Professor Manorama Dev, Dr. Samuel Bhattacharya and Dr. Basudha ...... NJ), the ICT malaria Pftest (ICT diagnostics, Sydney), and the Determine⢠test (Abbot.
ROLE OF NESTED POLYMERASE CHAIN REACTION (PCR) IN THE CLiNICOEPIDEMIOLOGICAL DIAGNOSIS OF MALARIA IN NEPAL
A THESIS SUBMITTED TO THE GRADUATE DIVISION OF THE UNIVERSITY OF HAWAI'IIN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE IN BIOMEDICAL SCIENCES (TROPICAL MEDICINE)
AUGUST 2004
by Moti Lal Chapagain
Thesis Committee: Sandra P. Chang, Chairperson Kenton Kramer William L. Gosnell
ACKNOWLEDGEMENTS
The field work was supported by the East West Center field study grant. It was impossible for me to conduct this work without this support. I would like to express my sincere thanks to Mendl Djunaidy, the Associate Dean, Education Program, East West Center for providing this support.
Prof. Sandra P. Chang, the chairperson and Dr. William L. Gosnell and Dr. Kenton Kramer, the members of my Graduate Committee had helped and guided me throughout this study. It would not have been possible for me to complete this work without their continuous guidance, supervision and feedback. I would like to express my heartiest gratitude to all of them. Dr. William L. Gosnell, had also made me familiar with several laboratory techniques during my training period and took troubles of getting permissions from all concerned authorities and thus made it possible for me to conduct this study. Friendly care and untiring help that he extended to me throughout my stay in the department was commendable. I would like to extend my sincere thanks to him again.
Dr. Karen Yamaga provided me opportunity and resources needed to learn PCR. This was the major impetus for my thesis work. I am really grateful to her for all her help and support that I got throughout my study period.
Dr. Arwind Diwan's encouraging word, caring attitude and willingness to extend support whatever or whenever I needed was unforgettable. I would like to express my sincere gratitude to him.
iii
Todd Sasser and Ann Hashimoto had introduced me the research techniques and protocols and were always available to provide me instantaneous support whenever I needed them. It would not have been possible for me to conduct this study without their continuous help. I would like to take this opportunity to thank both of them. Similarly, Sheila Kawamoto and Karen Amii provided me all help and support that I needed during my stay in the Tropical Medicine department. I am really grateful to both of them.
Professor Sudhamshu Sharma Khanal, the Rector and Dr. Sanjib Sharma, the member secretary of Research Committee at B.P. Koirala Institute of Health Sciences (BPKIHS), Dharan, Nepal were kind to permit me to conduct this study. I am really grateful to both of them.
Professor Manorama Dev, Dr. Samuel Bhattacharya and Dr. Basudha Khanal at the Department of Microbiology at BPKIHS had helped me throughout my field work in Nepal. I am really grateful to all of them.
At last but not the least, Mr. Ranjit Kumar, the Laboratory Technician at BPKIHS took trouble of collecting samples, performing QBC and microscopy and maintaining all records throughout this study. I would like to express my sincere thanks to him.
iv
ABSTRACT
Malaria poses a diagnostic challenge to laboratories of both developed and developing countries. Microscopic examination of Giemsa stained blood smears is the most commonly used method of malaria diagnosis in Nepal with recent introduction of Quantitative Buffy Coat (QBC) at B.P. Koirala Institute of Health Sciences (BPKIHS).
Eighty-five randomly selected samples among 3182 patients referred to Clinical Laboratory Services (CLS) at BPKIHS for aBC malaria testing from June 18, 2002 to July 17, 2003 were analyzed by aBC malaria test, Giemsa stained thin smear microscopy and nested PCR. Out of 85 samples, 48 (56.47%), 17 (20.00%) and 24 (28.24% ) were positive by malaria QBC test, microscopy and PCR, respectively.
Among 24 Plasmodium genus specific PCR positive samples, 12 (50.00%) were P. falciparum, 7 (29.16%) were P. vivax, 4 (25.00%) were mixed infection with P. falciparum, and P. vivax.
One sample (4.17%) was positive for malaria by genus
specific PCR but was negative for all species specific PCR. Three P.falciparum and one P. vivax microscopic positive samples were found to have mixed infection with both P.falciparum and P. vivax by PCR.
Although aBC was able to pick up all microscopy positive cases, a large numbers of QBC positive cases were negative by both microscopy and PCR suggesting that QBC may have a high false positive rate.
v
The results of this study indicated that malaria was seasonal in Nepal, was much more common among males and maximum numbers of malaria positive cases were found in the age group of 15-30 years.
As expected, PCR detected much more cases than the microscopy technique and more importantly many more mixed infections were identified by the PCR in this study. However, the nested PCR was a very labor intensive procedure. Since, the existing microscopy has low sensitivity, and the aBC has very high false positive rate, PCR would be the most reliable alternative for clinical and epidemiological diagnosis of malaria in Nepal. However, its routine use in the clinical laboratory in Nepal in the present form does not appear to be feasible because of its high cost, long processing time and requirement to run several PCR cycles for each samples. However, it could be a valuable tool for quality control as it can be used in selective samples to verify the microscopy or QBC test results.
vi
TABLE OF CONTENTS
Acknowledgement.
iii
Abstract................................................................................................... . v Table of Content.
,
,
vii
List of Tables
ix
List of Figures
x
Abbreviations
xi
Chapter 1. Malaria: Introduction
1
History
,
1
Global situation
2
Malaria in Nepal.
3
The Plasmodium parasite
4
Transmission and Life cycle
5
Plasmodium genome
9
The vectors
,
,
11
Malaria: Pathophysiology and clinical course
12
Malaria: Diagnosis
14
Microscopy
15
Conventional microscopy
15
aBC and fluorescence microscopy
17
Malaria Rapid Diagnostic Devices (MRDD)
21
Molecular methods
26
Chapter 2. Objectives
30
vii
TABLE OF CONTENTS (CONTINUED) Chapter 3. Materials and Methods
31
Study site and study population
31
Sample collection, storage and transport
32
Diagnositic tests performed
33
aBC™
malaria Test.
Microscopy
33 "
,
'"
Polymerase Chain Reaction
'"
34 34
DNA extraction
34
DNA amplification
35
Identification of amplification products Chapter 4. Results
38 39
Sample profiles
42
aBC malaria test results
42
Microscopy results
43
PCR results
45
Plasmodium Genus specific PCR
45
Plasmodium Species Specific PCR
47
Summary of all diagnostic test results
50
Sensitivity and specificity
52
Chapter 5. Discussion and conclusions
55
Appendix: Profile and results summary of the sample population
59
References
62
viii
LIST OF TABLES
Tables 1.
Selective recent QBC studies
19
2.
Selective recent studies of MROOs
25
3.
Summary of the Selective recent PCR studies
28
4.
Monthly distribution of aBC request and aBC positive cases
.40
5.
Sample population with their mean age (SO) and sex distribution
.42
6.
aBC positive cases according to sex and relative parasitemia
.45
7.
Comparison of the results of the microscopy and species specific PCR .... .47
8.
Number and percentage of malaria positive and negative cases
50
9.
Sex distribution of malaria positive cases
50
10. Age distribution of the sample population and positive cases
51
11. Mean age (SO) of the positive and negative sample population
51
12. Performance of aBC when compared with microscopy
52
13. Performance of PCR as compared with microscopy
53
14. Performance of aBC compared with the combined Performance of both microscopy & PCR
54
IX
LIST OF FIGURES
Figures 1.
Page Expected sizes of the amplified DNA of Plasmodium species, Agarose gel (2%) electrophoresis of nest 2 PCR products
38
2.
Monthly distribution of aBC request and aBC positive cases
39
3.
Monthly distribution of aBC positive cases
40
4.
Proportion of different species of Plasmodium causing malaria in patients attending to BPKIHS
44
5.
Proportion of falciparum and non-falciparum malaria in Nepal (2001) .........44
6.
Agarose Gel electrophoresis with Plasmodium Genus specific Nest 2 PCR product S
46
7.
Agarose gel electrophoresis with P.falciparum specific primers
.48
8.
Agarose gel electrophoresis with P. vivax specific primers
49
x
ABBREVIATIONS AMP
Apical Merozoite Antigen
An
Anophelene
AO
Acridine Orange
BC
Before Christ
BPKIHS.
B.P. Koirala Institute of Health Sciences
CD
Cluster of Differentiation
CIDR
Cystine Rich Interdomain Region
CSP
Circumsporozoite protein
DALY
Disability Adjusted Life Year
DBF
Duffy Binding Like
DBP
Duffy Binding Protein
DNA
Deoxyribose Nuceic Acid
EBA
Erythrocyte Binding Protein
EDTA
Ethylene di-amine tetra acetic acid
ETS
External Transcription Spacers
HRP
Histidine Rich Protein
HSPG
Heparan Sulfate Proteoglycan
LSU.
Large Subunit
MRDD
Malaria Rapid Diagnostic Device
MSP
Merozoite Surface Protein
NBP
Normocyte Binding Protein
P
Plasmodium
PCR
Polymerase Chain Reaction
Xl
aBC
Quantitative Buffy Coat
RBC
Red Blood Cells
RBP
Reticulocyte Binding Protein
Rifin
Repetitive interspersed family
RNA
Ribonucleic Acid
rRNA
Ribosomal RNA
RT-PCR
Reverse Trascript Polymerase Chain Reaction
SSU
Small Subunit
STEVOR
Subtelomeric Variable Open Reading Frame
TNF
Tumor Necrosis Factor
WHO
World Health Organization
xu
CHAPTER 1 MALARIA: INTRODUCTION
1.1 HISTORY
The word "malaria" was derived in seventeenth century from an Italian word "mal'aria" meaning bad air because of its association with the ill smelling vapors from the swamps near Rome [1]. Malaria is probably one of the oldest diseases known and its history can be traced back almost to the beginning of the history of the mankind. Fossils studies revealed that the mosquitoes, the vector of malaria were present in Africa up to 30 million years ago [2].
