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BioMed Central
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Malaria Journal
Open Access
Research
Comparison of PCR and microscopy for the detection of
asymptomatic malaria in a Plasmodium falciparum/vivax endemic
area in Thailand
Russell E Coleman*
1
, Jetsumon Sattabongkot
1
, Sommai Promstaporm
1
,
Nongnuj Maneechai
1
, Bousaraporn Tippayachai
1
, Ampornpan Kengluecha
1
,
Nattawan Rachapaew
1
, Gabriela Zollner
1
, Robert Scott Miller
1
,
Jefferson A Vaughan
2
, Krongtong Thimasarn
3
and Benjawan Khuntirat
1
Address:
1
Departments of Entomology and Immunology, U.S. Army Medical Component, Armed Forces Research Institute of Medical Sciences,
Bangkok, Thailand,
2
University of North Dakota, Grand Forks, North Dakota, USA and
3
Ministry of Public Health, Nonthaburi, Thailand
Email: Russell E Coleman* - russell.coleman@na.amedd.army.mil; Jetsumon Sattabongkot - jetsumonp@afrims.org;
Sommai Promstaporm - sommaip@afrims.org; Nongnuj Maneechai - nongnujm@afrims.org;
Bousaraporn Tippayachai - bousarapornt@afrims.org; Ampornpan Kengluecha - ampornpank@afrims.org;
Nattawan Rachapaew - nattawanr@afrims.org; Gabriela Zollner - gabriela.zollner@na.amedd.army.mil;
Robert Scott Miller - robert.s.miller@us.army.mil; Jefferson A Vaughan - jefferson_vaughan@und.nodak.edu;
Krongtong Thimasarn - thimasarnk@searo.who.int; Benjawan Khuntirat - benjawank@afrims.org
* Corresponding author
Abstract
Objective: The main objective of this study was to compare the performance of nested PCR with
expert microscopy as a means of detecting Plasmodium parasites during active malaria surveillance
in western Thailand.
Methods: The study was performed from May 2000 to April 2002 in the village of Kong Mong Tha,
located in western Thailand. Plasmodium vivax (PV) and Plasmodium falciparum (PF) are the
predominant parasite species in this village, followed by Plasmodium malariae (PM) and Plasmodium
ovale (PO). Each month, fingerprick blood samples were taken from each participating individual
and used to prepare thick and thin blood films and for PCR analysis.
Results: PCR was sensitive (96%) and specific (98%) for malaria at parasite densities ≥ 500/µl;
however, only 18% (47/269) of P. falciparum- and 5% (20/390) of P. vivax-positive films had parasite
densities this high. Performance of PCR decreased markedly at parasite densities <500/µl, with
sensitivity of only 20% for P. falciparum and 24% for P. vivax at densities <100 parasites/µl.
Conclusion: Although PCR performance appeared poor when compared to microscopy, data
indicated that the discrepancy between the two methods resulted from poor performance of
microscopy at low parasite densities rather than poor performance of PCR. These data are not
unusual when the diagnostic method being evaluated is more sensitive than the reference method.
PCR appears to be a useful method for detecting Plasmodium parasites during active malaria
surveillance in Thailand.
Published: 14 December 2006
Malaria Journal 2006, 5:121 doi:10.1186/1475-2875-5-121
Received: 25 September 2006
Accepted: 14 December 2006
This article is available from: http://www.malariajournal.com/content/5/1/121
© 2006 Coleman et al; licensee BioMed Central Ltd.
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0
),
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Malaria Journal 2006, 5:121 http://www.malariajournal.com/content/5/1/121
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Background
The detection of asexual parasites by light microscopy of
Giemsa-stained thick and thin films remains the standard
laboratory method for the diagnosis of malaria [1,2].
