I Clin Pathol 1996;49:533-538

ACP Broadsheet No 148

Laboratory diagnosis of malaria

D C Warhurst, J E Williams

July 1996

IntroductionPersons with a history of travel in parts of theworld where malaria is endemic who developfever within weeks of return, may be sufferingfrom malignant tertian malaria (Plasmodiumfalciparum). P falciparum infections rarelypersist for more than one year untreated, butmay be fatal within days. In 1994 and 1995there were 11 and four, respectively, importedmalaria fatalities in the UK.The other three species of malaria infecting

humans may, after the initial feverish symp-toms have died down, recur after severalmonths to four years in the case ofP vivax andP ovale, which have dormant liver forms, andfor up to 40 or more years in the case of Pmalariae, which can persist in the blood ofuntreated persons.'Although a history of overseas travel is

usually a factor when malaria presents, itshould be remembered that malarial infectionsmay be acquired by blood transfusion, sharingsyringes, organ transplants, and accidentallaboratory inoculation. A significant number ofreports exist of malaria transmission havingtaken place in the vicinity of airports innon-endemic countries, owing to the acciden-tal importation of infected vector femaleAnopheles mosquitoes.

Currently, the accepted diagnostic techniquefor malaria is the examination of stained bloodfilms under the oil immersion lens of themicroscope. Serology plays a part in epidemi-ology and in various special investigations.2

Samples for blood film diagnosisFor malaria diagnosis blood should ideally betaken direct from the patient's finger or ear andthe smears prepared at the bedside or in theclinic. Films adhere better to the slides, leave aclearer background after lysis, and parasite andred cell changes are minimal.

If it is necessary to use anticoagulants, thenthe films should be made as soon as possible,certainly less than three hours, after the bloodwas drawn. EDTA is superior to otheranticoagulants for this purpose. Parasite andred blood cell morphology can be seriouslyaffected if the blood has been in anticoagulantfor too long.3

Further development of the sexual stagesmay occur (even within 20 minutes under theright conditions) and the male gametes re-

leased into the plasma may be mistaken forother organisms, such as Borrelia. They may bedistinguished from Borrelia by the central loca-tion of the red-staining nucleus, and theabsence of notable, repetitive sinuous curves.

If parasitised blood is left at warm laboratorytemperature, red cells may be invaded byreleased merozoites. This may lead-for exam-ple, to the occurrence of appreciable numbersof "accole" forms, characteristic of P falci-parum, in blood parasitised by P vivax.

Heavier parasitaemias left for several hoursmay lead to the serious deterioration of thealready delicate parasitised erythrocytes, owingto a build-up of acid in the blood sample.Those parasitised with early P vivax forms mayshrink or become crenated. Later stages of theparasite may become compact, and the eryth-rocyte membrane may become very delicate sothat it stretches when the film is prepared. Pro-longed exposure, for more than 10-12 hours,may result in the total destruction of the laterstages of P vivax, making recognition difficultor, in some instances, impossible.Blood should, if possible, be taken during or

after pyrexia, and before the administration ofantimalarial drugs. Drug treatment, besidesmaking parasitaemia harder to detect, causesconfusing morphological changes in the para-sites. Chloroquine-for example, causesclumping of pigment vesicles and can lead toother species being mistaken for Pfalciparum.Where a bone marrow biopsy specimen is

available, this may be a fruitful specimen formalaria diagnosis. Buffy coat smears may beuseful for the detection of scanty maturingstages and gametocytes, but are not recom-mended routinely.For routine diagnosis of malaria, four thin

and four thick blood films should be prepared.One of the thin films, stained with Giemsa,should be adequate for detection of normalparasitaemias and for determination of species.To save time, where essential, the rapid Field'stechnique may be used, but should be backedup by Giemsa stained thin films. The thickfilms should be stained using Giemsa or Field'sstain. Ifproblems with diagnosis arise the otherfilms are available for further study. Inaddition, the PHLS Malaria Reference Labo-ratory would appreciate having unstainedfilms, both thick and thin, for confirmation.(Malaria is a notifiable disease in the UK.) If

This Broadsheet has beenprepared by the authors atthe invitation of theAssociation of ClinicalPathologists who reserve thecopyright. Further copies ofthis Broadsheet may beobtainedfrom thePublishing Manager,J'ournal of ClinicalPathology, BMA House,Tavistock Square, LondonWClH 9JR.