Egyptian mummies more than 3000 years old with enlarged
spleen presumably due to malaria has been found [1] and malaria antigen was detected from the skin and lung samples of mummies from 3200 and 1304 B.C [3]. Several ancient writers described many features now known to be characteristics of malaria in their writings and it is most likely that humans and malaria have probably evolved together [1].
Although people were likely to be exposed to malaria since ancient times, virulent malaria probably originated relatively recently and started having a major impact on human survival since the advent of Agriculture approximately 10,000 years ago. Malaria was probably originated in Africa and accompanied the human migration to the Mediterranean shores, India and South East Asia [4] and had possibly spread to southern Europe via the Nile Valley. It might have also reached Europe from Asia because of the close contact between Asia Minor and European people [1]. The intermittent fever, probably due to malaria was common in Europe by the 15th century. 1
Although malaria was unknown in the New World before the Columbus era, Plasmodium vivax and P. malariae were probably introduced to the Americas by European explorers,
whereas P. falciparum accompanied the African slaves. Malaria became worldwide by the early 18th century [1].
1.2 MALARIA: GLOBAL SITUATION Malaria transmission was largely controlled in many parts of the world during the 1950s and 1960s by the World Health Organization (WHO) led malaria eradication campaign coupled with improvement in sanitation and hygiene. However, this success was not sustained because of the cost of the program, community resistance to repeat spraying of their houses and emergence of resistant vectors and parasites [5] and the malaria eradication campaign was officially declared a failure in 1972 [6]. As a result malaria has returned to areas from which it had been eradicated, and is also spreading into new areas, such as Central Asia, and Eastern Europe. The world is now facing a rapidly increasing malaria burden and more people are now dying of malaria than thirty years ago. Currently, malaria is the most important tropical parasitic disease. It is a public health problem in more than 90 countries and over 2.4 billion people comprising 40% of world's population live in malaria endemic areas and are at risk of getting this deadly disease [7]. There are an estimated 300 to 500 million clinical cases of malaria every year resulting in 1 to 3 million deaths, mostly of children [8]. According to WHO, malaria was responsible for 2.2% of all deaths in 2003 and there were an estimated 1.27 million deaths worldwide attributable to malaria [9].
Eighty percent of all clinical cases and
about 90% of all parasite carriers of malaria infection occur in Sub-Saharan Africa [7]. P. falciparum, is the predominant species of malaria in Africa. It is less common in other
2
parts of the world. P. vivax, is responsible for over 50% of malaria cases outside the Africa and accounts for 70-80 million clinical cases annually [10]. P. vivax is common in Southeast Asia, Central and South America, and particularly on the Indian subcontinent. It also accounts for around 10% of cases in Eastern and Southern Africa but less common in West Africa, presumably due to the presence of Duffy-negative blood group variants [10].
Although, malaria remains largely under control in most developed countries, increasing international travel has been contributing to imported malaria in these countries.
In
recent years, malaria epidemics have occurred in several countries and the global situation is worsening. Malaria is the second most common killer tropical communicable disease and is the second major cause of loss of Disability Adjusted Life Years (DALYs) among the infectious diseases. In any given year, nearly ten percent of the global population will suffer a case of malaria and in many African countries about 10% of hospital admissions are for malaria, as are 20-30% of doctor's visits [7].
1.3 MALARIA IN NEPAL Nepal has vast plain, inner valleys and terrain with tropical and subtropical climate supported by high monsoon rain. Prior to the early 1950s, many inner valleys and terrain were full of forest and the condition was very favorable for malaria. It was estimated that approximately two million malaria cases occurred in Nepal annually during the early 1950s with case fatality rate of 10-15% [11].
3
Malaria control projects were initiated in Nepal in 1954 and 1956 and a malaria eradication program was launched in 1958. A remarkable success was achieved from the inception until the late 1960s and the number of recorded cases dropped to only 2,500 in 1970. However, this success could not be sustained and currently, about 70 percent of Nepal's 25 million people live in areas with unstable malaria transmission. There are an estimated 30,000 clinical cases with over 15 deaths per annum in Nepal although less than 10,000 cases are reported annually. Although rare in the past, Nepal has witnessed increasing outbreaks of P. falciparum malaria resistant to chloroquine in recent years. Moreover, P. falciparum resistant to the second line drug SulfadoxinePyrimethamine has also been documented in the outbreak areas [11].
1.4 THE PLASMODIUM PARASITES
The protozoan parasite Plasmodium species are the causal agents of malaria. Plasmodium genus is classified in the order Haemosporidia, class Hematozoa, phylum
Apicomplexa [12]. The Genus Plasmodium consists of nearly 200 known species that infects reptiles, birds and mammals including the four species causing human malaria. The evolutionary origin of Plasmodium is disputable. It is not very clear whether the Plasmodium species were derived from lateral transfer from other vertebrate parasites or
evolved directly from ancient parasites of the marine invertebrates from which the chordates evolved [12]. Pathophysiological understanding of malaria began unfolding when Charles Louis Alphonse Laveran discovered the malaria parasite in 1880 and Ronald Ross, a British surgeon working in India demonstrated that malaria was transmitted by mosquitos [1].
4
1.4.1 Transmission and Life Cycle
The Plasmodium parasite has a very complex lifecycle, involving asexual stages within human hepatocytes and erythrocytes and a sexual stage within the mosquito gut. Malaria is typically transmitted by the bite of an infected female Anopheles mosquito, however, it can also be transmitted transplacently during pregnancy, by blood transfusion or needle sharing [13]. When an infected Anopheline mosquito bites a human host, salivary fluid containing the sporozoites is inoculated into the subcutaneous tissue or blood stream. Sporozoites then travel through the blood stream to the liver microcirculation, where they come into contact with the sinusoidal cell layer consisting of endothelial and Kupffer cells. These cells separate the sporozoites from their target hepatocytes and it is unclear how the sporozoites reach the hepatocytes as the sporozoites (1/lm in diameter) are larger than the fenestrations (pore sizes 0.1 /lm) in the sinusoidal cell layer [14]. It has been suggested that the Kupffer cells may engulf the parasites and the parasites may traverse the Kupffer cells to reach hepatocytes [14]. Sporozoites may invade hepatocytes using region II of the circumsporozoite protein (eSP-II) on their surface as the ligand for binding to heparan sulfate proteoglycans (HSPGs) and the low-density lipoprotein receptor on the hepatocyte surface [15]. HSPGs extend from the Disse space to the sinusoidal lumen through endothelial fenestrae and may help the sporozoites to specifically target the hepatocyte [14]. Once inside the hepatocytes, these sporozoites mature into liver-stage trophozoites and then into schizonts
before rupturing the infected liver cell and releasing 20,000-40,000
merozoites into circulation in about 10 to 15 days depending upon the species [16]. Tissue schizonts are a central feature of all four plasmodial species that infect humans; they amplify the infection by producing large numbers of merozoites from each sporozoite-infected hepatocyte. During the exoerythrocytic cycle some of the sporozoites
5
of P. vivax and P. ovale may differentiate into hypnozoites and remain dormant for several weeks to months after the initial infection. When these dormant hypnozoites mature to tissue schizonts and release infectious merozoites, they produce a symptomatic blood stream infection and cause relapsing malaria [14].
The merozoites are small, ovoid cells 1.5-2.5 J..lm long and 1.0-2.0 J..lm wide, having slightly narrower apical pole. Three types of membrane bound organelles namely rhoptries, micronemes and dense granules are located at the apical pole and appear to play a role in binding and entry of the merozoite into RBCs [16]. Each merozoite released from the liver is capable of invading a human red blood cell and establishing the asexual cycle of replication in that red cell with the release of 24 to 32 merozoites at the conclusion of a 48- or 72-hour asexual cycle. All clinical symptoms of malaria are caused by this asexual erythrocytic cycle of Plasmodium. P. vivax merozoites exclusively invade the reticulocytes whereas P. falciparum merozoites can invades RBCs of all ages and thus can produce severe parasitemia [16,17].
Invasion of erythrocytes by the merozoites is a complicated process involving several steps [17]. Many merozoite ligands and RBC membrane receptors are likely to be involved in this invasive process but the details are still sketchy. It appears that P. vivax reticulocyte binding proteins 1 and 2 (PvRBP-1 and PvRBP-2) specifically target the reticulocytes and initiate loose attachment between the merozoite and erythrocyte. This initial interaction between merozoite and RBC enables the merozoite to reorient its apical pole towards the red cell surface and may trigger the release of Duffy Binding Protein (DBP) from the micronemes of P. vivax merozoite. The DBP of P. vivax merozoite binds with the Duffy factors on the erythrocytes and forms a tight junction and 6
initiates invasion. Similarly, the Normocyte Binding Proteins (NBPs) of P. falciparum merozoites help their merozoites specifically to target normocytes and initiate loose attachment between the merozoite and erythrocyte. This initial interaction between merozoite and RBC enables the merozoite to reorient its apical pole towards the red cell surface and may trigger the release of Erythrocyte Binding Antigen-175 (EBA-175) from the micronemes of P.falciparum merozoites. The EBA-175 specifically binds with the sialic acid residue of
Glycophorins A on the erythrocyte and initiates invasion of
merozoite [16,17].
