compensate for the accelerating effect of an increased proportion of plasma.

Sedimentation can be observed to take place in three stages: a preliminary stage of at least a few minutes during which time rouleaux occur and aggregates form; then a period in which the sinking of the aggregates takes place at a constant speed; and finally, a phase during which the rate of sedimentation slows as the aggregated cells pack at the bottom of the tube. It is obvious that the longer the tube used, the longer the second period can last and the greater the sedimentation rate may appear to be.

Although ESR is a non-specific phenomenon, its measurement is clinically useful in disorders associated with an increased production of acute-phase proteins. In rheumatoid arthritis or tuberculosis it provides an index of progress of the disease, and it is of considerable value in diagnosis of temporal arteritis and polymyalgia rheumatica. It is often used if multiple myeloma is suspected, but when the myeloma is non-secretory or light chain, a normal ESR does not exclude this diagnosis.

An elevated ESR occurs as an early feature in myocardial infarction.¹⁵ Although a normal ESR cannot be taken to exclude the presence of organic disease, the vast majority of acute or chronic infections and most neoplastic and degenerative diseases are associated with changes in the plasma proteins that lead to an acceleration of sedimentation. An increased ESR in subjects who are HIV seropositive seems to be an early predictive marker of progression toward acquired immune deficiency syndrome (AIDS).¹⁶ The ESR is less helpful in countries where chronic diseases are rife; however, one study has shown that very high ESRs (higher than 100 mm/h) have a specificity of 0.99 and a positive predictive value of 0.9 for an acute or chronic infection.¹⁷ The ESR is influenced by age, stage of the menstrual cycle and medications taken (corticosteroids, contraceptive pills). It is especially low (0–1 mm) in polycythaemia, hypofibrinogenaemia and congestive cardiac failure and when there are abnormalities of the red cells such as poikilocytosis, spherocytosis, or sickle cells. In cases of performance-enhancing drug intake by athletes (discussed below) the ESR values are generally lower than the usual value for the individual and as a result of the increase in haemoglobin (i.e. the effect of secondary polycythaemia).

Whole blood viscosity

The viscosity of whole blood reflects its rheological properties; it is influenced by PCV, plasma viscosity, red cell aggregation and red cell deformability. It is especially sensitive to PCV, with which it is closely correlated. The clinical interpretation of its measurement must also take into account the interaction of the red cells with blood vessels, which greatly influences blood flow in vivo.

Guidelines for measuring blood viscosity and red cell deformability by standardized methods have been published.²¹ Rotational and capillary viscometers are suitable for measuring blood viscosity; deformability can be measured by recording the rate at which red cells in suspension pass through a filter with pores 3–5 mm in diameter.

Heterophile antibodies in serum: diagnosis of infectious mononucleosis

Infectious mononucleosis (IM) is caused by Epstein–Barr virus.²² The immune response that develops in response to virus-infected cells includes not only antibodies to viral antigens but also characteristic heterophile antibodies. Before the nature of this reaction was understood, Paul and Bunnell²³ demonstrated the antibodies as agglutinins directed against sheep red cells. They are, in fact, not specific for sheep red cells but also react with horse and ox, but not human, red cells. They are IgM globulins, which are immunologically related to, but distinct from, antibodies that occur in response to the Forssman antigens. The latter are widely spread in animal tissue; they occur at low titre in healthy individuals and at high titre in serum sickness and in some leukaemias and lymphomas.^24,25 In these non-IM conditions, the antibody can be absorbed out by guinea pig cells. Thus, for the diagnosis of IM, it is necessary to demonstrate that the antibody present has the characters of the Paul–Bunnell antibody (i.e. it is absorbed by ox red cells but not by guinea pig kidney). This is the basis of the absorption tests for IM (‘monospot’ test). Immunofluorescent antibody tests have been developed to distinguish the IgM antibody, which occurs at high titre in the early phase of IM and diminishes during convalescence, from the IgG antibody, which persists at high titre for years after infection^26,27 and which also occurs in the non-IM infections.^22,28

