Development of a highly sensitive genus-specific qRT-PCR assay for detection and 1

quantitation of Plasmodium by amplifying RNA and DNA of the 18S rRNA genes 2

3

Edwin Kamau1*

, LaDonna S. Tolbert2, Luke Kortepeter

1, Michael Pratt

1, Nancy Nyakoe

3, 4

Linda Muringo3, Bernard Ogutu

3, John N. Waitumbi

3 and Christian F. Ockenhouse

1. 5

6

1Division of Malaria Vaccine Development, United States Military Malaria Vaccine Program, 7

Walter Reed Army Institute of Research, Silver Spring, Maryland, United States of America. 8

2Division of Entomology, Molecular Diagnostics, Walter Reed Army Institute of Research, 9

Silver Spring, Maryland, United States of America. 10

3Walter Reed Project, Kenya Medical Research Institute, Kisumu, Kenya 11

12

Running title: Total Nucleic Acids PCR Assay for Malaria Diagnosis 13

Key words: Plasmodium, Malaria Diagnosis, real-time PCR, Microscopy, Malaria Elimination 14

Abbreviations: Real-time PCR (qPCR), quantitative reverse transcriptase real-time PCR (qRT-15

PCR), Limit of Detection (LOD), nucleic acids (NA), Malaria Research and Reference Reagent 16

Resource Center (MR4), American Type Culture Collection (ATCC), reverse transcriptase (RT) 17

*Corresponding author 18

Division of Malaria Vaccine Development, Center for Molecular Diagnostics and Genomic 19

Studies, Walter Reed Army Institute of Research, 503 Robert Grant Ave, Silver Spring, 20

Maryland, United States of America. 21

22

Telephone number: (301) 319 7572 23

E-mail: Edwin.kamau@us.army.mil 24

Copyright © 2011, American Society for Microbiology and/or the Listed Authors/Institutions. All Rights Reserved.J. Clin. Microbiol. doi:10.1128/JCM.00276-11 JCM Accepts, published online ahead of print on 8 June 2011

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Abstract 25

26

A highly sensitive genus-specific quantitative reverse transcriptase real-time PCR (qRT-PCR) 27

assay for detection of Plasmodium has been developed. The assay amplifies total nucleic acids 28

(RNA and DNA) of the 18S rRNA genes with a limit of detection of 0.002 parasites/µL using 29

cultured synchronized ring stage 3D7 parasites. Parasite densities as low as 0.000362 30

parasites/µL were detected when analyzing clinical samples. Analysis of clinical samples 31

showed that detection of 18S rRNA genes from total nucleic acids increased the analytical 32

sensitivity of the assay by more than a log-fold compared to DNA only. When clinical samples 33

with no parasites present by microscopy were analyzed by qRT-PCR, 90% (117 of 130) were 34

positive for presence of Plasmodium nucleic acids. Quantification of clinical samples by qRT-35

PCR using total nucleic acid versus DNA was compared to microscopy. There was a 36

significantly greater correlation of parasite density to microscopy when DNA alone was used 37

compared to total nucleic acid. We conclude that analysis of total nucleic acids by qRT-PCR is a 38

suitable assay in detection of low parasite levels in patients with early-stage malaria and/or 39

submicroscopic infections and could greatly benefit malaria diagnosis, intervention trials, 40

malaria control and elimination efforts. 41

42

43

44

45

46

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Introduction 48

49

Malaria remains one of the most devastating infectious diseases in the world. After many years 50

of neglect, various philanthropies such as Roll Back Malaria (http://www.rbm.who.int) and 51

Gates Foundation have now committed to its eventual eradication (7). The success of these new 52

initiatives hinges in part with the use of effective diagnostic and surveillance methods (11). 53