RBC invasion process of the merozoite consists of four steps: a. Initial recognition and loose attachment of merozoite to erythrocyte membrane. b.
Juxtrapositioning of apical prominence of merozoite to the RBC membrane and formation of tight junction.
c. Movement of junction and invagination of RBC membrane d. Resealing of parasitophorous vacuole membrane and RBC membrane after completion of invasion [16].
Additionally, Merozoite surface proteins (MSP) and Apical Merozoite Antigen (AMP) particularly MSP-1 and apical AMA-1 also playa role in erythrocyte invasion [16,17].
Alternatively, some intraerythrocytic parasites develop into the sexual (gametocyte) forms necessary to complete the life cycle in the Anopheline vector. Although the studies on Plasmodium gametogenesis are not consistent, they together suggest that the Plasmodium is committed to sexual development in the preceding asexual schizogony
7
rather than differentiating following invasion of the erythrocytes by uncommitted merozoites [18,19,20]. Both innate and environmental factors may play an important role in conversion of asexual parasites to the sexual development [18,19]. Although the exact role of different environmental factors on
gametogenesis is not clear
immunological stress, host response to the parasites and cell debris and chemotherapy may play important roles
[21].
It has shown that chloroquine
may induce
gametocytogenesis whereas sulfadiazine and sulfamethazine may induce extended period of high gametocytemia [21]. Immature gametocytes of P. falciparum are sequestered in the body tissues and it has been suggested that this sequestration is mediated by expression of a
gametocyte-specific variant of var gene family, which
adheres to CD36/1CAM on endothelial lining of capillaries [21]. Sequestration prevents the clearance of gametocyte infected RBCs by spleen and other phagocytic tissues [21]. It appears that the mature gametocytes are arrested in Go phase of cell cycle and are thus resistant to most antimalarials. They may remain in the blood stream of human host even after the treatment with standard schizonticides and may continue to infect the mosquito vector [21].
When mature gametocytes are taken up by a female Anopheline mosquito with a blood meal, microgametocytes and macrogametocytes mature inside the mosquito gut to form male and female gametes respectively. The microgamete then fertilizes macrogamete and produces a diploid zygote. The zygote matures to an ookinete and undergoes a meiotic reduction division to produce the haploid sporozoites that migrate to the salivary gland and are prepared to infect the human host during the next blood meal [16].
8
1.4.2
Plasmodium genome
Malaria parasites are extremely successful protozoan parasites. During their complex life cycle, they propagate in very distinct environments and invade different cell types [22]. Genome mapping would provide an insight about the genes involved in parasite/host interactions. An international effort was initiated in 1996 to sequence the genome of the P. falciparum clone 307 and its genome sequence was published in 2002 [23]. Similarly,
the P. vivax clone Salvador I, is currently being sequenced using a whole genome shotgun strategy [24]. The P. vivax nuclear genome is estimated to be 30 Mb in size and is distributed among 14 chromosomes. It is estimated that the G+C content of the P. vivax genome is about 45% and is being sequenced using a whole genome shotgun
strategy [24].
The P. falciparum 307 has nuclear genome of 22.8 Mb distributed over 14 chromosomes, ranging in size from 0.643 to 3.29 Mb. There are approximately 5,300 protein encoding genes with an average gene density of 4,338 bp and mean gene length of 2.3 kb when introns are excluded. P. falciparum genome is the most A+T rich genome sequenced to date and the overall A+T content is 80.6% which is even higher (-90%) in introns and intergenic regions [23]. The P. falciparum genome is haploid throughout the majority of the life cycle and also contains a 5.9 kb mitochondrial DNA, and a plastid-like 35kb circular DNA [23].
The P. falciparum chromosomes are bounded by telomeres [22] and vary significantly in length resulting mostly from the variation in the subtelomeric regions.
9
Subtelomeric
regions vary probably because of recombination between different parasite clones during meiosis in the mosquito. 8everal genes responsible for antigen variation are located in the subtelomeric region. P.falciparum has 208 genes (3.9%) known to be involved in immune evasion including the three highly variable families of genes: var, fit, and stevor [23].
There are 59 var genes coding for P.falciparum erythroid membrane protein 1 (PfEMP1) that are exported to cell surface of infected RBCs and mediate adherence to the host endothelial receptors and thus promote sequestration. PfEMP-1 proteins are the target of protective antibody response and transcriptional switching between var genes permits antigenic variation and provides a means of immune evasion. The var gene products play important roles in pathogenesis of malaria. The var genes have three recognizable domains: 'Duffy binding like' (DBL), 'cystein rich interdomain region' (CIDR) and 'constant2' (C2) [23].
The fit genes which are 149 in numbers encode for rifin (repetitive interspersed family) proteins. The rifin proteins are also expressed on the surface of infected RBCs and contribute for antigenic variation. The 28 stevor genes encode the sub-telomeric variable open reading frame (stevor) and are less polymorphic than rifins. The exact function of the rifins and stevors is not known [23].
Plasmodium parasites contain several single 188-5.88-288 rRNA units distributed on
different chromosomes. Expression of each rRNA unit is developmentally regulated and different sets of rRNA are expressed at different stages of life cycle. 8-type rRNA genes are expressed primarily in mosquito vectors during the sexual stages whereas A-type 10
genes are expressed during the asexual cycles in the human hosts [23]. The genes within the unit are always identically placed, separated and flanked by spacer regions. The gene order within the unit is thus External Transcribed Spacer (ETS), 18S, Internal Transcribed Spacer 1 (ITS1), 5.8S, ITS2, 28S and ETS and the unit is transcribed as a single polygenic transcript. The transcript is then cut to yield the three mature rRNA molecules: 5.8S, 18S & 28S. The 18S and 28s are called small (SSU) and large (LSU) subunits respectively [25]. Seven loci encoding rRNA were identified in the P. falciparum genome [23].
Almost two-thirds of Plasmodial proteins are unique to this parasite reflecting greater evolutionary distance between Plasmodium and other eukaryotes. Very little is known about the genomes of other Plasmodium species.
1.5
THE VECTORS
Human malaria parasites are transmitted exclusively by a few species of a single mosquito genus, Anopheles [26]. Although, there are more than 500 species of Anopheles mosquitoes only fewer than 50 are competent to transmit Plasmodium parasites to human [27]. An. gambiae, An. arabiensis, An. funestus are the main vectors responsible in Africa whereas An. albimanus, An. culicifacies, An. dirus, An. anthropophagus are the main vectors responsible for malaria transmission in the rest of the world. Four main species of Anopheles mosquitoes incriminated as the vectors of malaria in Nepal are: An. minimus, An. f1uviatilis, An. maculates, and An. annularies. These vectors greatly differ in their ability and efficiency to transmit malaria because of their differences in habitat, feeding habit and life cycle [11].
11
1.6
MALARIA: PATHOPHYSIOLOGY AND CLINICAL COURSE
Malaria is an intravascular infection caused by one or more of the four Plasmodium species that infect humans. However, it is commonly caused by either P. falciparum or P. vivax, and in about 5 to 7% of cases it is due to mixed infection caused by more than one species. P. falciparum may produce overwhelming parasitemias by invading red cells of all ages [28] and it poses a much greater risk of death than do P. vivax, P. ovale, or P. malariae infections [15]. Cytoadherence and sequestration are central to the
pathophysiology of P. falciparum infection, occurs mainly in the venules of vital organs including the brain, heart, kidney and intestine [29,30]. As P. falciparum parasites mature, knobs appear on the surface of the parasitized red cells and facilitate the cytoadherence of P. falciparum-parasitized red cells to endothelial cells in capillaries and postcapillary venules of the vital organs and thus responsible for vital organs failure[31].
P. ovale and P. vivax are clinically and morphologically very similar and they can be considered identical for clinical purposes. P. vivax and P. ovale infect only reticulocytes which constitute only 1% of normal RBCs in the blood and the magnitude of the parasitemia they produce is often low [28,32]. However, the hemolysis associated with P. vivax or P. ovale infection may stimulate hematopoiesis increasing the number of reticulocytes in the circulation. This may permit parasitemias substantially greater than 1 or 2%. However, P. vivax or P. ovale parasitized red blood cells have no knobs and thus they do not cause microvascular complications [30].
12
P. malariae rarely produces acute illness in normal hosts, often causes chronic infection with low parasitemia that may persists for 20-30 years or more but does not cause microvascular diseases. However, it can cause an immune complex glomerulonephritis [15].
Cyclic fevers are the hallmark of malaria and typically have three stages: The first "cold or chilling stage" that lasts for about 15 minutes to several hours, the second "hot stage" lasting for several hours, and the final "sweating" stage associated with diaphoresis, resolution of fever, and marked fatigue. The fever typically occur at the time of red blood cell schizonts rupture to release merozoites and occurs every 48 hours with P. vivax or P. ova/e infection (tertian malaria) and every 72 hours with P. rna/ariae infection (quartan
malaria). Although P. fa/ciparurn also has about 48 hours parasite cycle, it often produces continuous fevers with intermittent irregular spikes [15]. The exact cause of fever
in
malaria
is
not
known.
However,
it
is
believed
that
the
glycosyl
phosphatidylinositol anchor that connects parasite proteins to the parasite or red blood cell surface is exposed (or released) at the time of merozoite release and stimulates the production and release of TNF-alpha by macrophages [33,34].TNF-alpha then produces the fever associated with synchronous parasite release at the end of the asexual cycle[34].
