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Molecular and Biochemical Parasitology, 61 (1993) 315-320
© 1993 Elsevier Science Publishers B.V. All rights reserved. / 0166-6851/93/$06.00
Short C o m m u n i c a t i o n
High sensitivity of detection of human malaria parasites by the use of nested
polymerase chain reaction
Georges S n o u n o u *'a, Suganya ViriyakbOSOla'l , Xin Ping Z hu a, William Jarra a, Lucilia
Pinheiro b, Virgilio E. do Rosario , Sodsri T h a i t h o n g and K. Neil B r o w n
aDivision of Parasitology, National Institute for Medical Research, The Ridgeway, Mill Hill, London NW7 1AA, UK; blnstituto
de Higiene e Medicina Tropical/Centro de Malaria e outras Doen~cas Tropicais, Lisbon, Portugal; and CWHO Collaborating
Centre on the Biological Characterization of Malaria Parasites, Institute of Health Research and Faculty of Science,
Chulalongkorn University, Bangkok, Thailand
(Received 27 April 1993; accepted 8 July 1993)
Key words: Plasmodium falciparum; Plasmodium vivax; Plasmodium malariae; Plasmodium ovale; Polymerase chain reaction;
Diagnosis; Mixed infection; Epidemiology
Accurate knowledge of the geographical and
longitudinal distribution of the four Plasmodium species infecting man is of crucial
importance, since they differ greatly with
respect to their biology and clinical manifestations [1,2]. When parasite levels are very low
and in the detection of mixed species infections, the information obtained by microscopy
is restricted, and in some cases biased, by the
inability to devote the necessary amount of
time to the examination of blood smears. DNA
based methods for the detection of parasites,
mainly the clinically significant Plasmodium
falciparum, have been developed in order to
overcome these limitations [3,4]. We have
previously described a PCR method for the
sensitive detection of the four human malaria
parasite species based on the sequence of their
author. Tel. 081-959 3666; Fax: 081-906 4477.
i Present address: University of California San Diego,
Infectious Diseases Section, Medical Services (11 IF), Veterans
Administration Medical Center, 3350 La Jolla Village Drive,
San Diego, CA 92161, USA.
Abbreviations: PCR, polymerase chain reaction; ssrRNA,
small subunit ribosomal RNA.
small subunit ribosomal RNA (ssrRNA) genes
[5]. In this article we report the attainment of
higher sensitivity of detection of these four
Plasmodium species, 10 parasite genomes,
through the use of nested PCR amplification.
We also present a simplified method for the
preparation of the PCR amplification template
from field blood samples.
The amplification scheme employed as well
as the sequence of the oligonucleotides used,
which are all based on the ssrRNA genes [6-9],
are presented in Fig. 1. Two genus-specific
primers rPLU5 and rPLU6, are used for the
first cycle of amplification. An aliquot of the
product thus obtained is used for a second
amplification cycle, in which each parasite
species is detected separately using speciesspecific primers. Experimental details are given
in the legend to Fig. 1. The presence of
amplification product is detected by simple
ethidium bromide staining following agarose
gel electrophoresis. A specific PCR product is
only obtained when DNA from the corresponding species is present in the reaction (Fig.
2A). No amplification is observed with human
DNA alone. The size of the specific PCR
product is different for each of the species: 205
First Amplification Reaction
rPLU 6
4-- rPLU 5
ca 1200 bp
S e c o n d Amplification Reactions
rVIV 1
rFAL 2
rVIV 2
Fig. 1. Schematic representation of Plasmodium ssrRNA genes and the nested PCR protocol used. The species-specific
oligonucleotide primers were designed to hybridise to the genes coding for only one of the two ssrRNA types present in the
Plasmodium genome [6-9]. rOVA2 was obtained from an upublished partial sequence of the ssrRNA gene (kindly provided by
Dr A. Waters, Leiden University, The Netherlands). Synthesis of the oligonucleotides was performed on a 380B DNA
Synthesizer (Applied Biosystems, Foster City, USA). All PCR reactions were carried out in a total volume of 20/A. In all cases
amplification was performed in 2 mM MgCI2/50 mM KC1/10 mM Tris pH 8.3 (HC1)/0.1 mg ml n gelatin/125 #M of each of
the four deoxyribonucleotide triphosphates/ 250 nM of each oligonucleotide primer/ 0.4 unit of AmpliTaq Polymerase
(Perkin Elmer Cetus, USA). 1 #1 of the purified template D N A was used for the first reaction, in which the fragment spanned
by rPLU5 and rPLU6 is amplified. A 1-#1 aliquot from the product of the first PCR reaction was then used as a template in
each of the four separate reactions in which the species-specific primer pairs are employed. The PCR assays were performed
using a heating block (PTC-100, MJ Research Inc., USA). The cycling parameters for the first amplification reaction were as
follows. Step 1, 95°C for 5 min; step 2, annealing at 58°C for 2 min; step 3, extension at 72°C for 2 min; step 4, denaturation at
94°C for 1 rain; repeat steps 2-4 24 times, then step 2, and finally step 3 for 5 min. On termination of the amplification cycle,
the temperature was reduced to 20°C. For the subsequent four species-specific amplification reactions, 30 cycles were
performed as above.