However, despite the revolutionary gains from molecular approaches in diagnosis of malaria, 54

microscopy remains the gold standard for malaria diagnosis, clinical trials efficacy evaluation 55

and epidemiological surveys not withstanding it shortcomings (15, 16). 56

In expert hands, microscopy has a detection limit of 10 to 50 parasites/µL (9, 12), but the 57

average microscopist has a detection limit of about 100 parasites/µL, thereby limiting the use of 58

microscopy in cases of low parasite burden (2). Studies have shown that even at submicroscopic 59

infections, mosquitoes do get infected and can potentially transmit malaria (13, 19). Therefore, 60

as we move to the era of malaria control and elimination, highly sensitive methods with high 61

throughput capabilities will be critical in parasite detection and surveillance. Such methods will 62

be important in quantifying the extent of submicroscopic infections and giving a better insight 63

into the dynamics of malaria transmission. 64

Molecular techniques for detection of specific Plasmodium nucleic-acid sequence have 65

enabled measurement of infections that are a log-fold lower than microscopy or antigen detection 66

tests (18, 20). For example, polymerase chain reaction (PCR) assays allow explicit identification 67

of malarial species and can be easily adapted for high throughput application. Real-time PCR 68

(qPCR) especially has improved the application of PCR because the assay is fast, has very low 69

risk of contamination, is highly sensitive, specific and quantitative. These qualities are not only 70

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ideal for diagnosis and clinical trials efficacy evaluation, but will increasingly become needed for 71

use in epidemiological surveys and to evaluate the success of malaria control and elimination 72

campaigns. Superiority of PCR as a diagnostic tool for detection of Plasmodium over 73

microscopy has been extensively described. For example in a recent systematic review of 74

surveys of endemic populations in which P. falciparum prevalence was measured by both 75

microscopy and PCR-based technique, the prevalence of infection measured by microscopy was 76

shown to be on average, 50.8% of that measured by PCR (17). 77

Most PCR assays target DNA of the Plasmodium multicopy 18S ribosomal RNA (rRNA) 78

genes, which, due to their high copy number and mosaic of conserved and variable regions, 79

provide an ideal molecular target for malaria genus and species identification and quantification. 80

However, even slight genetic variation within 18S rRNA gene sequences of the same species has 81

been problematic. The Plasmodium genome lacks the long tandemly repeated arrays of rRNA 82

genes found in other eukaryotes. Instead, it contains several, single 18S-5.8S-28S rRNA units 83

distributed on different chromosomes with sequence encoded by rRNA gene in one unit differing 84

from the sequence of the corresponding rRNA in the other units (8). Additionally, the expression 85

of each rRNA unit is developmentally regulated, with different sets of rRNAs being expressed at 86

different stages of the parasite life cycle (10, 21). As such, it is important to consider all these 87

factors when designing a nucleic-acid sequence based assay which targets 18S genes. 88

In this study, we describe development of a highly sensitive genus-specific quantitative 89

reverse transcriptase real-time PCR (qRT-PCR) assay for detection of Plasmodium and shown 90

that amplification of total nucleic acids (RNA and DNA) of the 18S rRNA genes significantly 91

increases analytical sensitivity of the assay. We also compared quantification of clinical samples 92

in detection of Plasmodium by qRT-PCR and microscopy. 93

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Material and Methods 94

Samples. 95

Samples used in this study were obtained from a Phase IIb pediatric clinical trial conducted 96

between March 2005 and April 2006 at the KEMRI/Walter Reed Project, Kombewa Clinic in the 97

Kombewa Division of Kisumu District, Nyanza Province, Western Kenya. The study was 98

approved by the Ethical Review Committee of the Kenya Medical Research Institute, Nairobi, 99

Kenya and Walter Reed Army Institute of Research Institute Review Board, Maryland, United 100

States. 101

Details about this study have been described elsewhere (23). Briefly, EDTA-treated 102

blood samples were collected from study participants at enrolment (Day 0) and one month after 103

administration of the third and final vaccination. In addition, blood was also drawn during 104

unscheduled clinical visits from children who were sick and suspected to have malaria. For 105

assessment of malaria, a peripheral blood smear was obtained from subjects who presented to the 106