The human host is usually asymptomatic during the pre-erythrocytic and the first few erythrocytic cycles of the parasites. Then, the host has vague non-specific symptoms such as malaise, headache, myalgia, weakness and anorexia at about 11 days after the sporozoites inoculation when there are an estimated 20-50 parasites/".t1 of blood that are
13
detectable by microscopy [35]. The fever usually begins two days later, when an adult harbors about 108 malaria parasites in the body (20-20,000 parasites/!!I). However, in endemic areas people with partial immunity usually tolerate
parasitemias up to
10,000/!!1 or 107 per ml of blood without feeling ill [36,37]. P.falciparum may be associated with several vital organs failure including cerebral malaria, renal failure, pulmonary edema, hypoglycemia and severe anemia and is often life threatening [15]. Similar to Plasmodium falciparum, P. vivax may also cause severe anemia, but major complications like cerebral malaria, hypoglycemia, metabolic acidosis and respiratory distress do not occur [10]
1.7
MALARIA: DIAGNOSIS
Accurate diagnosis is the key to the effective management of any infectious disease including malaria. However, malaria presents a diagnostic challenge to laboratories of both developed and developing countries. Endemic malaria, population movements, and travelers coupled with the changing patterns of morphological appearances of malaria species due to drug pressure or strain variation, all contribute to this challenge [38]. There has been a rapid development in the field of malaria diagnosis in the last couple of decades and currently, several malaria diagnostic options are available although, most of these options are feasible only in the research laboratories in the developed world. Current methods of malaria diagnosis include light microscopy, fluorescent microscopy and Quantitative Buffy Coat (QBC), flow cytometry, automated blood cell analyzers, detection of antigens and antibodies, molecular methods and laser adsorption mass spectrometry [39].
14
1.7.1 Microscopy 1.7.1.1 Conventional Microscopy
Malaria parasites could be demonstrated in the patient's blood either by the time honored conventional microscopy or a more recently introduced fluorescent microscopy. Microscopic examination of blood smears stained with Giemsa, Wright's, or Field's stain has been the method of malaria diagnosis ever since the discovery of malaria parasites by Laveran in 1880 [1] and introduction of the thick smear by Ronald Ross in 1903 [40].
The method is simple, does not require highly equipped facilities and often enables differentiation among four human Plasmodium species while providing parasite density and stage [39]. Although, blood obtained by pricking a finger or earlobe is the ideal because of the higher density of developed trophozoites or schizonts in blood from these capillary-rich areas [41], blood obtained from venipuncture and stored with EDTA may conveniently be used for malaria diagnosis [38].
Thick blood film concentrates the layers of RBCs by 20-30 times and provides enhanced sensitivity detecting up to five parasites/Ill [42]. Thus, it is better suited to detect low level of parasitemia, recrudescence or relapse [38]. However, lysis of RBCs during the staining may make species identification more difficult [38]. Thin blood film provides the fixed monolayer of RBCs with preserved morphology and makes this procedure more specific than thick film in identifying the Plasmodium species. However, thin blood film is much less sensitive than the thick blood smear [38].
15
Although several methods for estimation of parasitemia have been described using either the thin or thick blood films [43], there is no single, universally accepted standard method for the quantification of parasites and parasitemia is often estimated using different methods, making it difficult to compare the performance of the different diagnostic tools [44]. Expression of the parasitemia as a percentage infection of RBC is probably the method of choice particularly for non-endemic areas, where parasitemia is usually low [38]. Since, 1t11 of blood normally contains 5 x 106 RBC; a 1% parasitemia will contain 50,000 parasites/til of blood. This may be corrected to exact counts if the total RBC count per microliter is known [38].
Routine microscopy, when examined carefully by an expert microscopist is a sensitive (can detect as low as 10-30 parasites per III of blood) [41], relatively inexpensive (US $ 0.12 to 0.40 per slide examined) method that can differentiate among various species and also provide a permanent record useful for quality control [45]. This conventional light microscopy of a well-prepared and well-stained thick and thin blood films has thus stood the test of time and is still considered the "gold standard" for malaria diagnosis [46] as no other reliable and relatively rapid method for the detection and quantification of the parasite burden has been available [47].
However, there is no international agreement on the precise number of microscopic fields to be examined [48] and the microscopic "gold standards" varied widely [49] making it difficult to make valid comparisons. It is a labor intensive, time-consuming procedure requiring well-trained and well-supervised technicians, a condition often difficult to meet in many developing countries particularly in rural settings [45]. There are
16
often long delays in providing the diagnostic results to the clinicians and clinical decisions are often taken without the benefit of the results.
Moreover, substantial
number of patients with low levels of parasitemia may be missed by this method [50]. Screening the blood donor by this method can not reliably prevent transfusion transmitted malaria as many asymptomatic blood donors in endemic areas may harbor the parasites at low levels which may not be detectable by routine microscopy [51] but still be high enough to cause disease in the blood recipients. Moreover, the sensitivity actually achieved in the busy routine examination clinics is 10 fold or more lower (-100 parasiteshll) than that achieved in the research laboratories by expert microscopists [52,53].
1.7.1.2 QBC and Fluorescent microscopy Alternative microscopic methods for malaria diagnosis have been introduced with the hope of overcoming the limitations of conventional microscopy. aBC® (Quantitative Buffy Coat) (Beckton Dickinson, Franklin Lakes, NJ) is one such system [54]. In QBC, 55-65!!1 of whole blood is centrifuged at 12,000 rpm X g for 5 minutes in a commercially
supplied capillary tube coated internally with Acridine Orange (AO) [55]. The tube is then placed on the ParaViewer® microscopic tube holder and scanned for parasites in the interface between RBCs and Granulocytes using a standard white light microscope equipped with the ParaLense® U-V Microscopic adaptor. Since Acridine orange (AO) has an affinity for the nucleic acids, it differentially stains the malaria parasites [56] and since malaria infected RBCs, being lighter than the normal RBCs but heavier than the granulocytes, concentrate in the upper most layer of the erythrocytes during centrifugation [57]. Moreover, a cylindrical float that has the same density as that of
17
WBCs is also inserted into the capillary tube and thus compresses the buffy coat and the lighter portion of the RBCs against the wall of the capillary tube during centrifugation
[58]. Thus, a large volume of potentially infected blood could be observed through the capillary tube using a special, long-focal-Iength objective (paralens) with a fluorescence microscope [58,59].
Several trials using aBC techniques have been performed under laboratory and field conditions [54,58,59,60,61]. It has been claimed that the malaria diagnosis by aBC was unambiguous [58], easier to use and faster than the conventional microscopy. The aBC often detected more malaria cases than the microscopy and thus may have higher sensitivity than conventional microscopy (Table 1) [58,59,60,61].
The malaria detection limit of the aBC was substantially lower than that of the microscopy and aBC detected infection when concentration of samples was halved a mean of 3.3 times beyond the dilution detectable by conventional microscopy [58]. However, negative aBC results in samples haVing high parasitemia have also been reported [62]. aBC is more technically demanding and requires specialized equipment to separate the cell layers by centrifugation and a good fluorescence microscope and is costly (-2.25$ per test vs. $0.02 for field stain) [61]. There is no standard method of parasite quantification in aBC although attempts have been made to express relative quantity of parasites in the samples [60,61]. Pinto et al [60] used the plus (+) system as follows: + (1+) ++ (2+) +++ (3+) ++++ (4+)
-< 1 parasite per aBC field -1-10 parasites per aBC field - 11-100 parasites per aBC field - > 100 parasites per aBC field
18
Table 1. S
'f the find·
Country, sample population, & size (N) France, malaria suspected N=529 Brazil Symptomatic Hospital based samples from endemic areas & asymptomatic volunteers, N=402 India, Symptomatic patients attending clinic from endemic areas, N=2274 Colombia, low endemic, field samples N=833 GSTF=833 aBC=749 PCR=448 Kenya, Seasonal malaria, symptomatic, patients attending malaria clinics N=213
f
Select"---
------
t QBC stud·
Reference standard & Positive cases no&(% J PCR =136 (25.71%)
QBC Positive No. (%)&/or sensitivity & specificity 104 (19.66%)
GSTF
Sensitivity= 91.7% Specificity=88.95 0/0
Leishman stained thick & thin smears =239 (10.5%) GSTF (100 fields) = 97 (11.64%) PCR=98 (21.87%)
328 (14.6%)
Field & Giemsa stained thick films
60 (28.17%)
52 (24.41%)
108(14.42%)
Comments
Ref.
All 32 aBC -ve but PCR +ve cases were also -ve by microscopy None of the PCR -ve sample was +ve by aBC
[63]
HnPCR sensitivity= 97.4% Simple PCR= 84.6% PCR was +ve in 24% of asymptomatic blood donors, aBC was negative in all. Positive rate in asymptomatic individual from endemic area were 28.5%, &20.1 % for HnPCR & aBC respectively Species identification was not possible in 26 (7.9%) of cases. Examination time (microscopic) required for aBC was only 1-2 minutes but smear examination required 10-12 minutes. All aBC -ve cases were also neaative by microscopy aBC had sensitivity (9.8%) & specificity (94.7%) when compared with PCR 10 samples were positive by both aBC & microscopy but negative by PCR, 5 samples were positive only by aBC (? False positive) 19.6% of GSTF & 77.2% of aBC -ve samples were +ve by PCR Sensitivity= 88-98% (overall) 98-100% HP (parasites >320/J.11 of blood) 50-90% LP ((parasites ::;;320/J.11 of blood) All were Pf, 2% of aBC tubes cracked or spilled The agreement between two examiners was low, particularly lower parasitemia (HP=98%, LP=60%)
[51 ]
GSTF=Giemsa Stained thick/thin blood smears, QBC=Quantitative Buffy Coat, PCR= Polymerase Chain Reaction, HP= High Parasitemia, LP= Low Parasitemia
19
[60]
[64]
[61]
Alternatively, Lima et al [61] graded the parasitemia as ''few''
(~
2 parasites per field)
~,
"Moderate" (3-5 parasites per field) or "many" (> 5 parasites per field). However, the practical significance of these classifications in terms of their correlation with in vivo parasitemia is unknown.