PHLS MalariaReference Laboratory,London School ofHygiene and TropicalMedicine

Correspondence to:Dr D C Warhurst, Reader inMedical Protozoology,London School of Hygieneand Tropical Medicine,Keppel Street,LondonWC IE 7HT.

Accepted for publication17 March 1996

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clinical evidence of malaria is strong, yet para-

sites are not found in the films taken initially,then further films should be taken at six hourlyintervals.

Stained thin blood filmPRINCIPLETwo methods are described. A rapid method,where an urgent diagnosis is required, and a

longer method which provides the best stainingfor the successful identification of species.

Preparation of the smear and the staining issimilar to that used for normal haematology,except that Giemsa stain is used, and dilutionis made in alkaline buffer (pH 7.2) instead ofthe usual slightly acidic buffer used by mosthaematology laboratories.

EQUIPMENTStaining tray, constructed so that the slidesmay be stained face downwards in a small vol-ume of solution; 20 ml disposable syringe and5 cm x 19 g blunt needle; Coplin jars.

STAINS AND REAGENTS* Giemsa stain solution (BDH, R66; Product35086). This has variable batch quality andeach new batch should be checked against a

known P vivax before routine use. Field's stainsolution A (BDH; Product 35056) and Field'sstain solution B (BDH; Product 35057).* As an alternative to commercial Field's stain,the solutions may be made up as follows: (1)methylene blue (medicinal), 0.8 g; Azure I, 0.5g; Na2HPO4, 5.0 g; KH2PO4, 6.25 g; distilledwater, 500 ml. (2) Eosin, 1.0 g; Na2HP04, 5.0g; KH2PO4, 6.25 g; distilled water, 500 ml.The solutions should be filtered after beingallowed to stand overnight.* Phosphate buffered distilled water (pH 7.2):KH2PO4, 0.7 g; Na2HPO4, 1.0 g; distilledwater, 1.0 litre.* Absolute methanol (Analar).STAINING METHOD

Giemsa

1 Prepare thin blood film as for routinehaematology. Ensure that the film has a good"tail" and does not reach the edges of the slidelaterally.2 Allow the film to dry in air and fix withmethanol for 30 to 60 seconds.3 Tip off excess methanol and place face downon a staining tray.4 Using the 20 ml syringe and blunt needle,dilute the stock Giemsa 1 in 10 with buffereddistilled water. Mix well and expel air.5 Infiltrate the stain, using the syringe andneedle, under the slide, taking care not to traplarge air bubbles. Stain for 40-45 minutes.6 At the end of the staining time, rinse theslides briefly with tap water and allow to draindry in a vertical position. A x5O or x63 oilimmersion objective is invaluable for prelimi-nary examination of blood films.The syringe method for dilution of Giemsa isstrongly recommended, as once the stain isdiluted with water, precipitation of the stainbegins, which is hastened by exposure to air.Staining face-downwards also reduces precipi-tation, and any that does develop falls away

from the smear. Cleanly stained smears arevery important when searching for small,intracellular parasites.The buffered water at pH 7.2 must be used

for the dilution of stain for blood parasites. It isonly at this alkaline pH that proper differentia-tion of parasite nuclear and cytoplasmic mate-rial takes place, as well as the staining of cyto-plasmic and membrane changes in infected redblood cells (for example, Schiuffner's dots andMaurer's clefts). It must be stressed that acidicstaining is not suitable for proper diagnosis ofblood parasites on thin films.

Rapid Field's methodfor thin films1 Prepare and fix film as for the Giemsa tech-nique.2 Flood slide with 1 ml Field's stain B (redsolution) diluted 1 in 4 in buffered water (pH7.2).3 Immediately add an equal volume of Field'sstain A (blue solution), mix well and stain forone minute.4 Rinse in tap water and drain dry.