Although detection of parasites in symptomatic patients
reporting to local malaria clinics is the primary means
used for malaria diagnosis in Thailand, use of active
(cross-sectional) surveillance provides a tool for detection
of patients with asymptomatic malaria and relatively low
parasite rates. In Thailand, active surveillance is used in
remote areas where individuals may have difficulty in
reaching a malaria clinic – in this situation malaria clinic
personnel make periodic visits to a given village and
examine blood smears from all individuals present in the
village. In Thailand and many other malaria endemic
regions, there are problems and limitations associated
with reliance on microscopic diagnosis of malaria for
both active and passive case detection [3-5]. These include
lack of skilled microscopists, variation in individual train-
ing and/or experience, limited supply of microscopes and
reagents as well as variation in equipment maintenance,
and inadequate quality control [6]. When symptomatic
patients with a relatively high parasitaemia (e.g., >500
asexual parasites/µl) report to a malaria clinic for treat-
ment, microscopy can provide an accurate diagnosis that
is used to initiate appropriate chemotherapy. However,
accuracy of microscopy can decrease significantly at lower
parasitaemia levels [4]. Parasitaemia rates in asympto-
matic patients are often quite low – in a recent study in
western Thailand, the median asexual parasitaemia rate in
82 Plasmodium falciparum and 98 P. vivax-positive individ-
uals was 848 and 155 asexual parasites/µl blood, respec-
tively [7].
Polymerase chain reaction (PCR)-based methods have
been used since the early 1990's for the detection of Plas-
modium parasites in human patients. Several experimental
assays using various primers and extraction and detection
techniques have been reported [8-12]. In general, PCR
was more sensitive and specific than examination of thick
or thin blood smears, particularly in cases with low para-
site rates or mixed infections [2,8,10,13-16]. PCR was also
more sensitive than the QBC method [2] and various dip-
stick assays [17].
Although the majority of studies have shown that PCR is
both sensitive and specific for the diagnosis of malaria,
there are limitations that can affect the accuracy of the
method. Selection of appropriate primers, methods used
for collection and storage of blood samples, and extrac-
tion methods can all affect PCR performance. Jelinek et al.
[18] reported that the sensitivity of PCR was as much
linked to parasite density as was microscopy. They found
that sensitivity of PCR was affected by both parasite den-
sity and by geographic differences in parasite populations.
Other studies have reported that PCR may occasionally
yield false negative results [9,19,20]. Barker et al. [20]
carefully analysed discrepancies between microscopy and
PCR, and although true false negative PCR results did
occur, the majority of discrepancies resulted from prob-
lems with microscopy. Most recently, Scopel et al. [21]
determined that use of DNA extracted from thick blood
smears resulted in poor detection of malaria parasites,
particularly with parasite densities less than 20/µl.
In this study the performance of nested PCR for the detec-
tion and identification of Plasmodium parasites was evalu-
ated in a malaria endemic village in western Thailand,
with expert laboratory microscopy used as the reference
standard. The goal of the study was to evaluate perform-
ance of PCR at low parasite densities in a population that
was primarily asymptomatic.
Methods
Study site
The study was performed from May 2000 to April 2002 in
the village of Kong Mong Tha (98°33'16" E and
15°10'17" N), located in Laivo Sub-District, Sangkhlaburi
District, Kanchanaburi Province, western Thailand. Kong
Mong Tha is an isolated village accessible only by boat or
foot for 10 months of the year. Plasmodium vivax (PV) and
Plasmodium falciparum (PF) are the predominant parasite
species in this village, followed by Plasmodium malariae
(PM) and Plasmodium ovale (PO). During a recent two-
year study, 92% (320/346) of parasitemic individuals in
this village were asymptomatic at the time blood was col-
lected [22]. The study was approved by the Ethics Com-
mittee of the Ministry of Public Health (MOPH),
Bangkok, Thailand, and by the Human Subjects Research
Review Board of the United States Army, Fort Detrick,
Maryland, U.S.A.