Screening Tests for Infectious Mononucleosis

The reagents for IM screening are available commercially in diagnostic kits from several manufacturers. Guinea pig cells can be also be manufactured locally as described in previous editions. Some kits are based on agglutination of stabilized horse red cells or antigen-coated latex particles to which IM antibody binds. An extensive evaluation of 14 slide tests for the UK Medical Devices Agency (MDA), showed them to have a sensitivity between 0.87 and 1.00 and specificity of 0.97 to 1.00, with an overall accuracy (positive and negative) in the order of 91–100%.²⁵ False-positive reactions have been reported in malaria, toxoplasmosis, and cytomegalovirus infection; autoimmune diseases; and even occasionally without any apparent underlying disease.^29,30 False-negative reactions occur if the test is carried out before the level of heterophile antibody has increased or conversely when it has decreased. False-negative reactions may also occur in the very young and the very old. In the UK MDA study the best performance was obtained with the Clearview test (Unipath), which uses latex-labelled bovine erythrocyte glycoprotein. IM heterophile antibody binds to this to form a complex that presents as a band in the result window (Fig. 6.3). The test can be performed with diluted whole blood as well as with plasma or serum.

Figure 6.3 Screening for infectious mononucleosis with Clearview slide test. The sample is added to the bottom window, where, if positive, antibodies combine with bovine erythrocyte glycoprotein attached to blue microspheres; migration to the test window (centre) then occurs and here the complexes are bound to immobilized bovine erythrocytic glycoprotein, producing a blue line. The blue line in the upper window is a positive control.

Screening tests are also available based on enzyme-linked immunosorbent assay (ELISA) and immunochromatographic assay. These tests are more elaborate than the slide screening test described earlier, but they are less likely to give a false result.

Clinical Value

Tests for the heterophile antibody are useful for diagnosis. Antibodies are often present as early as the 4th to 6th day of the disease and are almost always found by the 21st day. They disappear as a rule within 4–5 months. There is no unanimity as to how frequently negative reactions are found in ‘true’ IM. Occasionally, the characteristic antibodies develop very late in the course of the disease, perhaps weeks or even months after the patient becomes ill. It is also known that a positive reaction may be transient and that the antibodies may be present at such low titres that they may be missed or may produce anomalous agglutination reactions when associated with the naturally occurring antibody at similar titres. For all of these reasons, it is difficult to state categorically that any particular patient has not or will not produce antibodies. Antibodies specific for Epstein–Barr virus have been demonstrated in the serum of 86% of patients with clinical and/or haematological features of IM.³¹

There is no substantial evidence that sera containing agglutinins in high concentration giving the typical reactions of IM are ever found in other diseases uncomplicated by IM. In particular, the heterophile antibody titres in the lymphomas are similar to those found in unselected patients not suffering from IM.³²

Demonstration of lupus erythematosus cells

Antinuclear antibodies, or antinuclear factors (ANF), occur in the serum in a wide range of autoimmune disorders, including systemic lupus erythematosus (SLE). Descriptions of the LE cell test, which has now been superseded by immunological tests, can be found in the 8th and earlier editions of this book.

Erythropoietin

Erythropoietin regulates red cell production. It is a heat-stable glycoprotein with a molecular weight of about 34 kDa. It is produced mainly in the kidney. Only a small quantity is demonstrable in normal plasma or urine.