Walter Reed Project’s Kombewa Clinic with fever or a history of fever within 48 h, or an illness 107

that the attending doctor suspected might be due to malaria infection. After Giemsa staining, thin 108

and thick blood smear slides from each sample were independently examined by three expert 109

microscopists for detection of Plasmodium and count where applicable. All malaria 110

microscopists were fully trained and were required to pass a competency and proficiency test 111

prior to reading slides for the study. Detection of asexual parasitemia of >0 parasites/µL resulted 112

in the diagnosis and treatment for malaria. The parasite density presented in this study is the 113

average density obtained by the three independent (blinded from each other’s results) 114

microscopists. Two hundred µl of blood were aliquoted and stored in -20°C until required. 115

Genomic nucleic acid was extracted from whole blood using the QIAamp DNA Blood Mini Kit 116

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(Qiagen, Valenica, CA) as recommended by the manufacturer. Extracted nucleic acids (NA) 117

were stored at -20°C until required. 118

Primers and Probes design. 119

Primer and probe sets were based on 18S rRNA sequences deposited in GenBank and were 120

designed using web-based software Primer3 v.0.4.0 (http://frodo.wi.mit.edu/primer3/) and/or 121

Primer Express Software (Applied Biosystem, Foster City, CA). The Plasmodium genus primers 122

and probe were designed to amplify all units of rRNA distributed in all the chromosomes; 1, 5, 7, 123

11 and 13. They were also designed to amplify the two types of Plasmodium 18S rRNA genes, 124

the S-type, expressed primarily in the mosquito vector, and the A-type, expressed primarily in 125

the human host (8, 22). The region of sequences selected were highly conserved and only found 126

in the Plasmodium genus. The sequence of the forward primer is 5´-127

GCTCTTTCTTGATTTCTTGGATG-3´ and the reverse primer is 5´-128

AGCAGGTTAAGATCTCGTTCG-3´. The probe sequence is ATGGCCGTTTTTAGTTCGTG, 129

labeled with 5´FAM (6-carboxyfluorescein) and 3´TAMRA (6-carboxytetramethyl-rhodamine) 130

as the reporter and quencher respectively. For the P. falciparum species-specific primers and 131

probe, we used previously published sequences but used VIC instead of FAM as the reporter dye 132

(18, 20). 133

qRT-PCR and qPCR. 134

Amplification and real-time measurements were performed in the Applied Biosystem 7500 135

analytical PCR system with the following thermal profile for qPCR: 10 min at 95°C; 40 cycles of 136

15 s at 95°C; 1 min 60°C. For qRT-PCR, a 30 min cycle at 50°C was added as the initial step for 137

the reverse transcription process. For the reaction, 1 µL of template was added to 9 µL of 138

reaction master mix containing 1X QuantiTect Probe RT-PCR Master Mix (QIAGEN, USA), 0.4 139

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µM of each primer, 0.2 µM probe and 4 mM MgCl2. For qRT-PCR assay, QuantiTect RT Mix (a 140

blend of Omniscript and Sensiscript Reverse enzymes) was added to the reaction master mix as 141

recommended by the manufacturer at a rate of 1 µL per 100 µL of the reaction master mix. 142

Standard curves. 143

For standard curve, cultured highly synchronized ring stages 3D7 parasites were used in order to 144

emulate infected human blood samples. The percent parasitemia of the ring stage was 145

determined by flow cytometry and microscopy. To determine the parasite/µL in culture material, 146

we multiplied percent parasitemia with the number of RBCs/µL which was counted by Coulter 147

analysis (Coulter AC·T 5 diff CP; Beckman Coulter, Inc., Miami, FL). The limit of detection 148

(LOD) for the PCR assays was established by creating a standard curve using cultured 149

synchronized ring stages 3D7 parasites that were serially diluted using uninfected whole blood 150

prior to total nucleic acid (NA) extraction. When analyzing and quantifying clinical samples, 151

each ninety-six well plate was ran with the standard 3D7 NA which was serially diluted five-fold 152

starting at 20,000 to 0.256 parasite/µL. Total nucleic acid (RNA and DNA) was extracted using 153