Patients and sample collection
Adults (≥18 years old) and children (≥1 to <18 years old)
living in Kong Mong Tha were enrolled in the study.
Informed consent was obtained from all adults willing to
participate in the study, with a parent/guardian giving
consent for children. Three teams of investigators went
from house to house during a 4-day period each month
that the study was conducted. Each individual was ques-
tioned about signs and symptoms of malaria, travel his-
tory, and medications taken during the prior two weeks.
Fingerprick blood samples were taken from each partici-
pating individual and used to prepare thick and thin
blood films. An additional 3–5 drops of blood were spot-
ted onto filter paper for subsequent PCR analysis. Isoc-
ode
®
Stix (Schleicher and Schuell BioScience, Inc., Keene,
New Hampshire) were used for the collection of blood
during the first 5 months of the study and 903 Whatman
Filter Paper (Schleicher and Schuell BioScience, Inc.,
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Keene, New Hampshire) thereafter. Both Isocode Stix and
Whatman Filter Paper provide an acceptable matrix for
the collection of whole-blood samples for subsequent
processing for malaria diagnosis by PCR [19,23]. After air-
drying for several hours, blood samples were stored at
room temperature in double zip-lock plastic bags for sub-
sequent DNA extraction.
Microscopy
Thick/thin films were stained with 10% Giemsa solution
and examined at ×1,000 under oil immersion by an expert
microscopist (N.M. or N.R.) with over 10 years of experi-
ence. Each microscopist correctly identified the species of
Plasmodium as well as parasite densities in 100% (50 of
50) of slides in a blinded panel of samples provided prior
to the start of the study. The microscopist was blinded to
any clinical diagnosis and PCR results during the course of
this study. The parasite density was counted per 500 leu-
kocytes and was then expressed as the number of tropho-
zoites per microliter by assuming a leukocyte count of
7,000/µl. Kain et al. [9] found that this value provided a
good estimate of leukocyte density for villagers in Western
Thailand. The initial thick film was considered negative if
no parasites were seen after 500 leukocytes were counted.
A high quality (Olympus) microscope with an incandes-
cent light source was used. Each film required approxi-
mately 20 minutes to read.
PCR amplification
DNA was extracted from the filter paper/Isocode Stix sam-
ples using the methods described by Plowe et al. [24].
Nested PCR amplification was performed as described by
Kimura et al. [25]. Both genus (P1F and P2R) and species
specific primers (FR, MR, OR, and VR) were designed from
the small subunit ribosomal RNA gene as previously
described [25]. Briefly, 200 ng of DNA (0.05 ug for 8,000
wbc in 1 µl of blood) was used as a template in the first
amplification step of the nested PCR and 1 µl of a 1:20
dilution of the first PCR product was used for the second
amplification. All reactions were performed in a 20 µl vol-
ume consisting of 1× PCR Gold buffer II (50 mM of KCl,
15 mM of Tris-HCl, pH 8.0), 1.5 mM of MgCl2, 200 µM
of each dNTP, 0.4 µM of each primer (P1F and P2R for the
first PCR and P1F and one of each reverse primer–FR, OR,
MR, and VR–for the second PCR), and 0.25 unit of Ampl-
iTag Gold (Applied Biosystems, Foster City, CA, USA). The
PCR product was resolved by electrophoresis in 2% agar-
ose gels, stained with ethidium bromide, and observed
under ultraviolet transillumination. The expected PCR
products are 140–160 bp for the first step and 110 bp for
the second one. Technicians conducting the PCR were
blinded to any clinical diagnosis or microscopy results.
Treatment of infected patients
The name, age, and house number of all individuals with
a malaria-positive blood film were provided to employees
of the local Thai Ministry of Public Health (MOPH)
malaria clinic. All malaria-positive individuals were
treated according to the malaria treatment protocols of
the Thai MOPH. Treatment follow-up was provided by
Thai MOPH malaria clinic employees and in accordance
with Thai MOPH standard procedures.
