Background
The genus
Plasmodium consists of over 200 widely distributed species, of which at least six are now known to regularly infect humans:
Plasmodium falciparum,
Plasmodium vivax,
Plasmodium malariae,
Plasmodium ovale wallikeri,
Plasmodium ovale curtisi,
Plasmodium knowlesi [
1]. Previously, malaria was thought to be primarily caused by just the four species,
P. falciparum,
P. vivax,
P. malariae and
P. ovale, and only very rarely by other
Plasmodium species. However, the entry of PCR-based techniques into malaria diagnostics has changed this view. It has led to the discovery of the existence of
P. ovale wallikeri and
P. ovale curtisi [
2], and the demonstration that
P. knowlesi is the predominant cause of malaria in regions of South East Asia. Interestingly, recent research indicates that also
P. cynomolgi may be more common in South East Asia than previously suspected [
3,
4]. In addition, human susceptibility to
Plasmodium simium and
Plasmodium brasilianum has been shown.
Plasmodium simium is found throughout South America, while
Plasmodium brasilianum seems restricted to a specific part of Brasil [
5‐
7]. Globally,
P. falciparum is responsible for the vast majority of malaria deaths and cases of severe malaria. Malaria caused by other
Plasmodium species is not typically as life threatening, but can also be lethal. In particular, severe and fatal cases of malaria caused by
P. vivax and
P. knowlesi are well recognized [
1].
The gold standard of malaria diagnosis is the examination of stained thick and thin blood films by light microscopy. The quantification of malaria parasites can be used to make clinical management decisions as well as monitoring response to treatment [
8]. Microscopic diagnosis of malaria is prone to human error owing to its subjective nature. An inherent weakness of microscopy is the dependence on morphological features to distinguish
Plasmodium species. Even under ideal conditions, reliable distinction of the infecting
Plasmodium species can be very difficult, if not impossible. Particularly,
P. vivax and
P. ovale cannot always be easily differentiated based on morphology, distinguishing
P. knowlesi from
P. malariae can be very challenging,
P. ovale wallikeri and
P. ovale curtisi are morphologically identical,
P. cynomolgi is morphologically indistinguishable from
P. vivax, and
P. simium and
P. brasilianum cannot be distinguished by microscopy from
P. vivax and
P. malariae, respectively [
9‐
11]. Also, the detection limit of microscopy is not always sufficient to detect low-density malaria infections. Submicroscopic infections are typically asymptomatic, but not benign. Such cases should be considered as chronic malaria, and should be treated accordingly. Untreated chronic malaria has significant health consequences for the individual, and, as it could potentially be transmitted, affects public health [
12‐
15].
A plethora of polymerase chain reaction (PCR)-based nucleic acid amplification tests (NAATs) to detect, quantify and identify Plasmodium parasites in blood have been described in the literature. Rapid NAATs with a turnaround time of less than one hour are now available, and some NAATs can be used for high throughput of samples. In addition, NAATs are used to detect anti-malarial drug resistance markers. NAATs offer several significant advantages over standard microscopy. Importantly, the sensitivity of NAATs for the detection of Plasmodium parasites in blood is superior to any other diagnostic test. The general ability to detect far less than ten Plasmodium parasites per microlitre of blood provides the sensitivity to rule out malaria with high negative predictive value in clinical settings. In endemic settings, where the goal is to eradicate malaria, the required sensitivity to reliably detect submicroscopic infections can be met. Also, NAATs tend to be highly specific. Identification of Plasmodium species goes beyond what is possible with conventional microscopy. For example, for differentiating Plasmodium species with similar morphologies, detecting mixed Plasmodium infections, utilization of poor-quality or ambiguous samples, and when differentiation of Plasmodium species or strains is impossible based on morphology alone.
However, an important pitfall to be aware of is that NAATs not specifically designed to identify a certain Plasmodium species may fail to detect it or cause misidentification. Individual patient treatment may not always be affected, but such errors could ultimately prompt public health concerns by compromising malaria control and elimination.
Other concerns regarding the implementation of NAATs are the special equipment and reagents required, and the need for skilled laboratory technicians able to perform testing and to interpret the results of these relatively complex tests.
Furthermore, test characteristics vary widely, and although the World Health Organization (WHO) International Standard for
P. falciparum [National Institute for Biological Standards and Control (NIBSC)-code 04/176] allows for comparison among the available assays, there is a lack of standardization and validation. As science and technology proceed, NAATs are bound to play an increasingly important role in malaria diagnosis, research, and epidemiology. At present, it is clear that NAATs cannot completely replace traditional microscopy [
16‐
18].