Thick blood filmThick blood films allow a rapid examination ofa relatively large volume of blood, enabling thedetection of even scanty parasitaemias of allblood parasites. A well prepared thick bloodfilm gives more than a 10-fold increase in sen-sitivity over thin films.As with thin films, two techniques are given:

a rapid Field's technique (giving excellent col-oration, but more dependent on the skill of theoperator for optimum results) and a slowerGiemsa stain method which is more consistentin its staining quality.

PRINCIPLEAn unfixed dried film, about five to six redblood cells thick, is made and the haemoglobinlysed out either before or during the stainingprocess. The malaria parasites in the film arestained with little interference from the largenumbers of red blood cells present, and can beseen against a relatively clear background.

PREPARATION OF THICK BLOOD FILM1 A drop of blood, 3-5 mm in diameter (3-5,ul) is put into the centre of a 76 x 26 mm slideand spread, with the corner of another slide ora swab stick, to cover an oval area ofapproximately 10-15 mm diameter.2 The final density of the smear should allownewsprint to be just visible through it.3 Thoroughly dry the smear, horizontally, inan incubator at 37°C for one hour.STAINING METHODGiemsa1 Do not fix the dry film, but place it gently ina Coplin jar containing buffered water (pH7.2) and allow to lyse until no more haemo-globin can be seen falling away from the smear(usually three to five minutes).2 Remove from water, place face-down onstaining dish, and stain with Giemsa diluted 1in 10 with water, as for the thin film method.3 Stain for 30 minutes; then rinse briefly withtap water and drain dry. Examine film asdescribed for a thin blood film.

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Field's stain1 Do not fix the dry film.2 Dip film for three seconds in Field's solutionA.3 Wash gently in a jar of tap water and quicklydrain off excess water onto absorbent paper.4 Dip film for three seconds into Field's solu-tion B.5 Wash gently in a jar of tap water, and allowto drain dry in a vertical position.6 When dry, examine as for a thin blood film,paying particular attention to the lower half ofthe film, where haemoglobin from the lysismakes the best background colour contrast fordetection of the parasites.Films must be absolutely dry before lysis, orelse the blood smear is likely to detach from theslide. The films should be washed, in steps 3and 5, until no more colour comes away fromthe film.

Blood films from suspected haemor-rhagic fevers or HIVThe recommended technique is as follows:

THICK FILMS1 Fix the dried smear directly in 10% bufferedformalin for 10 minutes.2 Wash three times (total three minutes) inbuffered water and stain with Giemsa as usual.THIN FILMS1 Fix in methanol for five minutes.2 Fix in formalin as above, wash and stain withGiemsa.CommentNuclei of malaria parasites stain darker by thistechnique, and the characteristic red blood cellstippling is not always satisfactorily stained.

Examination of filmsThe theoretical increase in efficiency of exam-ining a thick film rather than a thin one is by afactor of 44 because of the larger volume ofblood examined. However, when the param-eters of thin and thick blood films were studiedby Dowling and Shute4 they found thatcompared with thin films, 60% of parasiteswere missing or obscured on thick films, andthis rose to 80% for very low parasitaemias. ForP falciparum gametocytes this value rose to86% or more. In practice, for ring stages, thickfilms are 11 times more sensitive than thin.Because of superior preservation of morphol-ogy, recognition of parasite species on the thinfilm is more reliable.

In 100 high power (xlO0) fields of the thick-est part of a thick film, which takes an expertabout three minutes to examine, approximately0.28 ,ul blood is present and, if one parasite isseen, there are estimated to be four to fiveparasites/4l. In a 30 minute examination of athick film, the microscopist is estimated to havea 2/3 chance of detecting one parasite/,ul.Where detection of parasite presence is crucial,reexamination of the film by another, equallyskilled operator can increase sensitivity.5

In a diagnostic laboratory, where blood filmsare regularly examined for malaria and otherparasites, it has been found effective to ensurethat each thick film is examined consecutively

by two persons, each of whom will scan 100high power fields. In special circumstanceswhere the parasite count is thought to be verylow and approaching one parasite/l, themicroscopists may examine 500 high powerfields each. In the experience of the authorsmore extensive examinations are rarely justi-fied, even in the Reference Laboratory.The thin film is extremely valuable for the

specific determination of the malaria parasite,for counting the higher parasitaemias and alsoas a fail-safe for the thick film procedure. Toenhance the possibility of parasite detection onthe thin film, particularly for P ovale and Pvivax, examine the edges and tail of the film.