Non-concordant results
All samples with non-concordant results were re-evalu-
ated by both microscopy and PCR. The individual reading
the blood films or running the PCR was blinded to the ini-
tial test results.
Data analysis
Epi-Info version 6 [26] was used to calculate test perform-
ance and acceptability evaluation indices of PCR, with
microscopy used as the reference standard. For this analy-
sis, it was assumed that microscopy is always correct. Per-
formance indices followed those used by Tjitra et al. [27]
and were calculated for the following microscopic diag-
noses: total malaria (all species of Plasmodium), PF
malaria (to include mixed infections), and PV malaria.
Variables measured included the number of true positives
(TP), number of true negatives (TN), number of false pos-
itives (FP), and number of false negatives (FN). Sensitivity
was calculated as TP/(TP + FN), specificity was calculated
as TN/(TN + FP), the positive predictive value (PPV) was
calculated as TP/(TP + FP), and the negative predictive
value (NPV) was calculated as TN/(FN + TN). Test accu-
racy, the proportion of all tests that gave a correct result,
was defined as (TP + TN)/number of all tests. Reliability
was expressed as the J index ((TP × TN) - (FP × FN))/((TP
+ FN)(TN + FP)). Results were considered false positive if
microscopy detected PF and the PCR assay detected PV,
and visa versa.
In order to evaluate repeatability of each diagnostic
method, the performance of each assay were evaluated
using data from samples with non-concordant results.
Repeatability of each assay was calculated by comparing
the second set of test results with the initial test results.
Sensitivity and specificity were evaluated using the initial
test results as the reference standard.
Results
A total of 672 individuals (305 females and 367 males)
participated in the study. The age range was 1 to 92 years
old (mean = 22.1 years; median = 15.3 years). Sixty per-
cent (407/672) of individuals enrolled in the study had a
least one microscopy-positive blood film during the 2
years that the study was conducted. Seventy-one percent
(291/407) had 1 or 2 positive films, 21% (86/407) had 3
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or 4 positive films, and 7% (30/407) had 5 or more posi-
tive films.
Of the 8,590 blood films collected over the course of the
study, 7.7% (665) were found to have malaria parasites by
microscopy. Of the Plasmodium-positive slides, 39.8%
(265) were PF, 58.0% (386) were PV, 1.5% (10) were PM,
and 0.6% (4) were mixed PF/PV infections (Table 1). Of
the PV-positive blood films, 79.8% (308/386) had asex-
ual parasites alone, 19.4% (75/386) had both asexual par-
asites and gametocytes, and 0.8% (3/386) had
gametocytes alone. For PF-positive films, 86.0% (228/
265) had asexual parasites alone, 13.2% (35/265) had
asexual parasites and gametocytes, and 0.8% (2/265) had
gametocytes alone, while 91% of PM-positive films had
both asexual parasites and gametocytes. The mean density
of PF parasites was 892.7/µl (SEM = 164.3), with a range
from 28–14,000/µl. For PV, the mean density was 265.5/
µl (SEM = 66.5), with a range from 14–14,000/µl, while
for PM, the mean density was 324.8/µl (SEM = 128.4),
with a range from 28-1,134/µl. Prevalence rates for both
PF and PV were highest at the start of the study and in gen-
eral decreased over the 24 months that the study was con-
ducted.
An overall comparison of PCR and expert laboratory
microscopy for active malaria surveillance is presented in
Table 1. There was 87% (242/278) agreement on the spe-
cies of parasite present in samples that were positive by
both PCR and microscopy; however, a high proportion
(58.2%, 387/665) of all microscope-positive samples
were negative by PCR and a high proportion (52.8%, 311/
589) of PCR-positive samples were negative by micros-
copy. Ninety-eight percent (381/387) of the microscope-
positive, PCR-negative samples had fewer than 250 para-
sites/µl blood.