The objective of the present study was to determine the analytical performance of the novel MC004 real-time PCR assay for malaria diagnosis (MRC Holland, Amsterdam, the Netherlands). This single-tube assay has been developed for the purpose of simultaneously detecting all Plasmodium species known to infect humans, and discrimination between P. falciparum, P. vivax, P. malariae, P. ovale wallikeri, P. ovale curtisi, P. knowlesi (including differentiation of three strains) and P. cynomolgi (including differentiation of three strains). In addition, this study shows how this assay might be used to estimate the parasitaemia of at least P. falciparum.
Methods
Analytical specificity study design
The analytical specificity of the assay was determined by testing samples received from reference institutes or referral laboratories [the Dutch Foundation for Quality Assessment in Medical Laboratories (SKML), UK NEQAS, Streeklab Haarlem—Laboratory for Medical Microbiology, the Biomedical Primate Research Centre, the London School of Hygiene & Tropical Medicine, and Labor Dr. Wisplinghoff]. In total, 40 reference samples were tested that included all seven
Plasmodium species (
P. falciparum,
P. vivax,
P. knowlesi,
P. malariae,
P. cynomolgi,
P. ovale wallikeri, and
P. ovale curtisi) at various levels of parasitaemia, and four different non-
Plasmodium parasites:
Babesia microti,
Loa loa,
Trypanosoma brucei rhodesiensis, and
Leishmania donovani infantum. Specimen types included ethylenediaminetetraacetic acid (EDTA) whole blood, isolated DNA, and Giemsa-stained blood or bone marrow slides (Table
1). The results of the MC004 assay were compared with the results of the reference laboratory.
Table 1
Analytical specificity panel
Dutch Foundation for Quality Assessment in Medical Laboratories (SKML) The Blood- and intestinal parasites scheme (n = 7) | P. falciparum (2019.1B) | Blood smear | 1.9% |
P. falciparum (2019.3E) | EDTA whole blood | Unknown |
P. falciparum (2019.3F) | EDTA whole blood | Unknown |
P. vivax (2018.1G) | EDTA whole blood | Unknown |
P. ovale (2017.4B) | Blood smear | Unknown |
P. knowlesi (2016.3B) | Blood smear | 4.2% |
Trypanosoma brucei rhodesiensis (2019.1A) | Blood smear | Not applicable |
UK NEQAS (n = 3) The Blood parasitology scheme | P. malariae (5796) | Blood smear | Unknown |
Loa Loa (5211) | Blood smear | Not applicable |
P. falciparum + P. vivax (5844) | Blood smear | 0.2% |
UK NEQAS The Malaria molecular scheme (n = 8) | No parasites (5220) | EDTA whole blood | Not applicable |
No parasites (5222) | EDTA whole blood | Not applicable |
No parasites (5363) | EDTA whole blood | Not applicable |
No parasites (5364) | EDTA whole blood | Not applicable |
P. vivax (5365) | EDTA whole blood | Unknown |
P. vivax (5366) | EDTA whole blood | Unknown |
P. falciparum (5221) | EDTA whole blood | 0.0002% |
P. knowlesi (5219) | EDTA whole blood | 0.02% |
Streeklab Haarlem—Laboratory for Medical Microbiology (n = 17) | P. falciparum | Blood smear | 4.5% |
P. falciparum | Blood smear | 1.3% |
P. falciparum | Blood smear | 0.5% |
P. falciparum | Blood smear | 9.9% |
P. falciparum | Blood smear | < 0.1% |
P. falciparum | Blood smear | Gametocytes |
P. vivax | Blood smear | Unknown |
P. vivax | Blood smear | Unknown |
P. vivax | Blood smear | Unknown |
P. ovale | Blood smear | Unknown |
P. ovale | Blood smear | Unknown |
Leishmania donovani infantum | Bone marrow smear | Not applicable |
Leishmania donovani infantum | Bone marrow smear | Not applicable |
No parasites | Blood smear | Not applicable |
No parasites | Blood smear | Not applicable |
P. knowlesi | Blood smear | Unknown |
Babesia microti | Blood smear | Not applicable |
Biomedical Primate Research Centre (n = 2) | P. knowlesi (H strain) | Isolated DNA | Not applicable |
P. cynomolgi (M strain) | Isolated DNA | Not applicable |
The London School of Hygiene & Tropical Medicine (n = 2) | P. ovale wallikeri | Isolated DNA | Not applicable |
P. ovale curtisi | Isolated DNA | Not applicable |
Labor Dr. Wisplinghoff (n = 1) | P. knowlesi | EDTA whole blood | 0.3% |
MC004 assay technology