A pure form of human erythropoietin from recombinant DNA (r-HuEpo) is available for diagnostic assay methods by ELISA, enzyme immunoassay and radioimmune assay. Commercial kits are available that are reliable and sensitive,³³ although there is some inter-method variability.³⁴ Results are expressed in international units by reference to an international (WHO) standard. This was originally a urinary extract, and a preparation is available with a potency of 10 iu per ampoule.³⁵ The present standard has been established for r-HuEpo with a potency of 86 iu per ampoule.³⁶

Autonomous in vitro erythropoiesis

When mononuclear cells from blood or bone marrow are cultured, erythroid colonies (CFU-E) will normally develop only when erythropoietin is present in the culture medium. However, growth will occur in erythropoietin-free medium in primary polycythaemia. This provides a method for distinguishing primary from secondary polycythaemia.^45–47

Mononuclear cells are collected from a blood sample by density separation (see p. 65) and added to an appropriate serum-free liquid culture medium^46,47 or collagen gel medium,⁴⁸ which is then divided into two portions. To one portion is added 1 iu/ml of erythropoietin. Both portions are plated and incubated for 7 days at 37°C. They are then stained with benzidine and examined directly under an inverted microscope or after spreading onto slides. The numbers of benzidine-positive cell clusters in the erythropoietin-free and erythropoietin-containing samples are counted and compared. A diagnosis of primary polycythaemia is indicated if there is an approximately equal growth in both samples. A method has been described in which flow cytometry with immunofluorescence is used to detect growth of the erythroid cells after only 2–5 days of culture.⁴⁹

Thrombopoietin

Thrombopoietin regulates megakaryocyte development and platelet production. It is a protein produced by the liver and has been purified from serum.⁵⁰ It is considerably larger than erythropoietin, with a molecular weight of about 335 kDa. A recombinant human thrombopoietin (rhTPO) has been produced and used to prepare a monoclonal antibody and develop a sensitive and specific ELISA. This has been used to measure thrombopoietin in normal serum and serum from patients with various blood disorders.⁵¹ The normal range (mean and 2SD) is 0.79 ± 0.35 fmol/ml for men and 0.70 ± 0.26 fmol/ml for women. It is increased in thrombocytopenias and is especially high (18.5 ± 12.4 fmol/ml) in aplastic anaemia with severe thrombocytopenia. In essential thrombocythaemia, thrombopoietin is in the range of 1.01–4.82 fmol/ml.⁵¹

Haematological tests in sports medicine

Assays to detect the illicit use of hormones such as erythropoietin in endurance sports are becoming increasingly important.^52,53 Some of the current test methods for detecting erythropoietin used in this manner are described below.

Blood doping, often called induced erythrocythemia, is the practice of increasing the number of red blood cells through an intravenous infusion of blood, thereby enhancing performance in sports through the increase in red blood cell mass with an increased oxygen-carrying capacity.⁵² Alternatively, recombinant hormones such as erythropoietin can be used to induce erythropoiesis. Blood doping is an illegal practice as it provides an unfair advantage of endurance and performance over other athletes;^52,54 It is also potentially dangerous due to the abnormal increase in red cell mass and the risk of acquiring infection from contaminated blood.⁵⁴ The International Doping Test and Management⁵⁵ collaborates with the World Anti-Doping Agency and other international and national sports authorities to coordinate the collection of blood and urine samples to be analysed in accredited laboratories.⁵⁶

The full blood count, which includes the reticulocyte count, haemoglobin and packed cell volume, is the cheapest and most effective way to screen for suspected use of erythropoietin⁵⁵ as abnormal blood counts may suggest doping. The sports administrative bodies collect blood profiles on individual athletes and these are monitored over time so changes in blood parameters due to doping can be detected. Compared to population-derived upper limits of the reference range such as the 0.5 Hct limit or the 2.5% reticulocyte limit, the fluctuations allowed are smaller when using individual-based cut-offs.^52,54 Blood samples need to be analysed within 24 h of venesection⁵² and the security and confidentiality of blood samples obtained should be maintained at all times. The abnormalities in the blood profile which may suggest doping are outlined in the following section.