Table 2 presents a comparison of PCR with expert micro-
scopy for the detection of PF and PV at various parasite
densities. At parasite densities of 500/µl or greater, PCR
correctly identified 96% (45/47) of microscopy-positive
PF samples and 100% (20/20) of microscopy-positive PV
samples; however, only 10% (67/659) of positive films
had parasite densities this high. Performance of PCR
dropped off markedly with decreasing parasite (both PF
and PV) densities, with sensitivity dropping to approxi-
mately 20–25% at densities of <100/µl (Table 2). Forty-
four percent (118/269) of all PF-positive blood films and
75% (294/390) of all PV-positive films had parasite den-
sities of <100/µl.
There was no significant difference (X
2
test, P < 0.05)
between Iso-code Stix and the Whatman Filter Paper for
subsequent detection of PF or PV at parasite densities of
>100/µl; however, use of the Iso-Code Stix resulted in the
detection of significantly (X
2
test, P < 0.05) more PV-pos-
itive films than did use Whatman filter paper at parasite
densities of 1–99/µl and significantly more PF and PV in
microscopy negative samples.
When using expert microscopy as the reference standard,
PCR was both sensitive and specific for the detection of
both PF and PV at parasite densities above 500/µl. How-
ever, at parasite densities below 500/µl, sensitivity of PCR
dropped off markedly (Table 3). The non-concordance
between microscopy and PCR at parasite densities below
100/µl was remarkable (Table 4). A total of 118 samples
with fewer than 100 PF parasites/µl blood were detected
microscopically. PCR failed to detect 94 (80%) of these
samples. Conversely, a total of 120 PF positive samples
were detected by PCR – microscopy failed to detect 96
(80%) of these samples. Similar results were obtained
with PV samples, with both microscopy and PCR failing
to detect 75–80% of positive samples detected by the cor-
responding method. Although quantitative PCR was not
used in this study, it can be assumed that the majority of
PCR-positive samples that were microscopy-negative con-
tained fewer than 100 parasites/µl blood.
Table 1: Comparison of PCR and expert laboratory microscopy for active malaria surveillance.
No. of samples with the following result by PCR (% of total in row)
Expert microscopy resµlt P. falciparum P. vivax P. malariae P. ovale Mixed Negative Total
P. falciparum 102 (38.4%) 13 (4.9%) 0 (0.0%) 0 (0.0%) 7 (2.7%) 143 (54.0%)
a
265
P. vivax 4 (1.0%) 131 (33.9%) 1 (0.3%) 0 (0.0%) 8 (2.1%) 242 (62.7%)
b
386
P. malariae 0 (0.0%) 0 (0.0%) 8 (80.0%) 0 (0.0%) 0 (0.0%) 2 (20.0%) 10
Mixed 3 (75.0%) 0 (0.0%) 0 (0.0%) 0 (0.0%) 1 (25.0%) 0 (0.0%) 4
Negative 70 (0.9%) 214 (2.7%) 7 (0.1%) 2 (<0.1%) 18 (0.2%) 7,614 (96.1%) 7,925
Total 179 (2.1%) 358 (4.2%) 16 (0.2%) 2 (<0.1%) 34 (0.40%)
c
8,001 (93.1%) 8,590
a
98% (140/143) of microscopy-positive P. falciparum samples that were PCR negative had <250 parasites/µl.
b
99% (239/242) of microscopy-positive P. vivax samples that were PCR negative had <250 parasites/µl.
c
Includes the following mixed infections: 28 P. falciparum/P. vivax, 3 P. vivax/P. ovale, 2 P. vivax/P. malariae and 1 P. falciparum/P. malariae.