bp for P. falciparum, 120 bp for Plasmodium
vivax, 144 bp for Plasmodium malariae and
approx. 800 bp for Plasmodium ovale. A
fragment of approx. 1.2 kb which is observed
in all reactions corresponds to the product of
the first amplification reaction. For P. ovale
the level of non-specific amplification is
relatively higher than that obtained for the
other three species. It is, however, only
observed in the presence of P. ovale DNA.
Selection of the primers specific for P. ovale
was restricted, since;only a partial sequence of
this parasite's ssrRNA gene was known [9], the
unpublished sequence of another portion of
the gene was very kindly made available by Dr.
Andy Waters of Leiden University (The
Netherlands). A fainter band (Fig. 2) observed
with all four species, of a slightly higher
molecular weight than the specific product, is
an artifact resulting from the presence of an
excess of target D N A (Fig. 2C).
In order to establish the minimum number
of parasites that could be detected, samples
with known numbers of P. falciparum parasites
were employed. These were generated from a
ten-fold serial dilution of highly synchronous,
Fig. 2. Specificity and sensitivity of the PCR detection assay. (A) Nested PCR amplification for the demonstration of the
specificity of the primers employed. Control genomic DNA from P. falciparum (F), P. vivax (V), P. malariae (M), P. ovale (0)
and human blood (H) were prepared as in [5]. Molecular size markers, a 100-bp ladder, flank the experimental lanes. (B)
Nested PCR assay for the detection of in vitro cultured P. falciparum ring stage parasites. The numbers of parasites per
aliquot assayed is given above each lane. (C) Nested PCR amplification using diluted control DNAs. (D) Product of
amplification of diluted control DNAs, using the PCR assay previously described [5]. The control DNAs used in C and D are
diluted as indicated between these two panels, the undiluted stocks for these DNAs have been used for the reactions presented
in panel A. The species-specific oligonucleotide primer pairs used are given for each panel. Electrophoresis of product was
perfomed in 3% (3:1) Nusieve agarose/Agarose gels [5].
ring stage, in vitro cultured parasites, using
whole blood as diluent. Accurate erythrocyte
counts, obtained by flow cytometry, and the
exact parasitaemia in the sample, were used to
calculate the number of parasites per #1 of the
original culture aliquot. PCR template was
prepared by the boiling method described later
in this article. The result of the nested PCR
detection of P. falciparum in these samples is
presented in Fig. 2B. A specific amplification
product was observed in all the samples in
which the expected number of parasites was
one or higher. A constant quantity of specific
product was observed for samples containing
decreasing numbers of parasites. Thus, nested
PCR amplification results in an 'all or none'
detection of parasites in a given sample. Blood
samples from P. vivax, P. malariae or P. ovale
infected patients were not available. However,
the approximate parasite levels present in the
infected chimpanzee blood samples from which
control DNA was purified [5], were known.
The pattern of amplification (Fig. 2C) obtained for the dilution series of these DNAs
(including a series from P. falciparum purified
DNA), was observed to be comparable to that
obtained with the defined P. falciparum
samples. A similar sharp cut-off in the
generation of the specific PCR products,
indicating a similar 'all or none' detection of
the three species, at levels corresponding to
between 50 and 0.1 parasites per sample, was
also observed. Since the number of parasite
genomes present in each sample cannot be
accurately determined, we conservatively conclude that the presence of ten parasites can be
detected by the nested PCR method presented
here. Using the same diluted DNA templates,
we observe an increased sensitivity of detection
by nested PCR (10-fold for P. vivax and P.
malariae, 100-fold for P. ovale), as compared
to that obtained by the previously published
PCR assay (Fig. 2, Panel D). In contrast to the
nested PCR protocol, the intensity of the
amplified fragment obtained by the original
method decreases as the parasite numbers in
the samples diminish.