Principles of parasite detection

The essential method for a definitive diagnosis of malaria remains the finding of parasites in a blood film, followed by the identification of the particular species by morphology. Microscopic diagnosis can be a satisfying process in which parasites are identified then are assigned to their species based on fine morphological detail. Examination of thick or thin blood films remains the most sensitive routinely applied technique where microscopic expertise is available. Only brief outlines of how microscopic diagnoses may be reached are given in this chapter, and for more detailed accounts, readers are referred to a parasitology textbook. Where microscopic expertise is not available, antibody-based ‘rapid diagnostic tests’ (RDTs) are increasingly applied. RDTs have widespread application in developing countries, but increasingly are gaining use in all laboratories to screen potentially infected samples and to supplement microscopic diagnosis. It is important, however, that users appreciate the strengths and limitations that govern selection and interpretation of RDTs. Molecular approaches to malaria diagnosis and in particular to species identification, employ PCR-based assays and have the capability to be highly sensitive and specific.⁶³Cost and time constraints, however, mean that at present molecular techniques are employed principally in reference or research laboratories. Molecular techniques in parasite diagnosis are not discussed in this text.

In addition to the Plasmodia that give rise to malaria, the other important parasites to be found in the blood are leishmaniae, Babesia, trypanosomes, microfilaria, and ehrlichiae. Thick blood films are also useful for the detection of microfilaria. When they are used for this purpose, it is important to scan the entire film using a low-power objective, or parasites may be missed. Examination of wet preparations of blood can also be used for diagnosis of microfilariae and has the advantage that the parasites are easily detected because they are moving. A stained film is necessary for confirmation of species. Wet preparations are also useful for the detection of trypanosomes and the spirochaetes of relapsing fever. The presence of small numbers of trypanosomes or spirochaetes is revealed by occasional slight agitation of groups of red cells. Examination of a stained film confirms their nature.

Examination of blood films for parasites

Staining Thin Films

Thin films should be stained with Giemsa’s stain or Leishman’s stain at pH 7.2, not with a standard May–Grünwald–Giemsa stain. The pH is important to maximize the visibility of cytological features important for diagnosis (Fig. 6.4).

Figure 6.4 Effects of staining pH on parasite staining characteristics (P. vivax). At pH 7.2 (A) the parasite is readily distinguished from the adjacent blue/grey erythrocytes, chromatin staining is prominent and Schüffner’s dots are readily distinguished. At pH 6.9 (B) the parasite remains visible, but there is less contrast from surrounding erythrocyte cytoplasm, relatively weak chromatin staining, and poor resolution of Schüffner’s dots (which may not be visible at this pH).

Microscopic diagnosis of malaria

Morphological criteria for differentiation of malaria parasites are given in Table 6.2 and illustrated in Figures 6.5–6.8. Mixed species infections are well recognized, and microscopists should be alert to morphological features that cannot be reconciled to a single disease type (Fig. 6.9). Films for malaria must be made no longer than 3–4 h after blood collection. Films prepared later may still reveal parasites, but the parasite morphology will differ from fresh samples making species identification more difficult and in some cases causing the appearances of parasite stages not normally occurring in blood (Fig. 6.10).

Figure 6.5 Morphology of P. falciparum. Note that erythrocytes throughout this series are not enlarged or distorted. Early trophozoites (A) show high parasitaemia, with accolé (appliqué edge) forms, multiply infected erythrocytes and double chromatin dot forms. The late trophozoite (B) shows two thickened ring forms with characteristic Maurer’s dots (clefts) in the erythrocyte cytoplasm. A typical crescent (‘banana-shaped’) gametocyte is shown (C). Schizonts are not shown since they rarely circulate in P. falciparum infection.

Figure 6.6 Morphology of P. malariae. Early trophozoites (A) are small but less fine than those of P. falciparum; in this case the ring is irregular and chromatin dot is within the main parasite cytoplasm. A late trophozoite (B) spans the erythrocyte as a thick ‘band form’. A characteristic ‘daisy head’ schizont (C) has eight merozoites arrayed around the central pigment. Finally, the gametocyte (D) is typically small and does not fill the normally sized erythrocyte.