In view of the intense visual and mental con-centration involved in the examination ofbloodfilms, it is important that the operatives shouldbe comfortably seated with the arms suitablysupported. The microscope should have a bin-ocular head, and be correctly adjusted for eacheye. Not more than 20 films should beexamined without a break of 30 minutes doinganother, non-microscope activity. In the au-thors' view 50 thick films, of which the major-ity are scanty or uninfected, is probably toomany for one operative to examine on a dailybasis while retaining sanity and the ability todiagnose. A maximum of around 35 should beaimed at.

Counting the parasitaemiaA recognised way of estimating the parasitecount per jl on thick films is to use a standardvalue (8000/4d) for the white cell count as amultiplier. Thus, parasites in thick film fieldsare counted until 200 leucocytes have beenseen, the parasite count is then multiplied by40 to give parasites per pl of blood. This takesno account of the known loss of parasites in thethick film, which suggests it is an underesti-mate, but the "standard" white blood cell valueis probably too high, which helps to cancel outthis error. The method is widely used forestimation of the intensity of the infection. Toconvert approximately to percentage parasitae-mia, the number of parasites per 200 whitecells is divided by 1250.

Similarly, the parasitised cell percentage canbe estimated from thin films. Generally, thenumber ofinfected cells per 100 red blood cellsis counted. The infected cells are counted asone, whether they contain one, or several para-sites. Sufficient red blood cells are examined toobtain an estimate of the mean number perhigh power field in an evenly spread area of thefilm. Then, parasitised cells in 10 fields arecounted to give a mean parasitised cellpercentage.The use of an eyepiece graticule consisting

of a square covering most of the field, with asmaller square 1/9 the area of the large squarescribed in the corner will facilitate counting thenumber of erythrocytes per parasite, as follows:* Choose an area of the film showing evenlydistributed erythrocytes, as above.* Count the erythrocytes in the small squareand the parasitised cells in the large square.

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Table 1 Differential diagnosis ofPlasmodium species ofman in Giemsa stained thin films ofperipheral blood

P falciparum P vivax P malariae P ovale

TrophozoitesRing forms 0.15-0.5 of diameter of RBC; 0.3-0.5 of diameter of RBC which 0.3-0.5 of diameter of RBC which 0.3 of diameter of RBC which are

RBC normal size. Cytoplasm very are unaltered in size. Cytoplasm are unaltered in size. Cytoplasm unaltered in size. Cytoplasmfine with young rings; thick circle, thin. circle, thicker. circle, thicker.irregular in old rings. Marginal"accol6' forms, forms with twochromatin dots and multipleinfections common.

Growing forms RBC unaltered in size, sometimes RBC enlarged, stippled. Parasite RBC unaltered in size. Parasite RBC unaltered in size or slightlyspotted, pale. Parasite compact; amoeboid, vacuolated; pigment compact, rounded or band enlarged; stippled; may be ovalpigment dense brown or black fine and scattered golden brown. shaped; dark brown or black and fimbriated. Parasite compact,mass. pigments, often concentrates in a rounded; pigment fine brown

long line along one edge of the grains.band.

Mature schizonts RBC unaltered in size, sometimes RBC much enlarged, stippled. RBC unaltered in size. Parasite RBC frequently oval, fimbriated,spotted, pale. Parasites about 0.6 Parasites large, filling enlarged fills RBC completely; nuclei or enlarged and stippled. Parasite asof diameter of RBC; nuclei or RBC; nuclei or merozoites 12-24, merozoites 6-12, usually 8, for P malariae but does not fill themerozoites 8-24; pigment usually 16; pigment a golden sometimes forming a rosette; RBC entirely; pigment brownclumped, black. Not usually seen in brown central loose mass. pigment brown black central central clump.peripheral blood, clump.