The aBC test has been suggested to have a high rate of false positive [61], thus it has not yet generally accepted as a standard method for malaria diagnosis [51]. Although aBC capillary tubes are still commercially available, aBC equipment is no longer marketed [65]. The relative merits of QBC method versus standard light microscopy have been controversial [64, 51]. Although, aBC was claimed to be an unambiguous method" agreement between two examiners in interpretating the aBC was low and in one study as high as one third of the microscopy negative cases were reported positive by one examiner and negative by the other [61]. Furthermore, in QBC, parasitemia could not be easily quantified, samples could not be stored for latter examination and species identification has been problematic [58,66 ] without seeing the RBC morphology and inclusions [51]. Moreover, AD is a very intense fluorescent stain and it nonspecifically stains nucleic acids from all cell types. Thus it is important to distinguish fluorescencestained parasites from other cells and cellular debris containing nucleic acids [67].
20
Acridine orange (AD), has also been used as a direct-staining technique[59,68,69,70]. The simpler Kawamoto technique [56] employs an excitation filter mounted in the pathway of the transmitted light beam and can make use of strong sunlight as the exciting wavelength source [71]. However, acridine orange technique was not preferred over other methods because the lamp became excessively hot, the light was uncomfortable, and the staining was uneven [61].
In spite of some reservations about the use of aBC for malaria diagnosis, it remains a viable and rapid alternative to Giemsa staining as a screening test because of its higher sensitivity in order to eliminate negative slides that generally predominate in surveys. Microscopy would then be used for final confirmation and species identification of aBC positive samples [51,58].
1.7.2 Malaria Rapid Diagnostic Devices (MRDD)
Malaria rapid diagnostic devices (MRDD) have been recently introduced as an alternative to the microscopy for diagnosing malaria [45]. These devices use an immunochromatographic dipstick to detect Plasmodium-specific antigens in the patients' blood samples. The ParaSight®-F, the first malaria rapid diagnostic device (MRDD) introduced in the early 1990s, was capable of detecting Plasmodium falciparum only [72]. Significant improvements have been made since the introduction of the first device and it is hoped that MRDDs would offer accurate, reliable, rapid, cheap and easily available
alternatives to the traditional methods of malaria diagnosis [39]. Currently,
several MRDDs have been developed and available in the market and many of them
21
have capability to detect all four species and differentiate Plasmodium falciparum from non-falciparum. The major antigens targeted by these devices includes: Histidine Rich
protein II (HRP-II) [72], parasite lactate Dehydrogenase (plDH) [73], Plasmodium aldolase [38] and an unidentified antigen which is claimed to be specific for Plasmodium vivax [74].
Histidine rich protein 2 (HRP-2), a water soluble protein (antigen) produced by asexual blood stage and young gametocytes of Plasmodium falciparum [72], is the target of several rapid tests, such as the ParaSighFM F tests (Beckton Dickinson, Franklin lakes, NJ), the ICT malaria Pftest (ICT diagnostics, Sydney), and the Determine™ test (Abbot laboratories, Tokyo). HRP-2 is localized in the RBC cytoplasm and released into plasma in high amounts from the RBCs infected with P.falciparum, and may persist in blood up to 28 days after cure of infection and may not be a suitable target for devices that intend to monitor therapeutic response [75]. Moreover, Devices that target this antigen, could detect only Plasmodium falciparum. People in endemic area with low persistence parasitemia may be positive but the clinical value of such result is doubtful whereas some individual may have a gene deletion for the production of HRP-2 and will never give HRP-2 positive results [38].
Parasite lactate Dehydrogenase (plDH), an enzyme in glycolytic pathway is produced by the viable blood stage parasites of all four human Plasmodium species and each species produces a distinct isomer [38]. Thus, MRDDs that targets this enzyme could detect all four species, differentiate falciparum from non-falciparum and may be suitable for monitoring therapeutic response [37]. OptiMAL is a prototype of MRDD that targets this enzyme [76].
22
Plasmodium aldolase is also produced by all four species of Plasmodium and is an
enzyme in glycolytic pathway [38].
Monoclonal antibodies produced against
Plasmodium aldolase are pan-specific and have been used in combination with HRP-2
to detect P.falciparum as well as other Plasmodium species [38]. Previous studies with MRDDs based on this enzyme demonstrated the low sensitivity of detecting Plasmodium vivax, particularly at low parasitemia [38,77].
Performances of the several MRDDs have been reviewed [38,39,44,76]. These reviews of the earlier studies generally proved that the MRDDs have good sensitivity (>90%) and specificity (100%) for Plasmodium falciparum at parasite densities >100-500 parasites IIlI of blood but sensitivity decline rapidly at lower parasite densities and the performance
(sensitivity) for Plasmodium vivax is generally lower. A recent review by Murray et al [39] has shown that the overall sensitivity and specificity of OptiMal varies from 62% to 99% and 84% to 94%, respectively. Similarly, the overall sensitivity and specificity of ICT PflPvvaries from 63% to 84% and from 89-97%, respectively.
Performance characteristics of a few of the most recent studies using the prototypes MRDDs are summarized in the table 2. The recently introduced NoW® ICT P.f.lP. v. (Binax, Inc, Portland, ME) that detects P.falciparum HRP-2 and pan-Plasmodium aldolase was evaluated in symptomatic patients attending the malaria clinics in endemic areas in Thailand [76]. A similar product ParaSight f+v (Beckinson Dickinson) was also evaluated [74]. The overall sensitivity and specificity of the NoW® ICT P.fIP.v. and ParasSight f+v were excellent (>90%).
However, again as before the detection of
Plasmodium vivax was somewhat lower (sensitivity 87.3%). Similarly two recent studies
23
of the OptiMAL, one in malaria endemic areas in India [78] and the other in multiple hospitals in US [79]
were also very encouraging, the sensitivity approaching almost
100%. Although, the specificity was somewhat lower in field study in India [78], continuous improvement in the latest versions of the MRDDs is encouraging and MRDDs may become a good screening tool for malaria diagnosis.
Although the recent versions of the most MRDDs have good sensitivity with high parasitemia, their sensitivity is still low when parasitemia is low [80,81], they failed to detect gametocytes [76] and could not differentiate among various non-falciparum parasites. These antigen detection methods are fast and simple to perform and are good for use in clinical diagnosis. They are not yet reliable for conducting epidemiological studies, screening blood donors and monitoring drug effectiveness or development of resistance. Moreover, their high cost ($ 1.2 in developing countries) [50] is the main limiting factor for their wider use.
24
Device
d·
f
--
Location, Sample type & size
Gold Standard
NoW®ICT Malaria P.f/P.v. (Binax, Inc, Portland, ME
Thailand, Symptomatic, malaria clinic patients from areas, endemic N=246
microscopy
OptiMAL
symptomatic India, hospital and field clinic attendants in endemic areas N=86 (Hospital) N=75 (Field) hospitals, 6 US, symptomatic returned travelers N=216
Microscopy
Thailand & Peru , symptomatic malaria clinics patients N=4,873
Microscopy (+ve=2.051, 42.1%)
OptiMAL (DiaMed)
ParaSight f+v prototype FV99-2 (BD)
Microscopy
fMRDD
Sensitivity
Specificity (%)
Comments
Ref.
(%) Overall=94.3% Pf=100% Pv=87.3% But only 40% when parasitemia was
20parasites/1l1 detected
[76]
Hospital Pv=98.2 Pf=100 Field: Overall=67
Poor specificity in the field study Relation with level of parasitemia?
[78]
100
Parasitemia not reported, generally >0.01 % (50/1l1) Discrepant results were confirmed by PCR, were in agreement with OotiMal Three prototypes were used, specificity varied greatly among the prototypes, Performance of FV99-2 was the best
[79]
pf=93% pV=83%
100% when parasitemia was >500/ul Pf= P. falciparum, Pv= P. Vivax, US=United States, PCR=Polymerase Chain Reaction, N=number
25
[74]
1.7.2
Molecular methods
Molecular methods have gathered pace over the second half of the last decade and since then all major scientific disciplines have found molecular techniques increasingly useful in research, diagnosis, prognosis, and clinical management. The molecular revolution began in 1953 when Watson and Crick proposed the structure of doublestranded deoxyribonucleic acid (DNA) [82]. Subsequently, purified restriction enzymes were developed, which allows further manipulation of DNA [83]. Development of Southern blotting by E.M. Southern allows detection of sequences of DNA fragments separated by an agarose gel using radioactive probes[84].
In 1987 a unique DNA
amplification method called the polymerase chain reaction (PCR) was developed, which was followed by the development of reverse transcriptase PCR (RT-PCR) for the amplification of RNA. PCR has been further developed in recent years to provide realtime fluorescent methods using Taqman and Molecular Beacon chemistries and quantification of parasites by this approach has become possible [85,86,87,88].