Figure 6.7 Morphology of P. vivax. The red cells are enlarged, distorted and contain visible Schüffner’s dots in all these images. Note that the parasites are larger than in the other malarial forms. The early trophozoite (A) in this figure has the form of a thick ring with a large chromatin dot, but the later trophozoite (B) has a distorted ‘amoeboid’ form. Note that the large macrogametocyte (C) fills the enlarged erythrocyte.

Figure 6.8 Morphology of P. ovale. Distinguishing P. ovale from P. vivax is perhaps the most difficult morphological distinction in malaria diagnosis. For both species, the red cells are enlarged, distorted, and contain visible Schüffner’s dots. The features that help distinguish these forms are given in Table 6.2. In this figure, note that the parasites and erythrocytes are smaller than for P. vivax, and that in some cases the cytoplasm is fimbriated. Both early (A) and late trophozoites (B) have coarse ‘ring’ forms. The schizont (C) contains fewer than 16 developing merozoites. A microgametocyte (D) does not fill the erythrocyte and numerous dots can be seen in the erythrocyte cytoplasm.

Figure 6.9 Mixed infection of P. falciparum and P. ovale. Diagnosis depends on recognition of features that cannot be reconciled with a single malarial species. (A) A typical fine double dot ring form of P. falciparum coexists with an enlarged ovoid erythrocyte containing a thickened parasite and numerous Schüffner’s dots (P. ovale). (B) Gametocytes from the same two species with contrasting appearances and causing different effects on the host erythrocyte.

Figure 6.10 Delayed film preparation. These gametocytes of P. falciparum have very atypical morphology: showing rounding up and clumping (A) or exflagellation of microgametocytes (B). Biologically, such changes occur within the mosquito stomach during sexual reproduction of the parasite; when present on a blood film they indicate a delay in preparation.

Two other morphology-based screening methods can be used for malaria diagnosis.

Rapid diagnostic tests for malaria

Rapid diagnostic tests (RDTs) for malaria employ specific antibodies that detect malaria antigens in the blood of infected individuals. RDTs use small blood samples obtained by finger prick or by venepunture and employ a ‘lateral diffusion’ system similar to a pregnancy test to generate results. RDTs therefore display results in the form of visible ‘bands’ that can be interpreted by non-expert users with limited facilities. In general, a blood specimen to be tested (2–50 μl) is lysed in buffer solution containing one or more malaria-specific ‘detection antibodies’. The detection antibody is coupled to a visually observable label. Where specific antigen is present, a complex is formed between that antigen and its cognate labelled antibody. The labelled antigen–antibody complex generated is then bound by a second ‘capture-antibody’ that recognizes the same antigen, and which is immobilized as a line on the test strip. A positive result therefore generates a visible line of antigen–antibody complex. A separate immobilized capture antibody recognizes the labelled detection antibody alone; this control band will produce a line in the absence of malaria antigen and confirms that the test has been performed correctly and the result can be interpreted (Fig. 6.11).⁷²

Figure 6.11 Rapid diagnostic tests for malaria. An RDT of the ‘three line’ format, the image shows the output window. (A) The results of a negative test in which the positive control line shows a positive band to indicate the technical validity of the result. (B) The positive lines in all three positions, i.e. a technically interpretable result (control line positive), with P. falciparum-specific antigen detected (line 1), and pan-malarial antigen detected (line 2). This patient had a single-species infection (P. falciparum), although the test does not allow a mixed infection to be excluded.