Gametocytes RBC distorted. Parasite RBC enlarged, stippled. Parasite RBC unaltered in size. Parasite RBC slightly enlarged, stippled.crescentic. large, rounded, filling enlarged small, round, filling RBC. Parasite round, not usually filling

RBC. cell.Stippling Maurer's clefts Schiiffner's dots None. (Fine dots after prolonged James's dots

staining may be seen.)

RBC = red blood cells.

* Repeat the count on subsequent fields until50 parasitised cells have been counted or 200erythrocytes have been counted in the smallsquare.* Multiply the total erythrocytes in the smallsquares counted by 9. This is the denominatorto use when calculating the percentage parasit-ised erythrocytes.

CommentParasitaemia estimation is highly valuable forthe clinician, as it is an important determinantof treatment schedules for P falciparum. Forexample, if parasitaemia exceeds 10% in Pfal-ciparum, exchange transfusion may be indi-cated. If late trophozoites or early dividingforms of this parasite (usually containingnoticeable pigment deposits) are seen in theperipheral blood, this should be reported, as itmay indicate that the patient is in a criticalcondition.6The presence of gametocytes of Pfalciparum

in the absence of growing stages is clinicallysignificant in an untreated symptomatic patientas it may indicate the presence of a suppressedactive infection. Gametocytes seen in the bloodafter treatment, however, are ofno significance.

Interpretation of thin and thick filmsThe presence of intra-erythrocytic bodies, gen-erally consisting of a blue staining cytoplasmicarea closely associated with a small reddishstaining nuclear area and, in the larger, moremature parasites, the presence within theorganism of yellow-brown to black malariapigment, is diagnostic of malaria infection.During the intra-erythrocytic growth of themalaria parasite, it finally divides to give amaximum of 24 infective merozoites. The hostcell may show enlargement (P vivax and Povale), remain the same size, or shrink (some-times in Pfalciparum and often in P malariae).The erythrocyte membrane may develop sur-face markings (Schiiffner's and James's dots)which stain pink with Giemsa at pH 7.2 (Pvivax and P ovale). All stages of the parasite

may be seen in the peripheral blood in the caseof P vivax, ovale and malariae, but generallyonly the small ring parasites and (in olderinfections) the banana-like gametocytes arefound in P falciparum. In infections of Pfalciparum, a few intra-erythrocytic spots ap-pear, particularly noticeable in erythrocytesinhabited by the thicker ring forms. These aretermed Maurer's clefts, and should bedistinguished carefully from the finer, muchmore numerous Schfiffner's and James's dotsfound in P vivax and P ovale (table 1 and fig 1)

SerologySerology is not used routinely for diagnosticpurposes as blood films are quicker and detectactive infection. However, serology is valuablefor:* Transfusion blood screening, and the investi-gation of transfusion acquired infections. Se-rology is the only satisfactory techniquecurrently available for adequately screeningtransfusion blood, as an antigen or nucleic acidmethod would have to detect one malariaparasite in a unit of blood. A donor who visitedan endemic area more than six monthspreviously and has not developed antibody isunlikely to be carrying a malaria infection. Inthe unlikely case of an antibody negative donorbecoming infective from newly developing dor-mant hypnozoites in the liver, the organism isexpected to be P vivax or ovale, not the poten-tially fatal Pfalciparum.* Retrospective diagnosis.* Investigation of "cryptic" malaria.* In endemic areas, serology is invaluable forepidemiological purposes.For serological studies, blood stage antigen

is used, prepared from primate blood infec-tions or from Pfalciparum cultures in the labo-ratory. Falciparum antigen tends to cross reactwith antibodies directed against the other spe-cies, although generally at a lower titre, so thatit can be used as a general screen. The schizontstage of the blood cycle (obtained fromcultures in the case of Pfalciparum) is used for

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preparation of antigen as this gives a more sen-sitive test. The blood or culture should bewashed well with phosphate buffered salinebefore use to remove serum or plasma proteins.