Molecular techniques have been used in diagnosing Plasmodium for several years. Initial nucleic acid based probes were focused on P.falciparum targeting the 21-bprepetitive sequences found throughout the P. falciparum genome [89]. Although, these probes were very specific [90], their sensitivity was lower than that of microscopy particularly with the patients having low parasitemia [91]. Subsequently, nucleic acid probes targeting species-specific rRNA molecules and labeled with radioisotope were used to diagnose the malaria and a sensitivity detecting 10-50 parasites/J,.t1 was achieved [92]. Although, there were efforts to increase the sensitivity of nucleic acid based hybridization techniques [93], a number of candidate DNA probes were found to be less
26
useful than previously thought [91] and efforts were switched to PCR once it became available [94].
Snounou et al in 1993 used species-specific oligonucleotide primers
targeted to 18S Small Subunit of ribosomal RNA (SSU RRNA) gene of all human Plasmodium species for PCR and detected a large no. of mixed infections [95]. The
multicopy 18S small subunit rRNA genes of Plasmodium species that infect human have been found to be highly stable and conserved, do not cross react to human or other human pathogens' DNA or RNA [25,95]. In recent years several polymerase chain reaction (PCR) assays have been developed and evaluated for diagnosing malaria most often based on genus or species specific sequences of the parasites' 18S small subunit rRNA gene (95,96,97) and generally PCR particularly the nested PCR was found much more sensitive and specific than those of the other diagnostic methods when compared either with microscopy or rapid diagnostic tests [94,96,97,98] . Moreover, multiplex PCR, where more than one target sequences can be amplified by including more than one primer sets in a single reaction and thus potentially could save time and effort has been shown to be a valuable tool in infectious disease diagnosis [99] could also theoretically be used to diagnose malaria. Moreover, PCR can be a valuable tool for epidemiological diagnosis and if used for screening blood donors can dramatically reduce or eliminate transfusion transmitted malaria. It can also reliably be used for monitoring drug effectiveness or emergence of drug resistance in parasite populations.
27
f the Select"- - - - - - --- t peR stud·-- -Malaria prevalence (No. & % of cases rJositive) Country, sample population, Other & size(N) PCR nested Microscopy Non-endemic, Semi Italy, =61 (50%) symptomatic returned travelers PCR =62 Real Time PCR from endemic N=122 =60
Table3. S
Canada, travelers N=259
Symptomatic
return
France, symptomatic patients, N=183
Real PCR=208 (80.31%)
Nest PCR=209 (80.69%)
cPCR= 60 (32.78%)
Microscopy =48 (26.22%)
mcrt-PCR=60 (32.78%) Singapore, Symptomatic patients from malaria reference center & laboratory culture samples, N=153 Iran, symptomatic treatment seeking individuals from endemic areas N=120 Malaysia, endemic area, field survey N=129 Spain, non-endemic, symptomatic returned travelers, N=168
Real PCR=127 (83.01%)
Microscopy =125 (81.70%)
116 (96.67%)
107 (89.17%)
46 (35.6%)
37 (28.7%)
89 (53)
70 (41.7)
Comments
TaqMan based Real Time PCR with detection limit of 0.7 (Pf), 4{Pv) & 1.5{Po) per /-ll of blood., results within 2 hours 2 Real Time PCR -ve samples were P.malariae not designed to detect by this PCR Real time PCR detected 14.7% (9/61) misdiaonosed samples RealArt malaria PCR assay (Artus Biotech), sensitivity= 99.5% & specificity=100%, also detected 2 P. ovale samples with a variant 18S rRNA , assay time -
eo
L..
0.
«
~
LL
C")
C")
L..
~
C")
C")
0I
0I
C
:::J J
:::J J
eo 0
l-
127 174 280 310 136 3182
Performed
aBC
39
17
13
6
3
Positive
40
6
1
1
5
6
21
6
153
.!!l 45 l: .!!! 40 : . 35 ~ 30 ~ 25 ~ 20 o 15 lD o 10 0 5 ci z 0
-
-A-
Positive aBC
-
Jun- Jul-02 Aug02 02
Sep02
Oet02
Nov02
Dec02
Jan03
Feb03
Mar03
Apr03
Months & Years Fig. 3. Monthly distribution of aBC Positive cases (June 18,2002 to July 17,2003)
41
May03
Jun- Jul-03 03
4.1 SAMPLE PROFILES
Eighty-five samples comprising of both microscopy and aBC positive (n=17), only aBC positive (n=31) and both microscopy and aBC negative (n=37) were analyzed by nested PCR. The mean age of the sample population was 26.64 years (SO=16.85 years) and ages ranged from 1 day to 71 years. Among the sample population (N=85), males (n=50, 58.82%) slightly outnumbered females (n=35, 41.18%). Although females were slightly younger (22.97 years, SO=14.12) than males (29.22 years, SO= 18.22) (Table 5) the age difference was not statistically significant (F = 1.66 with 83 (49, 34) O.F., P = 0.12).
Table 5. Sample population with their mean age (SO) a nd sex distribution
Sex
Number (%)
Mean Age (SO) (In Years)
Male
50 (58.82%)
29.22 (18.22)
Female
35 (41.18%)
22.97 (14.12)
Total
85 (100.00%)
26.64 (16.85)
4.2 aBC malaria test results
Out of 85 samples, 48 (56.47%) were positive by malaria aBC test. Among them 35 (72.98%) were male whereas only 13 (21.02%) were female and more importantly, 70% of male (35 out of 50) and only 37.14% of female (13 out of 35) were positive by aBC malaria test. A majority (n=29, 60.42%) of aBC positive cases had lower parasitemia (1 + or 2+) and proportionately more female (10, 76.92%) had the lower parasitemia (1+ or 2+) than that of the male (26, 54.29%) (Table 6), however, the difference was not statistically significant (X2 =2.03; df=1 ,1; p=0.115).
42
Table 6. Breakdown of aBC positive cases (N=48) according to sex and relative . paras I'temla. No. of cases according to level of parasitemia 8ex
1+
2+
3+
4+
Male (n=35)
15 (42.86%)
4 (11.43%)
11 (31.43%)
5 (14.29%)
Female (n=13)
8 (61.54%)
2 (15.38%)
3 (23.08%)
0(0.00%)
Total (N=48)
23 (47.92%)
6 (12.50%)
14(29.17%)
5 (10.42%)
The mean age of aBC positive male (29.22 years, 80=18.22) & female (22.97 years, 80=14.12) was not significantly different from that of the sample population (table-2).
4.3 MICROSCOPY RESULTS
All aBC positive (153) and an equal number of randomly selected aBC negative samples were examined microscopically by a Jenner-Giemsa stained thin smear. Among the 306 samples thus examined only 17 were positive for malaria. None of the
aBC negative samples was positive by microscopy and all microscopy positive samples were also positive by aBC. Among the microscopy positive samples, 9 (53%) were P. falciparum, 7 (41%) were P. vivaxand one (6%) was a mixed sample containing both P. falciparum and P. vivax (fig. 4). The proportion of P. falciparum positive cases (59%)
was substantially higher than national average (7%) (fig. 5) but was similar to the overall falciparum rate in the Eastern region [11].
43
Mixed infection
P. v/vax Fig.-4. Proportion of different speci es of Pia smodi urn c ausin g malaria in patients attending to BPKIHS (June 2002-July 2003)
F ig.-5. Proportion of fa Icipa rum an d non-falcip arum malaria in Nepal (2001)
The high parasitemia (3+ or 4+) by aBC was a strong predictor of a microscopy positive result. Among the 19 patients with high aBC parasitemia (3+ or 4+) 16 (84.21 %) were positive by microcopy whereas only 1 patient (3.45%) among the 27 low parasitemia (1 + or 2+) group was positive by the microscopy. Interestingly, 80% (4 out of 5) of the patients with very high (4+) aBC parasitemia had either P. falciparum or mixed infection and only one (20%) of the same group was negative by the microscopy. On the other hand patients with moderately high parasitemia (aBC 3+) (n=14) were either equally infected with
P.falciparum (n=6, 42.86%) or P. vivax (n=6, 42.86%) and were rarely
negative by microscopy (n=2, 14.29%). Interestingly, only one patient with low parasitemia (1+ & 2+) was positive (3.45%) (P. vivax) by microscopy. 44
4.5 PCR RESULTS 4.5.1 Plasmodium Genus specific PCR Among the selected samples (N=85), 24 (28.24%) were positive by Plasmodium genus specific PCR targeting the 188 88U rRNA gene (Fig. 6, positive samples contain 240 bp band) of which only 19 (79.17%) & 14 (58.33%) were positive by aBC & microscopy respectively. However, 3 microscopy positive and 29 aBC positive cases were negative by the PCR but PCR was positive in an additional 5 aBC negative and 10 microscopy negative cases. In other words, PCR picked up seven more cases than microscopy but a substantial number of aBC positive cases were negative by PCR.
The higher parasitemia (3+ or 4+) on aBC test was also a strong predictor of the PCR positive result. Among the 19 patients with high aBC parasitemia (3+ or 4+)
14
(73.68%) were positive by the PCR whereas a large majority (88.46%) of patients with low parasitemia (1 + or 2+) were negative by the PCR. Interestingly, the PCR positive rate (17.51 %) among the lower parasitemic patients was not significantly higher than that of the aBC negative patients (13.51 %).
45
240 bp-
240 bp_ L
NC PC 15 16 17 18 19
20 21
22 23 24
25
26
27 28 29
240bp L
NC
PC
30
31
32
33
34
35
36
37 38 39
40
41
42 43 44 45
54
55
56
240bp L
NC
PC
46
47
4S
49
50
51
52
53
57
58
59
240 bpL
NC PC 60
61
62
63 64 65 66
67
68 69
70
71
82
83
72
73
240 bp-+ L
NC PC 74
75 76 77
78 79
80
81
84
85
Fig. 6. Agarose Gel (2%) electrophoresis of Nest 2 PCR product using Plasmodium Genus specific primers, L=100bp ladder, PC=positive control, NC= Negative control. PC and samples positive for Plasmodium was detected by 240bp (--+).