The MC004 assay (MRC-Holland, Amsterdam, the Netherlands) is a multiplex nucleic acid amplification test that detects mitochondrial
Plasmodium DNA encompassing the cyclo-oxygenase 3 (COX-3), cyclo-oxygenase 1 (COX-1) and cytochrome
b (CYTB) genes from
Plasmodium species that cause malaria in humans. The assay provides a qualitative result for the presence/absence of
Plasmodium DNA, and discriminates between 11
Plasmodium species/strains (
P. falciparum,
P. vivax,
P. malariae,
P. ovale wallikeri,
P. ovale curtisi,
P. knowlesi LT48,
P. knowlesi ATCC 30153,
P. knowlesi ATCC 30158
P. cynomolgi ATCC 30149,
P. cynomolgi KJ569866.1 and
P. cynomolgi KJ569868.1). The MC004 assay involves two main steps, which take place in a single tube: (1) asymmetric target amplification by two different primer sets (primer pair 1: 5ʹ-TCGCTTCTAACGGTGAACT-3ʹ/5ʹ-AAGCAAACACTAGCGGTGGAA-3ʹ and primer pair 2: 5ʹ-CAGTATAATATTGTAATTTGATCAGTATGAG-3ʹ/5ʹ-GGATATTGTATAAATGATGCTATATCAGGTA-3ʹ). Primer pair 1 was designed to amplify a 116 base pair fragment of all 11
Plasmodium species/strains. Primer pair 2 was designed to amplify a 212 base pair fragment of
P. vivax,
P. knowlesi, and
P. cynomolgi only. See Table
2 for the precise annealing positions of the primers. Amplification can be monitored real-time in the FAM detection channel due to the presence of a nonspecific green fluorescent nucleic acid dye that becomes fluorescent upon binding to double stranded DNA. (2) detection and differentiation of the amplicons by molecular beacon probe-based melting curve analysis. The assay uses three different probes, which are either Texas red-labelled (identical to the sequence of
P. knowlesi ATCC 30153), Cy5-labelled (identical to the sequence of
P. falciparum), or Cy5.5 labelled (identical to the sequence of
P. knowlesi LT48).
Table 2
Annealing positions of the forward and reverse primers of primer pairs 1 and 2
P. falciparum | LR605957.1 | Forward: 339–357 Reverse: 434–454 | Not applicable |
P. vivax | KY923424.1 | Forward: 1493–1511 Reverse: 1588–1608 | Forward: 1941–1971 Reverse: 2122–2152 |
P. malariae | AB489194.1 | Forward: 1560–1578 Reverse: 1655–1675 | Not applicable |
P. ovale wallikeri | HQ712053.1 | Forward: 1038–1056 Reverse: 1133–1153 | Not applicable |
P. ovale curtisi | HQ712052.1 | Forward: 1038–1056 Reverse: 1133–1153 | Not applicable |
P. knowlesi | LR701176.1 | Forward: 340–358 Reverse: 435–455 | Forward: 781–811 Reverse: 962–992 |
P. cynomolgi | AB444125.1 | Forward: 1605–1623 Reverse: 1700–1720 | Forward: 2051–2081 Reverse: 2232–2262 |
Specimen processing
The preferred specimen type is DNA extracted from 200 µL of (EDTA) human whole blood. DNA was isolated using the QIAamp DNA Blood Mini QIAcube Kit (Qiagen, Hilden, Germany) on a QIAcube instrument (Qiagen, Hilden, Germany) following the manufacturer’s specifications. DNA was eluted with 100 µL of elution buffer. Giemsa-stained blood smears were processed by pipetting 10 µL of 10 mM sodium hydroxide (NaOH pellets, 99%, Merck, Darmstadt, Germany) onto the surface of the blood smear. Subsequently, blood smear material was scraped off from the glass slide by making circular movements with a sterile lancet (Solofix, B Braun, Oss, Netherlands). The collected specimen was transferred to a sterile 2.0 mL tube (Eppendorf, Hamburg, Germany) that contained 100 µL of 10 mM sodium hydroxide. Processed specimen samples were stored at − 30 °C.
Amplification and detection
Each reaction was carried out in a final volume of 25 µL reaction mixture containing 23 µL of MC004 master mix (MRC-Holland, Amsterdam, the Netherlands) and 2 µL of template. The components of the master mix were: both forward primers (1.25 µM), both reverse primers (0.25 µM), all three probes (0.5 µM), fluorescent compound, dNTPs, reaction buffer, and Taq DNA polymerase. The following settings were used for the amplification step: 95 °C for 3 min, followed by 50 cycles of 95 °C for 15 s, 60 °C for 30 s and 68 °C for 40 s. The following settings were used for the melting curve step: a gradual temperature increase from 25 to 69.4 °C (0.4 °C per 5 s).