QIAamp DNA Blood Mini Kit using 200 µL of blood (or cultured material) and eluted in 200 µL 154

of water. For each experiment, we used 1 µL of NA template which is equivalent to 1 µL of 155

blood from a patient or cultured material. 156

Statistical analysis. 157

For statistical analysis, a two-tailed paired t test in GraphPad prism was used. 158

159

Results 160

Comparison of limits of detection between qRT-PCR and qPCR assays. 161

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Standard 3D7 NA was used to establish LOD which was set as the lowest NA concentration at 162

the threshold cycle number (Ct) at which normalized reporter dye emission rose above 163

background noise. For the genus-specific qRT-PCR assay, LOD was determined to be 0.002 164

parasites/µL and 0.0512 parasites/µL for qPCR assay (Fig. 1 (a) and (b)). For P. falciparum 165

species-specific assay, LOD was determined to be 1.22 parasites/µL for qRT-PCR and 2.44 166

parasites/µL for qPCR (data not shown). We assessed the reproducibility of the qRT-PCR and 167

qPCR genus-specific assays with respect to both intra- and inter-operator variability on replicate 168

samples conducted on different days. The qRT-PCR assay was found to be more sensitive over a 169

wide dynamic range of known parasite densities compared to the qPCR assay. Both assays were 170

highly reproducible with a mean coefficient of variation of 3% across between different 171

operators performing assays on different days (Fig. 2). Next, we randomly picked a clinical 172

sample that had been established to be P. falciparum positive by microscopy and assessed LOD 173

for both genus-specific and P. falciparum species-specific assays by serially diluting the sample. 174

The LOD for genus-specific assay was established to be 0.00661 parasites/µL for qRT-PCR and 175

0.0297 parasites/µL for qPCR (Fig. 3 (a) and (b)). For P. falciparum species-specific assay, the 176

LOD was established to be 1.82 for qRT-PCR and 3.41 parasites/µL for qPCR (data not shown). 177

We tested the specificity of the assays by ensuring the assay does not amplify human NA. 178

Establishing Assay sensitivity by inclusion of reverse transcriptase step in clinical samples. 179

We then compared the performance of qRT-PCR and qPCR in 603 clinical samples using genus-180

specific or P. falciparum species-specific assays in paired t test. There was a significant 181

difference in performance of qRT-PCR and qPCR for both genus-specific and P. falciparum 182

specific assays. For the genus-specific assay, the Ct values for qRT-PCR and qPCR were 183

significantly different from each other (P value <0.0001) with a mean ± SEM of 17.69 ± 0.2393 184

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and 22.44 ± 0.2373 respectively. The difference between the Mean Ct values for qRT-PCR and 185

qPCR assays was 4.757 ± 0.3370. For the P. falciparum species-specific assay, the Ct values for 186

qRT-PCR and qPCR were significantly different from each other (P value <0.0001) with a mean 187

± SEM of 25.27 ± 0.2564 and 27.12 ± 0.2343 respectively. The difference between the Mean Ct 188

values for qRT-PCR and qPCR was 1.756 ± 0.3713. To show how inclusion of the RT step into 189

the qPCR assay increases the sensitivity of the assay, the difference in Ct (∆Ct) for each clinical 190

sample between the qRT-PCR assay and the qPCR assay was plotted against the parasite density 191

as determined by thick blood smear (Fig. 4). Over 5 log-fold differences in parasite density, the 192

inclusion of reverse transcriptase enzyme into the qPCR assay increased the sensitivity of the 193

assay (samples with net Ct > 0). 194

Comparison of quantification by microscopy to qRT-PCR quantification. 195

We analyzed clinical samples that had no parasites based on microscopy using genus-specific 196

qRT-PCR assay. Of the 130 samples analyzed, 117 (90%) were positive by qRT-PCR with mean 197