For the indirect fluorescent antibody test(IFAT), antigen thick films are made on slides,dried, wrapped in tissue, and stored frozenunder desiccation. This antigen is very stableand will last for years provided thawing is car-ried out in a desiccator.For ELISA antigen, the infected erythro-

cytes are lysed with saponin in the presence ofprotease inhibitors, the erythrocyte material is

A

C

E

removed by centrifugation, nonidet P40 ex-tracts are prepared, and stored frozen. Antigenprepared for ELISA is relatively ephemeral andneeds to be used within six months, otherwisefalse positivity is observed. Commercial kitsare available.

Other methods of malaria parasitedetectionQUANTITATIVE BUFFY COATThe first of the practicable commercial kits fordiagnosis (quantitative buffy coat, QBC) wasbased on technology which has been available

B

D44

F

*

G H

Figure 1 Plasmodium falciparum: (A) small ring stage trophozoites and (B) late trophozoite and "accole"form,showing Maurer's clefts. Plasmodium vivax: (C) ring stage amoeboid trophozoites in enlarged cells showing developingSchiiffner's dots and (D) mature schizontfilling a heavily stippled, enlarged erythrocyte. Plasmodium ovale: (E)trophozoite showing an elongated, stippled andfimbriated erythrocyte and (F) almost mature schizont showing eight( ?)nuclei, not filling an elongated, stippled andfimbriated erythrocyte. Plasmodium malariae: (G) "bird's eye" ring stage and(H) mature schizont with five(?) merozoites filling an unenlarged, unstippled erythrocyte.

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for decades, the differential centrifugation ofblood and the acridine orange staining of themalaria parasite.7 Blood samples in acridineorange coated heparinised tubes are centri-fuged and the area just below the buffy coat isexamined in situ under a fluorescence micro-scope to detect the parasitised cells.

In studies in UK laboratory8 and in fieldconditions9 the sensitivity has been found to beabout the same as the Field's stained thick film.It is still necessary to use films for specificdeterminations, and it is recognised that thesensitivity for detection of species other than Pfalciparum is appreciably less. Howell-Jollybodies (nucleic acid remnants in red bloodcells in megaloblastic anaemia or other dys-haemopoietic conditions) may possibly givefalse positive results as they do in other fluores-cence staining methods.'0 If late parasite stagesor gametocytes are present they tend to localiseamong the highly fluorescing leucocytes of thebuffy coat and are easily missed. The QBCtechnique gives only a rough indication of theintensity of the infection, so for properdiagnostic work thick and thin films need to berun in parallel. The tubes do not remain read-able at their most sensitive for much more thana matter of hours, so for any permanent record,slides are necessary. It should also be remem-bered that centrifuging capillary tubes ofbloodposes a safety hazard.

It is easier to train workers to use the QBCthan to interpret thick films. The technique isquicker than preparation, drying, staining, andexamination of thick films. In some locations inthe UK the technique has been introduced foruse on call, because of its greater simplicity.The cost per capillary tube is £2.50. This andthe cost of the additional apparatus neededprobably rules out use in small rural clinics inthe developing world.

ANTIGEN DETECTIONA monoclonal antibody of high affinity againstP falciparum histidine-rich protein 2 (HRP2:AHH[AHHAAD]2), found in the membraneof infected erythrocytes and in the plasma ofinfected persons, has been used in two testsspecific for Pfalciparum infection. One of thesetests, a dipstick method ("ParaSight-F"),"' iscommercially available, costing £4.00 per test.The technique depends on the capture of theHRP2 by a line of monoclonal antibodyattached to a dipstick composed of nitrocellu-lose and fibre. A control dashed line ofHRP2 isalso present on the dipstick to monitor thedetection reagent. The bound HRP2 is de-tected by a dye labelled anti-HRP2 polyclonalantibody. This technique promises to be ofgreat value for the routine haematology labora-tory; in parasitology departments it will be par-ticularly useful for the detection ofPfalciparumin mixed infections. In field studies the dipstickhad >90% sensitivity when tested on bloodcontaining >60 parasites/,ul; positivity maysometimes persist for several days after treat-ment, and has rarely been reported in unex-posed persons.'2 More importantly, false nega-tive results have been reported in a small

number of patients even when parasitaemiaabove 10004d1 was present. It may be that highantibody level in plasma in some samples fromendemic areas is rendering the antigen inacces-sible, but this observation remains a worry. Itcannot be assumed that a negative ParaSight Ftest rules out malaria.