As a result of the very high sensitivity as well
as the 'all or none' effect of the nested PCR
assay, irreproducibility in parasite detection
was observed when aliquots from samples with
very low numbers of parasites (an average of 1
parasite #1-1 or lower) were analyzed. Successful amplification is only achieved when the
aliquot contains parasite D N A harbouring the
targeted ssrRNA gene. Detection of a particular parasite species was also obtained from
samples in which a minimum amount of DNA
from this species was mixed with a large excess
of D N A from the three other species as well as
human DNA (data not shown). Attempts to
include all the species-specific primers in one
reaction (duplex PCR) resulted in the failure to
detect the expected specific amplification
products from samples harbouring DNA
from the four malaria species.
Preparation of template DNA by phenol
extraction and ethanol precipitation [5] might
be considered unsuitable because of the
hazardous nature of the reagents and the cost
in time and materials it necessitates. A
simplified method for the preparation of
PCR template which lacks these disadvantages has been described [10]. Modifications to
this method, in which the parasites are boiled
to release the DNA, were introduced, and
duplicate samples from 25 of the blood
samples obtained from patients attending the
Malaria Clinic at Borai, a village in Chantaburi province, Thailand [5], were used to assess
the new procedure. For the boiling method
Detection of parasites from blood samples collected in the
Malaria Clinic at Borai (Thailand)
Sample N °
Single PCR a
Nested PCR
ethanol b
ethanol b
F = P. f a l c i p a r u m , V = P. v i v a x , M = P. m a l a r i a e and O
= P. ovale.
aAliquots of these DNA samples had been analyzed by the
previously described PCR assay [5].
bThe results obtained in these two columns were derived
using the DNA templates prepared by phenol extraction/
ethanol precipitation from whole blood kept cold following
collection [5].
~At the end of the 48 hour period of storage of the blood
samples at room temperature, saponin was added to a final
concentration of 0.05%. The parasites released from the
lysed erythrocytes were collected by centrifugation (6000 x
g for 5 rain. at room temperature). The supernate was
discarded and the parasite and leucocyte pellet resuspended
in 25/tl PCR buffer with MgC12 omitted. The mixture was
overlaid with mineral oil and subjected to 99°C for 10 min.
The nested PCR assay described above was then performed
with a 5 #1 aliquot of the boiled solution, and in a parallel
reaction with an equivalent quantity of D N A purified by
phenol extraction/ethanol precipitation from blood simultaneously collected from the same patients.
described here (see Table I), a 25 #1 aliquot
from each whole blood sample (approx. 1 ml)
collected from 25 patients, was taken and
thoroughly mixed with 1 ml of culture medium
with no added serum and in the presence of
heparin, immediately following vene puncture.
Many villages in endemic areas are only
accessible on foot, thus storage of the
collected samples at low temperatures is often
impractical. Also, return to the laboratory on
the day of blood collection is often not
possible. In order to reproduce these circumstances, these 25 samples were left, following
collection, for 48 h at room temperature, which
was 25-35°C, before processing by the boiling
method (Table I). The tube contents were
mixed by inversion 6 times during the storage
period. The remainder of the samples taken
from the same 25 patients were processed as
previously described [5]. Briefly, the samples
were kept at 4°C immediately after collection,
and stored at - 7 0 ° C upon return to the
laboratory on the same day. DNA from these
samples was prepared by phenol exctraction
and ethanol precipitation. The results from the
analysis of the two sets of samples are
presented in Table I. Thus, the results presented in the first two columns are derived
from DNA samples prepared as in [5], and
therefore provide a comparison between the
efficiency of detection using the nested PCR
method with that of the single PCR assay. The
results of the last column were obtained by
nested PCR analysis of the DNA template
obtained by the boiling method.
Parasites were detected in 15 samples, with
one case of mixed infection (P. malariae with
P. falciparum), when phenol/ethanol purified
template was analysed by the single PCR assay
(first column). Using nested PCR amplification
with the same DNA templates, 17 samples
were found to be positive for Plasmodium,
mixed species infections were brought to light
in eight patients, with a triple species infection
in one case and all four species present in
another (second column). When nested PCR
amplification was performed on the templates
obtained by the boiling method, 17 Plasmodium positive samples were also obtained (third
column). However, in this instance a mixed
infection was only observed in two cases.