Ct value of 19.60 with the lowest and the highest Ct values of 11.46 and 39.41. These Ct values 198

correspond to quantitative values of 4.65X104

parasites/µL and 0.000362 parasites/µL 199

respectively. We next determine whether inclusion of the reverse transcriptase enzyme into the 200

qPCR assay affected the quantification of the parasites in the blood over a range of parasite 201

densities (42-1.17E6 parasites/uL). From 466 clinical samples, we correlated the parasite density 202

as determined by microscopy with both qRT-PCR and qPCR genus-specific assays (Fig. 5) and 203

measured the statistical significance of each assay for either all samples or samples whose 204

parasite density was stratified into sub-groups. There was a statistically significant correlation 205

between parasite density measured by microscopy against either qRT-PCR (Fig. 5(a)) or qPCR 206

(Fig. 5(b)) molecular assays. However, the qPCR assay outperformed the qRT-PCR assay for 207

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each sub-group examined. The correlation was weakest at low parasite densities in both assays 208

with increasing divergence of the 95% confidence intervals as parasite density decreased. 209

Diluting clinical samples extends qRT-PCR dynamic range. 210

We observed that at high parasite densities, quantitative PCR reached a plateau limiting the 211

dynamic range of the qRT-PCR. We hypothesized that the dynamic range of the qRT-PCR can 212

be extended by further diluting the clinical samples. As such, we randomly picked 95 samples 213

with Ct values <18 and performed a tenfold serial dilution of the NA to 10-4

. The diluted samples 214

were analyzed by genus-specific qRT-PCR assay. Before dilution, the mean parasite equivalent 215

as determined with qRT-PCR assay was 2.09E+04 parasites/µL but after diluting, the mean 216

parasite equivalent increased to 4.33E+05 parasites/µL. Interestingly, the mean parasite density 217

of these samples by microscopy was 2.41E+05 parasites/µL, clearly showing that dilution of 218

extracted NA correlates well with microscopy at high parasite densities. These data represents 219

more than a log fold increase in the amount of parasite detected by qRT-PCR after diluting the 220

clinical samples, proofing our hypothesis to be true. 221

Discussion 222

The detection of parasites in sub-clinically subjects infected with either P. falciparum or P. vivax 223

malaria is a stated goal of the malaria vaccine and drug development programs within the Walter 224

Reed Army Institute of Research because of the increasing reliance on the human malaria 225

challenge model to assess the efficacy of malaria vaccines or the evaluation of new antimalarials 226

drugs. Because the threshold for fever and clinical disease from both P. falciparum and P. vivax 227

malaria is < 10 parasites/uL blood, newer and reliable diagnostic tests that are ultra- sensitive and 228

specific for malaria are needed to replace the Giemsa-stained thick blood smear, the current gold 229

standard test. It has been proposed that concomitant use of a molecular-based assay in detecting 230

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Plasmodium parasites would be an excellent safeguard against possible false negative results as 231

determined by expert microscopists (3). Here, we have described a highly sensitive qRT-PCR 232

assay for detection of Plasmodium genus. The assay detects both ribosomal RNA and DNA, 233

taking advantage of the high copy numbers of ribosomal RNA present per genome in a parasite. 234

The assay has LOD of 0.002 parasites/µL which represents one of the lowest LOD reported thus 235

far for detection of Plasmodium. During the analysis of the clinical samples in this study, we 236

detected the presence of parasite NA in samples that were below LOD as established using 237

standard P. falciparum 3D7 strain. For instance on two separate occasions the assay detected two 238

samples that had 0.000362 and 0.0004 parasites/µL, which to our knowledge is the lowest 239

parasite density reported thus far in detection of Plasmodium by PCR. 240

Under ideal conditions, during PCR amplification, the amplicon amount doubles every 241

cycle (i.e. increases by one log2) and one ∆Ct corresponds thus to a twofold increase in analytical 242

sensitivity (4). Here, we have shown that introduction of the RT step in the genus-specific assay 243

improved the mean analytical sensitivity of the assay approximately ten-fold. We have also 244

shown that introduction of the RT steps improves analytical sensitivity of P. falciparum species-245

specific assay by more than three-fold. Several PCR assays for detection of Plasmodium genus 246

have been previously described. For example, Elsayed et al., (6) using molecular beacon probes 247

developed an 18S rRNA-based assay that could detect 0.004 parasites/µL blood but required two 248

separate PCR reactions. However, careful consideration of the results reported is critical as there 249

are many variables which affect the sensitivity of PCR assays. The sensitivity of PCR assays in 250

detection of Plasmodium at genus or species level is commonly described in parasites/µL or 251

percent parasitemia. While many laboratories might follow similar protocols, factors such as 252

level of parasite synchronization of cultured samples, dilution ranges, methods of nucleic acid 253