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Table 3: Performance characteristics of PCR at different parasite densities relative to expert laboratory microscopy for active
surveillance for Plasmodium falciparum and Plasmodium vivax.
a
Trophozoites/µl
(total positive)
Sensitivity (95% CI) Specificity (95% CI) Positive Predictive
Value (95% CI)
Negative Predictive
Value (95% CI)
Accuracy J Index
P. falciparum
>5000/µl (14) 100.0% (73.2–100.0) 97.7% (97.4–98.0) 6.7% (3.9–11.3) 100.0% (99.9–100.0) 98% 0.98
>1000/µl (36) 97.2% (83.8–99.9) 98.0% (97.7–98.3) 16.8% (12.1–22.8) 100.0% (99.9–100.0) 98% 0.95
>500/µl (47) 95.7% (84.3–99.3) 98.1% (97.8–98.4) 21.6% (16.4–28.0) 100.0% (99.9–100.0) 98% 0.94
>100/µl (151) 58.3% (50.0–66.2) 98.6% (98.3–98.8) 42.3% (35.6–49.3) 99.2% (99.0–99.4) 98% 0.57
>1/µl (267) 41.6% (35.7–47.8) 98.8% (98.6–99.0) 53.6% (46.6–60.5) 98.1% (97.8–98.4) 97% 0.40
P. vivax
>5000/µl (4) 100.0% (39.6–100.0) 95.5% (95.0–95.9) 1.0% (0.3–2.8) 100.0% (99.9–100.0) 95% 0.95
>1000/µl (15) 100.0% (74.7–100.0) 95.6% (95.2–96.0) 3.8% (2.2–6.1) 100.0% (99.9–100.0) 96% 0.96
>500/µl (20) 100.0% (80.0–100.0) 95.7% (95.2–96.1) 5.1% (3.2–7.9) 100.0% (99.9–100.0) 96% 0.96
>100/µl (96) 71.9% (61.6–80.3) 96.2% (95.8–96.6) 17.6% (14.1–21.9) 99.7% (99.5–99.8) 96% 0.68
>1/µl (387) 35.9% (31.2–40.9) 96.9% (96.5–97.3) 35.8% (31.1–40.8) 97.0% (96.5–97.3) 94% 0.33
a
For this analysis we assume that microscopy is the gold standard (i.e., always correct).
Table 2: Performance of PCR relative to expert laboratory microscopy at different Plasmodium falciparum and Plasmodium vivax
parasite densities
P. falciparum P. vivax
Parasites/µl No. detected by microscopy
a
No. positive by PCR (%) No. detected by microscopy
a
No. positive by PCR (%)
>10,000 10 10 (100.0%) 3 3 (100.0%)
5,000–9,999 4 4 (100.0%) 1 1 (100.0%)
1,000–4,999 22 21 (95.5%) 11 11 (100.0%)
500–999 11 10 (90.9%)
b
55 (100.0%)
250–499 17 14 (82.4%) 19 15 (78.9%)
100–249 87 29 (33.3%) 57 34 (59.6%)
1–<99 118 24 (20.3%) 294 71 (24.1%)
Negative 8,321 96 (1.2%) 8,200 251 (3.1%)
Total 8,590 208 (2.4%) 8,590 391 (4.6%)
a
Data for each parasite includes 4 mixed infections
b
A single sample was negative for P. falciparum by PCR, however, this sample was positive for P. vivax and P. malariae by PCR
Table 4: Relationship between PCR and expert laboratory microscopy with Plasmodium falciparum and Plasmodium vivax parasite
densities of less than 100 parasites/µl.
P. falciparum P. vivax
Expert Microscopy Resµlt PCR Positive
a
PCR Negative PCR Positive
a
PCR Negative
Microscopy Positive 24 94 71 223
Microscopy Negative 96 8,225 251 7,949
a
Samples with parasite densities greater than 100/µl by microscopy are not included in this analysis.
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A total of 698 samples that had non-concordant PCR and
microscopy results when initially tested were retested by
both microscopy and PCR and the performance of each
assay calculated by assuming that the initial test result
with each method was correct. The agreement between
first and second test results was significantly better for
PCR (75.1% sensitivity, 91.9% specificity) than for micro-
scopy (33.5% sensitivity, 58.5% specificity). This data
clearly indicates that PCR was the more repeatable
method.