NUCLEIC ACID METHODSWe have selected three reports from a rapidlygrowing literature on nucleic acid detection ofmalaria.The probe method,'3 using alkaline phos-

phatase for the detection system, seems to bethe best of the non-radioactive probes reportedso far. This is applicable to carrying out largescale epidemiological surveys for P falciparumonly, and may be useful, though insensitivecompared with serology, for transfusionscreening. It is probably not practicable forclinical investigations.Two PCR'4 techniques are mentioned here,

the first of which is reported to be verysensitive indeed, and is applicable to largeruns."' It might be used epidemiologically for Pfalciparum and, although still too insensitive,for transfusion screening. The second is acolorimetric PCR'6 for a small subunit rRNAgene which detects and identifies all fourspecies of the parasite. It is particularly suitablefor reference studies or microepidemiology.

1 Gilles HM, Warrell DA (eds). Bruce Chwatt's EssentialMalariology. 3rd edn. London: Edward Arnold, 1993.

2 Draper CC, McLaren ML. Some applications of immu-noassays in tropical diseases. In: Voller A, Bartlett A. Bid-well D, eds. Immunoassays for the 80's. Lancaster: MTPPress, 1981:449-56.

3 Ree GH, Sargeaunt PG. Laboratory diagnosis of malaria.BMJ 1976; 1: 152.

4 Dowling MAC, Shute GT. A comparative study of thick andthin blood films in the diagnosis of scanty malaria parasi-taemia. Bull World Health Organ 1966;34:249-67.

5 Raghavan K. Statistical considerations in the microscopicaldiagnosis of malaria, with special reference to the role ofcross checking. Bull World Health Organ 1966;34:788-91.

6 Field JW, Shute PG. The microscopical diagnosis of humanmalaria. Vol 2. Kuala Lumpar: Government Press, 1956.

7 Levine RA, Wardlaw SC, Patton CL. Detection ofhaemoparasites using quantitative buffy coat analysistubes. Parasitol Today 1990;5:132-4.

8 Moody AH, Hunt-Cooke A, Chiodini PL. Experience withthe Becton Dickinson QBC II centrifugal haematologyanalyser for haemoparasites. Trans R Soc Trop Med Hyg1990;84:782.

9 Bawden M, Malone J, Slaten D. QBC malaria diagnosis:easily learned and effectively applied in a temporary mili-tary field laboratory. Trans R Soc Trop Med Hyg1994;88:302.

10 Facer CA. Rapid malaria diagnosis: an update. In: Rapidmethods and automation in microbiology and immunology.Andover: Intercept, 1994:14-143.

11 Shiff CJ, Minjas J, Premji Z. The rapid manual ParaSight-Ftest. A new diagnostic tool for Plasmodium falciparuminfection. Trans R Soc Trop Med Hyg 1993;87:646-8.

12 Beadle C, Long GW, Weiss WR, McElroy PD, Maret SM,Aggrey JO, et al. Diagnosis of malaria by detection of Plas-modium falciparum HRP2 antigen with a rapid dipstickantigen capture assay. Lancet 1994;343:564-8.

13 McLaughlin GL, Subramanian S, Lepers JP, RaharimalalaL, Deloron P. Evaluation of a nonisotopic DNA assay kitfor diagnosing Plasmodium falciparum malaria in Mada-gascar. AmJ Trop Med Hyg 1993;48:211-15.

14 Wilson SM. Application of nucleic acid based technologiesto the diagnosis and detection of disease. Trans R Soc TropMed Hyg 1993;87:609-11._

15 Barker RH, Banchongaksora T, Courval JM, Suwonkerd W,Rimwuntragoon K, Wirth DF. A simple method to detectPlasmodium falciparum directly from blood samples usingthe polymerase chain reaction. Am Y Trop Med Hyg1992;46:416-26.

16 Oliveira DA, Holloway BP, Durigon EL, Collins WE, LalAA. Polymerase chain reaction and a liquid phase noniso-topic hybridization for species specific and sensitive detec-tion of malaria infection. Am J Trop Med Hyg 1995;52:139-44.

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