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extraction, molecular assay detection (real-time PCR, molecular beacon, nucleic acid sequence 254

based amplification (NASBA), gel electrophoresis) all contribute to differences in sensitivities 255

making cross-platform and cross-study comparisons difficult. As we move toward considering 256

PCR assays as the gold standard for malaria diagnosis and surveillance not only in clinical 257

research, but also in future monitoring and evaluation efforts in malaria control and elimination 258

campaigns, stringent standardization of such details needs to be harmonized. The sensitivity of 259

any new molecular-based diagnostic test must be compared to the current gold standard, 260

microscopy of thick blood smears. The difficulties associated with malaria diagnosis by 261

microscopy are numerous and accuracy depends on the microscopists’ ability, training, 262

experience, motivation and availability of laboratory resources. Microscopy can miss a 263

substantial proportion of P. falciparum infections both in endemic and non-endemic populations. 264

Malaria microscopy is a highly perishable skill requiring continued training, practice, and testing 265

of microscopists to maintain high level proficiency (16). There is wide variation among “expert” 266

microscopists in assessing parasite density and distinguishing different parasite species. The 267

Walter Reed experience in Africa and Southeast Asia have demonstrated the need for uniform 268

training, quality assurance, and standardized reporting methods to minimize errors that occur 269

with microscopy (5, 15). Literature analysis shows that there are frequent discrepancies and 270

discordant results between microscopy and PCR techniques. In a recent review by Berry at al., 271

(3), it was noted that greater than 17% of thin blood smears examined by microscopy were 272

corrected after checking by PCR. Additionally, the rate of misdiagnosis varied from 20% to 50% 273

for P. malariae, P. ovale or P. vivax. In mixed infections, almost all microscopy results were not 274

accurate. Using the genus-specific assay, we found 90% of microscopy negative samples to be 275

positive by genus-specific qRT-PCR assay. This is an unusually high number compared to what 276

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has been previously reported especially considering that three independent qualified 277

microscopists evaluated each smear. The large discrepancy between qRT-PCR and microscopy 278

can be attributed to the high sensitivity of our genus-specific qRT-PCR assay, presence of 279

Plasmodium not recognizable by microscopy, presence of Plasmodium nucleic acid circulating in 280

subjects/patients, poor staining procedures, etc. 281

Some of the important parameters in guiding malaria treatment include detection of the 282

presence/absence of the parasite, identification of infecting species, and determination of the 283

level of parasitemia. As demonstrated in this study, each assay has its strengths and weaknesses, 284

and applying uniform “one size fits all” approach is both naïve and potentially misleading. The 285

biggest difference was found at the lowest or the highest densities, with qRT-PCR density 286

reaching a plateau sooner compared to microscopy. Since the qRT-PCR is much more sensitive 287

than microscopy, we hypothesized that qRT-PCR was more reliable than microscopy at lower 288

parasite densities but not at higher parasite densities. We showed this to be true when we diluted 289

NA from the clinical samples and consequently obtained higher parasite densities by qRT-PCR. 290

When samples were grouped based on individual log densities, the group with log density of 291

three was quantitatively more congruent for both qRT-PCR and microscopy than groups with 292

higher log densities. In this group, genus-specific qRT-PCR had a mean Ct value of 18.75. As 293

such, we propose that if a clinical sample is quantitatively analyzed by PCR and has Ct values 294

below 18.00 (this number may vary depending on the sensitivity of the assay), the sample must 295

be diluted further in order to obtain a more reliable quantification. In addition, diluting the 296

sample might improve the sensitivity of the assay by diluting-out PCR inhibitors that might be in 297

the sample. Although most extraction kits claim that purified NA is free of protein, nucleases and 298

other contaminants or inhibitors, PCR-inhibitory components in blood such as hemoglobin and 299