Discussion
Data from this study highlights the problem of using a
less-than-perfect diagnostic test as a reference standard.
Microscopic results were initially considered as the refer-
ence standards for true positive and true negative results,
with all subsequent statistical analysis based on this
assumption. Although there was good agreement between
PCR and microscopy at parasite densities of >500 para-
sites/µl, the majority of positive blood films in the village
of Kong Mong Tha had fewer than 250 parasites/µl blood.
There was extremely poor concordance between micros-
copy and PCR at these relatively low parasite densities.
The poor performance of PCR at low parasite densities
presumably reflected limitations of microscopy as much
as PCR. Many of the PCR results that were considered
false-positives and false-negatives for analysis were pre-
sumably true-positives and true-negatives (i.e., the micro-
scopy result was incorrect). When PCR was considered the
reference standard, the performance of microscopy was
just as poor as that observed for PCR when microscopy
was used as the reference standard.
Because it was difficult to determine whether microscopy
or PCR was the more accurate assay, all non-concordant
samples were retested in order to determine which
method was the more repeatable test (assuming that the
test that was more repeatable was more accurate). A panel
of 698 samples that had non-concordant results was
retested and the performance of each assay calculated by
comparing the results from the second test with those of
the initial test (i.e., the initial test was used as the reference
standard for statistical purposes). The data from this test
clearly demonstrated that PCR was a more repeatable, less
subjective test than was microscopy with parasite densities
of less than 250/µl. All performance criteria (sensitivity,
specificity, PPV, NPV, accuracy and reliability) were much
lower for microscopy than for PCR.
The limitations and shortcomings of microscopy are well-
documented [1,2], with significant problems existing
even in fairly sophisticated laboratories. A rigorous qual-
ity assurance program is essential if performance of micro-
scopy is to be maintained at a high level [2]. The two
microscopists who examined all blood films in this study
have a total of over 35 years of experience reading malaria
slides, and had recently completed and passed a quality
assurance test developed by the Walter Reed Army Insti-
tute of Research (R.S. Miller, personal communication).
In spite of having an expert team of microscopists, this
study highlights the difficulty in conducting an active sur-
veillance program in areas where infection rates and para-
site densities are low. Of the 8,590 blood films that were
collected, 98.7% were either negative (7,925) or had fewer
than 250 parasites/µl blood (556). Both microscopists
examined slides for an average of 12 hours per day for 3
days in a row during each 5-day trip to Kong Mong Tha –
it is not surprising that mistakes were made under these
conditions (long hours examining mostly negative
slides).
Conclusion
PCR is a less subjective test than is microscopy – this was
clearly demonstrated when the set of 698 non-concordant
slides was retested. Each performance indicator (sensitiv-
ity, specificity, PPV, NPV, accuracy and J index) was mark-
edly higher for PCR than for microscopy. Although
sensitivity of PCR can be related to parasite density
[17,28], data indicates that PCR is a viable method for
conducting active malaria surveillance in western Thai-
land.
Authors' contributions
REC, JS and BK were involved in all stages of this study.
SP, BT and AK participated in the coordination of the lab-
oratory work. NM, NR and GZ participated in the coordi-
nation of the field work. RSM, JAV and KT were involved
in the design of the study.
Disclaimer
The opinions of assertions contained in this manuscript
are the private ones of the authors and are not to be con-
strued as the official or reflecting views of the Department
of Defense or the Armed Forces Research Institute of Med-
ical Sciences.
Acknowledgements
Funding for this project was provided by the Military Infectious Diseases
Research Program of the U.S. Army Medical Research and Materiel Com-
mand, Fort Detrick, MD, and by NIAID Grant R01-AI48813-01A1 to Dr.
Jefferson Vaughn.
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