The antigen targets detected by RDTs fall into two groups. The first group of antigens are expressed in all malarial species. Antigens from this group therefore confirm malarial infection is present, but do not allow the parasite species to be determined. Antigens from this group are Plasmodium aldolase (PMA) or parasite lactate dehydrogenase (pLDH). The second group of antigens are specific for P. falciparum. Antigens from this group are histidine-rich protein-2 of P. falciparum (pfHRP2) or a P. falciparum-specific form of LDH (pfLDH). Antibodies from the two groups are used individually, or in combination, to produce two different test formats. A ‘two line’ test uses an anti-pfHRP2 band together with the positive control band to recognize P. falciparum only. A ‘three line’ test uses a P. falciparum specific antibody band, together with PMA or pLDH, together with the positive control antibody. The three line test can therefore indicate the presence of P. falciparum, or if P. falciparum is not present the test will indicate the presence of other malarial species. Since the second antibody is pan-malarial, the three line test does not distinguish between infection by P. vivax, P. ovale, or P. malariae, and will not detect mixed infection. For field diagnosis in malaria-endemic areas the selection of a two line (P. falciparum only) test, or three line (pan-malarial) test, will depend on local species prevalence. It is considered that if >90% of malaria cases in an area are caused by P. falciparum an RDT that detects only that species is appropriate for use. The introduction of such tests in malaria endemic areas also requires careful consideration of ease of use; cost; and limitations imposed by transport, distribution and storage.⁶⁸

In terms of diagnostic sensitivity, WHO requires RDTs to reliably detect infections of 100 parasites per microlitre of blood (95% sensitivity).⁶⁹ This is equivalent to the diagnostic sensitivity reasonably expected of a field microscopist diagnosing malaria in endemic regions.⁷⁰ Sensitivity becomes less reliable below 100 parasites per microlitre, and this contrasts with the ‘gold standard’ sensitivity achieved by an expert microscopist in good conditions, who should detect 5–10 parasites per microlitre.⁷⁰ Sensitivity does depend on species. For P. falciparum parasites, RDT sensitivity frequently exceeds 100 parasites per microlitre, although genetic variation of P. falciparum antigens may reduce sensitivity in some instances.⁶⁹ For other malarial species, sensitivity of detection is recognized to be less good, particularly for P. ovale and P. malariae where RDTs may not detect infections that are clinically significant. There is also a recognized incidence of false-positive reactions caused by cross-reaction with autoantibodies (particularly with rheumatoid factor) that varies between detection systems. In all instances, malaria antigens are recognized to persist for a number of weeks following successful treatment, and in these circumstances a positive result may not indicate current infection.

The lower sensitivity of RDTs when compared with expert microscopic diagnosis means that all positive and negative results should be confirmed using microscopy. Symptomatic parasitaemia in non-immune subjects may occur with fewer than 100 parasites per microlitre of blood, and repeat testing of negative samples may be required to confirm a diagnosis. Users should also be aware that the positive line on the test only indicates a correctly performed test, and does not confirm effectiveness of parasite detection or accuracy of the test. Awareness of diagnostic performance for the selected test in the user’s laboratory is therefore mandatory.⁶⁹

Leishmaniasis

Leishmania species are transmitted by the bite of an infected female sandfly and are associated with a variety of clinical conditions including visceral and mucocutaneous leishmaniasis. Visceral leishmaniasis may present to the haematologist as splenomegaly, hepatomegaly, fever, lymphadenopathy or pancytopenia, and it is being increasingly reported in patients with HIV infection. Serological studies are recommended as the initial diagnostic tests in suspected leishmaniasis. In advanced stages of the disease, parasites can be found in phagocytic cells in spleen, lymph nodes, bone marrow and peripheral blood. Culture of aspirated bone marrow is, however, a more sensitive diagnostic technique than microscopy.

Diagnosis of Leishmaniasis in the Haematology Laboratory

Leishmaniasis is diagnosed in the haematology laboratory by direct visualization of the amastigotes (often referred to as Leishman–Donovan bodies). Buffy coat preparations of peripheral blood or aspirates (see p. 64 for preparation of buffy coats) from marrow, spleen, lymph nodes or skin lesions should be spread on a slide to make a thin smear and stained with Leishman’s or Giemsa’s stain (pH 7.2) for 20 min (see p. 66). Amastigotes are seen within monocytes or, less commonly, in neutrophils in peripheral blood and in macrophages in bone marrow aspirates. They are small, round bodies 2–4 mm in diameter with indistinct cytoplasm, a nucleus, and a small rod-shaped kinetoplast (Fig. 6.12A). Occasionally, amastigotes may be seen lying free between cells.