46
4.5.2 Plasmodium Species Specific PCR Out of 24 Genus specific PCR positive samples, 23 (95.83%) were positive by the species specific PCR (fig.-7 and 8). Among 24 PCR positive cases, 12 (50.00%) were P. falciparum, 7 ( 29.16%) were P. vivax, 4 (25.00%) were mixed infection with P. falciparum, and P. vivax.
One sample (4.17%) was positive for malaria by genus
specific PCR but was negative for all species specific PCR in spite of repeated attempts. Three samples that were reported as P.falciparum positive by microscopy were found to be positive for both P.falciparum and P. vivax by PCR. Similarly, one sample reported as P. vivax by microscopy was also found be infected by both P.falciparum and P. vivax with the PCR. However, two P.falciparum positive samples and one P.falciparum and P. vivax mixed infection samples that were detected by microscopy were found to be negative by the PCR (Table 7).
.
Tabl e 7 Companson 0 f t he resu ts 0 f t he microscopy an d species-speciTIC PCR Species specific PCR P. falciparum >-
P. vivax
Mixed (P.f.+P.v.)
Total (microscopy) Negative
P.falciparum
4
0
3
2
9
P. vivax
3
3
1
0
7
Mixed
0
0
0
1
1
Negative
5
4
0
59
68
12
7
4
62
85
Q.
0 0
en
0 .... 0
~
Total (PCR)
47
205
205 bp-t L PC NC 1 2 3
205
bp~
7
bp~
L
15 18 29
PC NC 34 35 36 37 41 46 47
05bp_
L PC NC 48 50 51 52 53 L PC NC 54 55 56 60 62 Fig-7. Photograph of the 2% Agarose gel electrophoresis stained with 5% Ethidium bromide of the Nest 2 peR products amplified with the Plasmodium falciparum specific primers (rFAL1 & rFAL2). Positive samples gave 205bp band (--+).
48
120bp--1
L
NC
PC
2
3
7
15
18
29
120bp-
L
NC
PC
54
55
56
60
62
Fig-S. Photograph of the 2% Agarose gel electrophoresis stained with 5% Ethidium bromide of the Nest 2 peR products amplified with the Plasmodium vivax specific primers (rVIV1 & rVIV2). Positive samples gave 120bp band (~).
49
4.6 SUMMARY OF ALL DIAGNOSTIC TEST RESULTS From the result, it was apparent that the aBC detected the maximum number of malaria positive samples followed by the PCR and microscopy (Table 8)
Table 8. Number and percentage of the positive and negative cases as determined b»y th eth i· ree ma ana d·lagnosrIC t est s. Positive
Negative
Total (n=85)
aBC
48 (56.47%)
37 (43.53%)
85 (100.00%)
Microscopy
17 (20.00%)
68 (80.00%)
85 (100.00%)
PCR
24 (28.24%)
61 (71.76%)
85 (100.00%)
The study revealed that over three-fourths of the malaria positive cases were males (table 9) and the maximum number of malaria positive cases by all tests were found in the age group of 15-30 years (table 10). However, the proportion of cases in the different age groups was similar to the proportion of sample population and a further population based study is needed to see whether the malaria prevalence in Nepal varies with age.
Table 9. Sex distribution of malaria positive cases according to different malaria diagnostic tests. No. & (%) of positive cases Tests
Male
Female
aBC
35 (72.92%)
13 (27.08%)
Microscopy
14 (82.35%)
3 (17.65%)
PCR
17 (80.83%
7 (29.16%)
50
Tabl e 10" A~ge 0"ISt"b " 0 fth e sampi e popuIaf Ion an d positive cases rI ut.on Age Group
Total sample
aBC Positive
(Years)
No. (%)
No. (%)
Microscopy Positive
PCR Positive No. (%)
No. (%) 0-15
18 (21.18%)
10 (20.83%)
4 (23.53%)
6 (25.00%)
15-30
39 (45.88%)
23 (47.92%)
6 (35.29%)
10(41.67%)
30-45
16 (18.82%)
7 (14.58%)
4 (23.53%)
6 (25.00%)
45-60
8(9.41%)
6 (12.50%)
3 (17.65%)
2 (8.33%)
60-75
4 (4.70%)
2(4.17%)
0(0.00%)
0(0.00%)
All Ages
85 (100.00%)
48 (100.00%)
17 (100.00%)
24 (100.00%)
Although the Microscopy positive cases were slightly older (mean age=28.29 years, 80=18.26) than the aBC (mean age=25.98 years, 80=16.49), and the PCR (mean age=25.00 years, 15.49) positive cases (Table 11), the age difference was not statistically significant (p >0.05).
Table 11" Mean age (SO) of the positive and negative sample population Mean Aae(SD) in years Positive Negative p-value aBC 25.98 26.77 0.974 (16.49) (16.37) (F=1.01), OF (47,36) Microscopy 28.29 25.80 0.426 (18.26) (15.91) (F=1.32), OF (16,67) PCR 25.00 26.88 0.686 (15.49) (16.79) (F=1.17), OF (60,23)
51
4.7 SENSITIVITY AND SPECIFICITY Given the lack of universally available accepted gold standard for diagnosing malaria, the sensitivity and specificity of the aBC and PCR were first calculated against the microscopy, which has been often used as gold standard [46]. However, microscopy has been widely recognized as a less sensitive test and the evaluation of a new diagnostic tool only based on the microscopy would not reflect the true sensitivity and specificity of the tool to be evaluated. Since PCR is reported to be very sensitive and specific test for malaria diagnosis, sensitivity and specificity of microscopy and aBC were also calculated against the PCR as gold standard.
It is apparent from the result (table 12) that the aBC was able to pick up all microscopy positive cases and thus was 100% sensitive. Moreover, none of the aBC negative cases were positive by microscopy thus aBC had 100% negative predictive value. However, a large numbers of aBC positive cases were negative by the microscopy suggesting that the aBC was much less specific tests (54.41 %) for diagnosing malaria and its positive predictive value was just over 35 percent (table 12).
Table 12. Performance of QBC when compared with microscopy as a gold standard Microscopy
Total
Positive
Negative
Positive
17
31
48
Negative
0
37
37
17
68
85
()
lD
a
Total
...
SensitIVIty of aBC= 17/17 = 100.00 % Specificity of aBC = 37/68= 54.41 % Positive predictive value= 17/48= 35.41 % Negative predictive value= 37/37= 100.00%
52
On the other hand, PCR appeared much more specific than the aBC (85.25%) and had a good negative predictive vale (95.08%). Moreover, the specificity of the PCR (85.29%) was higher than the aBC and thus had higher positive predictive value (58.33%) (table 13).
Table 13. Performance of peR as compared with microscopy (gold standard) Microscopy
a:
0
a..
Total
Total
Positive
Negative
Positive
14
10
24
Negative
3
58
61
17
68
85
Sensitivity of PCR= 14/17 = 82.35% Specificity of PCR= 58/68= 85.29% Positive predictive value= 14/24= 58.33% Negative predictive value= 58/61 = 95.08%
53
Assuming that both microscopy and PCR are very specific tests, any sample, which was found positive by either of the test, was likely to be a true positive sample. Thus, sensitivity and specificity of aBC were also calculated considering all positive cases by either microscopy or PCR as the true positive cases. aBC detected 22 of the 27 true positive cases as determined by the combined performance of the PCR and microscopy and thus had a sensitivity of 81.48% and it also had a fairly good negative predictive value (86.49%). However, a large number of aBC positive cases (26) were negative by both
microscopy and PCR and thus aBC had a lower specificity (55.17%) and low
positive predictive value (45.83%) (table 14).
Table 14. Performance of QBC as compared with the combined performance of both microscopy & PCR Total
Microscopy & PCR Positive*
Negative**
Positive
22
26
48
Negative
5
32
37
27
58
85
()
m
a
Total
*Total cases positive either by microscopy or PCR or both are considered true positive. **Only cases negative by both (PCR & Microscopy) tests were considered true negative. Sensitivity of aBC= 22/27 = 81.48% Specificity of aBC= 32/58= 55.17% Positive predictive value= 22/48= 45.83% Negative predictive value= 32/37= 86.49%
54
CHAPTER 5 DISCUSSION AND CONCLUSIONS
A rapid, accurate, sensitive and affordable test for diagnosing malaria is important for effective clinical management. However, diagnosis of malaria is a complex task and there is no single tool that is suitable for all settings.
The results of this study indicated that malaria is seasonal in Nepal and maximum cases were seen during the summer (June through September). This finding is consistent with the national trend [11] and is probably due to favorable condition for mosquito vector breeding during the warm humid summer associated with the high monsoon.
Apparently, malaria was much more common among males and levels of parasitemia as determined by the aBC were higher among males than females.
Although these
findings need to be confirmed by the larger community based studies, if true these findings are in contrast to the malaria in Africa where female are considered more vulnerable to this disease [7]. Probably, the feminine gender role in Nepal may limit the exposure of females to mosquito bites and relatively protect them from malaria. However, the reason for low parasitemia among the malaria positive females when compared with the malaria positive males is not known and may reflect differences in disease susceptibility or reporting differences.