Controls
Positive and negative control samples were used to validate run validity. Each run included two negative controls: (1) with elution buffer added as template and (2) a fresh uninfected whole blood specimen processed as a separate sample. Positive control samples for each Plasmodium species come with the MC004 assay as provided by MRC Holland. The 1st WHO International Standard for P. falciparum DNA (NIBSC code: 04/176) diluted in EDTA whole blood (before DNA isolation) to a concentration of 1 × 10–3 IU/mL was used as a positive control for limit of detection. Each sample should produce an amplification curve, if no amplification curve is observed, this may be a sign of inhibition and renders the result invalid.
Instrumentation
The assay was performed on the CFX96 Touch Real-Time PCR Detection System (Bio-Rad, Hercules, CA, USA). Runs were analysed using the accompanying Bio-Rad CFX manager 3.1 software (Bio-Rad, Hercules, CA, USA). Cq-values were determined by the software’s Cq Determination Mode (settings: Single Threshold set at 2000; Baseline Setting: Baseline Subtracted Curve Fit).
Melting temperatures were determined using the software’s default settings. The throughput is approximately 80 specimens for 1 operator in about 3.5 h. Up to 12 specimens can be processed, including DNA isolation using the QIAcube, within four hours of arrival of the specimens at the laboratory.
Limit of detection study design
The limit of detection of the MC004 assay was determined by preparing a tenfold serial dilution of the WHO International Standard for
P. falciparum DNA (NIBSC, Hertfordshire, England) in uninfected EDTA whole blood to achieve a concentration range of 1 × 10
9 IU/mL to 1 × 10
–9 IU/mL. To clarify, this WHO standard was assigned an arbitrary unitage in International Units (IU), unrelated to the level of parasitaemia or DNA copy number [
19]. The lyophilized WHO standard, reconstituted in 500 µL of sterile water, was diluted in EDTA whole blood. Total volume of each dilution was 400 µL. DNA was isolated as described above, and samples were tested in three independent runs. The limit of detection was determined as the lowest concentration of
P. falciparum detected in all three runs. In addition, similar tenfold serial dilutions of patient samples with known parasitaemia were tested:
P. falciparum 1.3%
, P. falciparum 0.2%,
P. vivax < 0.1%
, P. malariae 0.3%,
P. ovale curtisi < 0.1%
, and
P. knowlesi ATCC 30158 0.3%. Samples were tested in two independent runs. The limit of detection was determined as the lowest concentration of the
Plasmodium species detected in both runs.
Detection of mixed infections study design
Mixed Plasmodium infections were created by adding together two DNA samples, each containing a different Plasmodium species (P. falciparum and a non-falciparum species) 1:1 (equal volume). When applicable, the parasitaemia was determined by experienced microscopists (P. falciparum 0.2%, P. vivax 0.1%, P. ovale wallikeri 0.2%, P. malariae 0.4%, P. knowlesi ATCC 30158 0.3%, P. ovale curtisi and P. cynomolgi KJ569868.1 isolated DNA diluted 100 times with sterile water). In addition, a two-fold serial dilution (1:2, 1:4, 1:8, 1:16, 1:32, 1:64, 1:128) of a P. falciparum DNA sample (initial parasitaemia was 1.3%) in sterile water was prepared to create mixed infections of P. falciparum and a non-falciparum species (non-falciparum samples as described above).
Quantification of parasitaemia study design
To quantify parasite density, a calibration curve was established by plotting the microscopically determined parasite densities of ten samples from patients with P. falciparum infections as a function of the Cq-value. The parasite densities were: 0.1%, 0.1%, 0.4%, 0.8%, 1.3%, 1.5%, 3.7%, 23.7%, 31.8%, and 35.8%. Cq-values were determined in three independent runs. The exponential equation (y = a*e(b*x)) of the line of best fit through the 30 data points was determined using Microsoft Excel for Mac 2016, where ‘y’ is the parasitaemia (percentage of infected erythrocytes), ‘x’ is the Cq-value (threshold cycle), ‘a’ and ‘b’ are the dimensionless coefficients that define the equation, and ‘e’ is the base of the natural logarithm.
Precision of the quantification of the parasitaemia study design
Within-run and between-run precisions [standard deviations (SD) and coefficients of variation (CV)] for the quantification of the parasitaemia were determined using four clinical samples infected with
P. falciparum. The parasitaemia levels of these four samples were estimated by real-time PCR to be around 0.1%, 0.2%, 2% and 27%, respectively. Precision was assessed by testing five replicates per run over five runs. Each run the samples were tested in different wells of the PCR plate. Within-run and between-run precision were calculated as described by Chesher, 2008 [
20].
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