Figure 6.12 Bone marrow and blood parasites. (A) Leishmaniasis (Leishman–Donovan bodies); (B) African trypanosomiasis; (C) American trypanosomiasis (T. cruzi); and (D) microfilaria.

Trypanosomiasis

Diagnosis of Trypanosomiasis in the Haematology Laboratory

Care should be taken when handling samples suspected of being infected with trypomastigotes because infection can occur if the organisms penetrate the skin. Several techniques are available for examining specimens for the presence of trypomastigotes.

Wet Preparations

If present in high concentrations, trypomastigotes can be seen thrashing among the cells on a fresh, unstained wet preparation of blood or lymph node fluid. Preparations should be examined within 4 h of sampling (this time can be extended if a few milligrams of glucose are added to the specimen) using a ×40 objective and a partially closed condenser iris or dark-field or phase contrast microscopy.

Thick Blood Films or Chancre Aspirates

Examination of a thick film allows more of the sample to be examined rapidly, but T. cruzi are easily damaged by the spreading of specimens for thick films. Thick films are prepared by spreading a drop of blood on a slide to cover a 15–20-mm diameter area and staining with Giemsa staining technique or Field’s rapid technique (see p. 62) as for malaria smears. Microscopically, T. b. gambiense and T. b. rhodesiense cannot be distinguished from each other; they are 13–42 mm long with a single flagellum, a centrally placed nucleus, and a small dot-like kinetoplast. T. cruzi measures 12–30 mm and has a larger kinetoplast than T. b. gambiense and T. b. rhodesiense (Fig. 6.12B,C).

Concentration Techniques

Quantitative buffy coat method

The QBC method⁷¹ is referred to on p. 110. After centrifugation, the tube should be left to stand upright for 5 min, and the plasma interface area is then examined for motile trypomastigotes. This has been suggested as the ‘gold standard’ for diagnosis.

Capillary tube method

Fill one or two micro-haematocrit capillary tubes with EDTA or citrated blood. Seal the ends and centrifuge for about 5 min as for microhaematocrit. Then lay the capillary tubes adjacent to each other on a microscope slide, and secure both ends onto the slide with adhesive tape (Fig. 6.13). Examine the plasma just below the red cell and buffy layer immediately for motile trypomastigotes using a ×20 or ×10 objective with the condenser iris partially closed or by dark-field microscopy.

Figure 6.13 Capillary tube concentration method. Method used for detecting trypomastigotes or microfilariae in blood.

Filariasis and loiasis

Filariasis involving the lymphatics is the cause of elephantiasis. It is caused by the filarial worms Brugia malayi, Wuchereria bancrofti and Brugia timori, whereas filarial infection of the subcutaneous tissues is caused by Loa loa. The larvae of these worms, microfilariae, are transmitted by mosquito to humans, where they can be found in the blood and where they show periodicity with fluctuating levels at different times of the day (Fig. 6.12D).

Babesiosis

Babesiosis results from a tick-borne intraerythrocytic protozoan, Babesia. Humans are infected by chance in the natural cycle of transmission between the tick and its domestic or wild animal host. It is especially prevalent in subtropical and tropical countries. The infection results in high fever accompanied by jaundice and severe haemolytic anaemia with haemoglobinuria; there is a leucocytosis with neutrophilia.

The parasites can be seen in the erythrocytes in Giemsa-stained blood films. Morphologically they are variable round or oval bodies that may be mistaken for the ring form of plasmodium. However, in babesiosis the dividing cells characteristically consist of two daughter cells held together by a thin strand of cytoplasm; also, no pigment occurs in erythrocytes infected with older stages of Babesia.

Ehrlichiosis

Ehrlichiosis is a tick-borne fever in which clusters of small organisms may be seen in Giemsa-stained blood smears. The detection of organisms within neutrophils or monocytes is important for its diagnosis.