55
Unlike the highly endemic African countries where young children are affected the most [7], maximum numbers of malaria positive cases were found in the age group of 15-30 years in this study population. Whether this finding is because this particular age group is more susceptible to malaria or more frequently exposed to mosquito bites is unclear from this study. A further field based study is needed to reach a reliable conclusion. However, malaria was seen at all ages in Nepal and immunity, if it exists, is not adequate to protect any particular age group from malaria. This is probably because of low and unstable epidemics of malaria in Nepal.
A high proportion of malaria cases at BPKIHS was found to have P. falciparum or mixed infections. Although, the exact reasons why the P. falciparum cases were substantially higher at BPKIHS than the national average are not clear, possible explanations could include: a) more sick patients might have visited hospital who were likely to have P. falciparum infection whereas mildly symptomatic P. vivax cases might have
treated locally and thus did not visit the University Teaching Hospital. b) Moreover, a substantial proportion of these patients may have been Bhutanese refugees taking shelter the Eastern Nepal who have a high P. falciparum rate [11] . Whatever may be the reason, it is an alarming signal and every clinician involved in treating these patients should be made aware of this fact.
A large numbers of aBC positive cases were negative by both microscopy and PCR. Although superior sensitivity of aBC could not be ruled out in the absence of acceptable
56
confirming methods, it is most likely that some aBC tests were falsely positive. In fact, a high false positivity rate of the aBC test has been suggested as the reason for its reported high sensitivity [61] and the agreement between two examiners in interpretating aBC results was often low [61]. The other possible reason for very high false positive aBC malaria test result at BPKIHS could be due to its recent introduction to BPKIHS, with the consequence that laboratory technician may not yet be well versed in its interpretations.
As expected, PCR has detected more cases than the microscopy technique and more importantly many more mixed infections were identified by the PCR in this study. However, the nested PCR was a very labor intensive procedure and sample contamination was a problem in initial experiment requiring the repetition of the PCR assays [63]. Since, the existing microscopy has low sensitivity, and the aBC has very high false positive rate, PCR would be the most reliable alternative for clinical and epidemiological diagnosis of malaria in Nepal. However, its routine use in the clinical laboratory in Nepal in the present form does not appear to be feasible because of its high cost, long processing time and requirement to run several PCR cycles for each samples. Further automation may make it more rapid and reliable. However, it could be a valuable tool for quality control as it can be used in selective samples to verify the microscopy or aBC test results.
Recently, a few ready to go malaria real time PCR Kits have become available in the market and the first few studies showed very promising results [85,87,88]. They are sensitive, fast and easier to use, can quantify the Plasmodium and can differentiate
57
among at least the three of the four human Plasmodium species [88]. The trend is such that, it is not unrealistic to expect availability of commercial kits capable of differentiating all four Plasmodium species in the near future. Furthermore, automated DNA extraction method could be incorporated with Real time PCR and thus would potentially solve the problem of malaria diagnosis in the non-endemic developed world.
However, even the availability of a very sensitive, specific, rapid and affordable diagnostic tool in detecting malaria parasites from human blood would not completely solve the diagnostic dilemma of the malaria in the endemic world. A great majority of people in the high endemic areas always harbor Plasmodium in their blood and detecting Plasmodium does not necessarily prove that the cause of illness is malaria. It was interesting to note in a recent study that there was a high prevalence of parasitemia even among the asymptomatic healthy volunteers from the endemic areas (28.5% by hemi-nested PCR) whereas the parasitemia positive rate among the symptomatic group was just over 50% [51]. Therefore, in a setting where Plasmodium parasites could not be detected from the blood of a symptomatic patient, malaria could safely be excluded as a potential cause of the symptoms and would encourage the clinicians to look for alternative causes. However, detecting Plasmodium parasites in symptomatic patients, although it is an additional supporting evidence for malaria diagnosis, it should not preclude the clinicians for searching for alternative causes of the symptoms. Moreover, PCR may be too sensitive in detecting Plasmodium that is present at non-pathogenic level and may also pose a problem in monitoring vaccine efficacy.
58
APPENDIX PROFILE AND RESULTS SUMMARY OF THE SAMPLE POPULATION SAMPLE 10
BP1 BP2 BP3 BP4 BPS BP6 BP7 BP8 BP9 BP10 BP 11 BP12 BP13 BP 14 BP1S BP16 BP17 BP 18 BP 19 BP20 BP21 BP22 BP23 BP24 BP2S BP26
AGE
3 13 3 20 8 43 25 68 30 69 10 24 18 55 59 15 16 29 22 18 34 48 22 14 22 29
SEX M F M F F M F M F M F M F F M F F M F F M M M M M M
Plasmodium Positive Positive Positive Negative Negative Positive Positive Positive Positive Positive Positive Positive Positive Positive Positive Positive Positive Positive Positive Negative Negative Negative Positive Positive Positive Positive
QBC Quantitative
4 3 3 0 0 1 1 1 2 1 1 1 1 1 3 1 1 4 2 0 0 0 4 1 3 1
MICROSCOPY Species Plasmodium falciparum Positive falciparum Positive falciparum Positive negative Negative negative Negative negative Negative negative Negative negative Negative negative Negative negative Negative negative Negative negative Negative negative Negative negative Negative falciparum Positive negative Negative negative Negative falciparum Positive negative Negative negative Negative negative Negative negative Negative mixed Positive negative Negative vivax Positive negative Negative
59
Plasmodium Positive Positive Positive Negative Negative Negative Positive Negative Negative Negative Negative Negative Negative Negative Positive Negative Negative Positive Negative Negative Negative Negative Negative Negative Negative Negative
PCR Species falciparum falciparum & vivax falciparum & vivax Negative Negative Negative falciparum Negative Negative Negative Negative Negative Negative Negative falciparum Negative Negative falciparum Negative Negative Negative Negative Negative Negative Negative Negative
SAMPLE 10
BP27 BP28 BP29 BP30 BP31 BP32 BP33 BP34 BP35 BP36 BP37 BP38 BP39 BP40 BP41 BP42 BP43 BP44 BP45 BP46 BP47 BP48 BP49 BP50 BP51 BP52 BP53 BP54 BP55 BP56
AGE
21 18 32 19 52 9
SEX F
M M M M M
4
F
37 19
M
6
57 39 55 21 18 18 59 25 20 28 38 22 11 30 17 25 35
F F
M M M M M M M F F F
M M M M M F
M
44
F
25
M M
9
QBC Quantitative Plasmodium 0 Negative 1 Positive Positive 3 2 Positive 1 Positive Positive 3 1 Positive Negative 0 Negative 0 Positive 3 1 Positive 1 Positive Negative 0 4 Positive Positive 3 1 Positive Positive 3 Negative 0 Negative 0 1 Positive Positive 3 Positive 3 Positive 1 Positive 3 Positive 3 Negative 0 NeQative 0 Positive 3 2 Positive Positive 2
MICROSCOPY Plasmodium Sl:ecies negative Negative negative Negative Positive vivax Negative negative Negative negative Negative negative Negative negative Negative negative Negative negative Positive vivax Positive vivax Negative negative negative Negative Negative negative Negative negative Negative negative Positive falciparum Negative negative negative Negative negative Negative Positive vivax Positive vivax Negative neQative Positive vivax Positive falciparum Negative negative Negative negative Positive falciparum Negative negative Negative negative
60
PCR Plasmodium Negative Negative Positive Negative Negative Negative Negative Positive Positive Positive Positive Negative Negative Negative Positive Negative Negative Negative Negative Positive Positive Positive Negative Positive Positive Positive Positive Positive Positive Positive
Species Negative Negative falciparum Negative Negative Negative Negative falciparum falciparum falciparum falciparum Negative Negative Negative vivax Negative Negative NeQative Negative vivax vivax vivax Negative vivax falciparum vivax vivax falciparum & vivax vivax falciparum
SAMPLE 10 BP57 BP58 BP59 BP60 BP61 BP62 BP63 BP64 BP65 BP66 BP67 BP68 BP69 BP70 BP71 BP72 BP73 BP74 BP75 BP76 BP77 BP78 BP79 BP80 BP81 BP82 BP83 BP84 BP85
AGE
19 55 33 1 26 25 10 1 24 21 17 43 4 25 35 25 14 18 34 71 35 28 8 35 16 17 4 66 25
SEX
M M M M M M F F F M F M M F F F F M F F F F F F M M M M M
Plasmodium Positive Positive Neaative Negative Positive Positive Negative Negative Neaative NeQative Neaative Negative Negative NeQative Negative Neaative Negative Neaative Negative Neaative NeQative Negative NeQative Negative Positive Neaative Negative Negative Negative
QBC Quantitative
1 1 0 0 1 4 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 2 0 0 0 0
MICROSCOPY Plasmodium Species Neoative neoative negative Neoative neaative Neaative Negative neaative NeQative neQative falciparum Positive negative Negative Negative negative Neaative neaative negative Negative Neaative neaative negative Negative neaative Neaative NeQative neQative Negative neaative neoative Neaative Negative neaative NeQative neQative Negative negative Neaative neaative neQative Neoative Negative negative NeQative neQative Negative negative Negative negative Neaative neaative Negative negative Negative negative NeQative neQative
61
Plasmodium Neoative Negative Neoative Positive NeQative Positive Negative Negative Neoative NeQative Neaative Negative Negative Neaative Negative Neaative Negative NeQative NeQative Negative Neaative NeQative Neaative NeQative Negative Negative Neaative NeQative Neaative
PCR Species Neoative Negative Neaative falciparum Neoative Negative NeQative NeQative Negative Neoative Negative NeQative Negative Neaative Negative Neaative Neoative Neaative NeQative Negative Negative Neoative Negative Negative Negative Negative Negative Neoative Negative
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