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lactoferrin are always of concern. Inclusion of amplification facilitators such as a single-300

stranded-DNA binding protein or Bovine serum albumin in PCR reaction might prove to be 301

helpful (1). 302

The Malaria Eradication Research Agenda (malERA) (http://malera.tropika.net/) has as 303

its stated objective, “Improved and new tools and strategies for monitoring, evaluation, and 304

surveillance are needed to track program intervention coverage, impact on cases and 305

transmission, and progress towards elimination/eradication. New survey tools may help to 306

measure transmission more simply and directly, thus enabling detection of 'hot spots' requiring 307

additional resource allocation.” As we move to the next phase of malaria control, we embrace the 308

objectives of malERA and believe that additional tools including ultrasensitive molecular assays 309

are needed to assess pockets of residual malaria prevalence and transmission. We believe that 310

such molecular tools will undoubtedly require assays that are inexpensive, easy to use under a 311

variety of detection platforms, and that require minimal training. We propose that the assays we 312

describe here can fulfill the Target Product Profiles (TPP) for two separate indications, one for 313

quality control and quality assurance programs in malaria diagnosis (i.e. Reference Centers) and 314

the second for surveillance in malaria elimination and eradication campaigns. The qRT-PCR 315

assay is a good fit for the modified TPP for eradication agenda: the assay is highly sensitive, 316

reliable and accurate, compatible with most PCR systems found in resource constraint 317

environments and importantly cost effective through reduction in the volume of the PCR reaction 318

thereby reducing the cost-of-goods. 319

In handling of clinical samples, NA extraction and storage of the samples should be 320

considered as a critical bottleneck towards standardization of PCR as a gold standard for malaria 321

diagnosis. We propose standardization of NA extraction procedures (methods) and protocol 322

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across the board as an initial step towards standardization of PCR as a gold standard for malaria 323

diagnosis. Since QIAamp DNA Blood Mini Kit is one of the most widely used extraction kit, we 324

propose adapting usage of this kit as a standard procedure for extraction of NA from whole blood 325

for malaria diagnosis. Additionally, we propose that publically-available resource such as ATCC, 326

MR4, or the WHO should prepare and supply standard NA such as 3D7 to be used worldwide by 327

all researchers and in clinical settings for quantitation and assessment of PCR sensitivity. 328

A radical changeover from traditional malaria microscopy to molecular PCR-based 329

assays will require careful thought and consideration regarding the impact that such a change 330

would have on the clinical development pathway of malaria vaccines or drugs that are assessed 331

in human challenge models. At the Walter Reed Army Institute of Research, we have used 332

standard malaria microscopy by qualified expert malaria microscopists that undergo proficiency 333

training examination before each clinical trial as the definitive method that determines whether a 334

subject will initiate treatment after the identification of two unambiguous malaria parasites in 335

Giemsa-stained thick blood smears. This approach has been validated and used successful in 336

over 1000 human subjects in approximately 57 human clinical challenge trials to date (2011). 337

Importantly, the use of expert microscopists that detect the appearance of malaria parasites in the 338

peripheral circulation prior to the onset of symptoms consistent with clinical malaria is an 339

important objective in our studies. Therefore we are keen to consider alternative diagnostic 340

assays other than microscopy which can detect the presence of parasites earlier than microscopy. 341

Our eagerness to employ such an assay will be tempered by the potential for PCR contamination 342

of products which may inadvertently give a false positive result in someone who actually has no 343

malaria parasites which could lead to a false conclusion on the effectiveness of a malaria vaccine 344

or drug intervention. Nevertheless, with stringent quality assurance programs, segregation plans 345

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for sample preparation and assay detection, and endorsement by regulatory agencies, we believe 346

the time has come to transition from microscopy to molecular-based PCR assays. 347