Failure to detect P. falciparum was observed
in three cases and P. vivax in 5 cases. In one
case P. falciparum had not been detected when
the phenol/ethanol purified DNA was used as
a template. Since very low parasite numbers
were detected following boiling (Fig. 2B), it is
unlikely that failure to amplify the parasite
DNA from the field samples is a consequence
of template preparation by the boiling method.
It is probably due to the storage of the blood at
room temperature before processing. In all
cases, the species that was not detected
following the 48 h storage was present in low
numbers, since it had not been detected by the
previously performed single PCR procedure.
The loss of sensitivity might be explained by
the fact that P. vivax cannot be maintained in
continuous culture, and some lines of P.
falciparum do not grow in vitro. It is thus felt
that the parasites originally present in the
collected blood were lost during the storage
period before template preparation. However,
in the one case (TD 281, Table I) where P.
falciparum was only detected following storage, it is conceivable that maturation of the
parasites throughout the 48-h period at close
to physiological temperature, could have
resulted in an increase of the parasites' DNA
content to the detection threshold. Maturation
of the parasite to the late trophozoite stages,
when DNA is replicated in preparation for
merozoite formation, might result in improved
sensitivity. Therefore, although it is advisable
to process the samples as quickly as possible
following collection, storage at room temperature for 12-24 h before DNA template
preparation, or cold storage, should not result
in an appreciable loss of sensitivity. The results
obtained by PCR were mostly confirmed by
careful and, for some samples, lengthy microscopic examination of blood smears obtained
at the time of collection. In two cases TD 273
and TD 276, the nested PCR diagnosis of P.
vivax and P. ovale could not be confirmed by
microscopy, nor could the PCR detection of P.
falciparum and P. vivax be similarly demonstrated in TD 282 blood smears. It is felt that
this aconsequence of the very sensitivity of the
nested PCR assay, in which approx. 200-fold
more blood cells are examined for the presence
of parasites than by conventional microscopy.
No evidence of cross-contamination was
observed in any of the reactions, which were
repeated more than twice and with the sample
order altered at each duplication. The efficiency of detection of low parasitaemias can be
further improved by the screening of a larger
volume of blood in each reaction. In a separate
experiment, PCR templates were obtained
from field samples by boiling as described
above. These templates were then left at room
temperature for 6 days before use in the nested
PCR assay, without any deleterious effect on
the sensitivity of parasite detection (data not
In conclusion we show that the use of nested
primers allows the detection and identification
of very low numbers of the four human
malaria parasites, without the requirement
for further blotting and hybridisation of the
PCR amplification product. The addition of a
PCR amplification step, is justified because of
the resulting high sensitivity of detection.
Thus, by comparison to the original report
[5], it was found that a greater incidence of the
parasites species, and in particular their
presence in mixed infections, might characterise the malaria epidemiological situation at the
Thai - Cambodian border region. The use of
the boiling method for template preparation
should allow consideration of this technique
for use in a wider variety of epidemiological
and clinical investigations. The use of centrifuges not requiring mains electricity might
allow sample processing to be started under
adverse field conditions. A re-evaluation of the
epidemiological data and the methods of
detection employed is clearly indicated in the
light of these results.
This work was funded by a grant from the
Commission of European Communities, ECAsean Scientific and Technical Cooperation,
Contract number Cl*0634/UK/SMA. We
thank the Malaria Division, Ministry of
Health (Bangkok, Thailand) for their cooperation. We are grateful for the hospitality and
generous collaboration of the staff of Malaria
Region 5 and the Malaria Clinic in Borai,
(Trad Province, Thailand), especially Mr.
Chartchai Palanant and Mr. Dokrak Thongkong. We are also indebted to the members of
staff at the WHO Collaborating Centre for the
Biological Characterization of Malaria Parasites, Chulalongkorn University (Bangkok,
Thailand) for all their help, and in particular
to Miss Napaporn Siripoon for the consistently excellent technical assistance she has
provided. L.P. and V.E. do R. were supported
by The Gulbenkian Foundation, JNICT/
Ci~ncia and INIC (Portugal).
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