In conclusion, we have described a highly sensitive quantitative real-time PCR assay for 348

detecting Plasmodium at genus level and shown that adding RT step significantly increases the 349

sensitivity of real-time PCR. We have shown that at high parasite density, quantitative PCR 350

assay reaches a limit of quantification which can be extended by further diluting the sample. Our 351

genus-specific qRT-PCR assay has proven to be extremely useful in detection of Plasmodium 352

while maintaining the same robustness and sensitivity. Towards this, the assay is currently being 353

used in our institute in support of human clinical trials for detection of both P. falciparum and P. 354

vivax. The assay is also being used in animal studies for detection of P. berghei, P. yoelli and P. 355

knowlesi. In addition, this assay was recently successfully used in detection of Plasmodium from 356

mosquitoes collected in the field for surveillance studies. 357

358

Acknowledgements 359

We would like to acknowledge Kathy Moch for supplying cultured highly synchronized ring 360

stages 3D7 parasites. 361

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children. PLoS Clin. Trials 1e32. 439

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Figure legends 460

Fig. 1. Amplification plot showing LOD for detection of Plasmodium. To establish LOD, 461

standard 3D7 NA was serially diluted five-fold starting at 2.00E+04 to 4.10E-04 parasite/µL and 462

then ran qRT-PCR or qPCR using genus-specific assay. (a) Amplification plot showing LOD by 463

qRT-PCR using standard 3D7 NA. The lowest amplification for qRT-PCR was 2.05E-03 464

parasite/µL and (b) for qPCR was 5.23E-02 parasite/µL. 465

466

Fig. 2. Reproducibility of the genus-specific assay is shown. Data from serially diluted standard 467

3D7 NA which was assayed on different days by different operators represented by different 468

color on the graph. Data shows that the assay is highly reproducible. 469

470

Fig. 3. Amplification plot showing LOD by qRT-PCR using a clinical sample that was serially 471

diluted five-fold. (a) The lowest amplification for qRT-PCR was 6.61E-03 parasite/µL and (b) 472

for qPCR was 2.97E-02 parasite/µL. 473

474

Fig. 4. Addition of the reverse transcriptase enzyme into the qPCR assay increases sensitivity. 475

The Ct values of clinical samples were determined using qRT-PCR and qPCR. The difference in 476

Ct (∆Ct) for each clinical sample was determined and plotted against the parasite density as 477

determined by thick blood smear. Increase in sensitivity of the assay by addition of RT step was 478

seen in all log groups. 479

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Figure 1 483

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Ct

values parasites/µL

14.9062 2.00E+04

16.4416 4.00E+03

18.5357 8.00E+02

21.0309 1.60E+02

23.559 3.20E+01

26.302 6.40E+00

30.0388 1.28E+00

32.7088 2.56E-01

33.9949 5.12E-02

36.2447 1.02E-02

37.8839 2.05E-03

Ct

values parasites/µL

20.1494 2.00E+04

22.5817 4.00E+03

24.7312 8.00E+02

27.4059 1.60E+02

29.605 3.20E+01

32.1082 6.40E+00

34.3021 1.28E+00

36.6053 2.56E-01

37.5859 5.12E-02

Genus-specific qRT-PCR assay using 3D7 NA (a)

Genus-specific qPCR assay using 3D7 NA (b)

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Figure 2 495

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parasite density (p/µL)

Ct

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Reproducibility of qRT-PCR from P. falciparum 3D7 parasite

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Figure 3 518

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Ct values parasite/µL

15.8998 3.99E+03

19.1753 604.76

20.9961 211.828

23.7627 43.0226

26.5109 8.83161

29.0228 2.07731

32.017 0.370064

37.2896 0.0177397

39.0027 6.61E-03

Ct

values parasites/µL

25.3387 118.962

27.69 24.9402

29.8554 5.91638

32.6855 0.902322

34.9499 0.200421

35.9695 0.101791

37.8223 0.0297205

(b)

(a)

Genus-specific qRT-PCR assay using clinical sample NA

Genus-specific qPCR assay using clinical sample NA

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Figure 4 537

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Effect of reverse transcriptase on sensitivty of real-time PCR assay

Increased

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Decreased

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Figure 5 550

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