Giemsa-stained thick and thin blood smears are the gold standard diagnostic method for malaria. Where microscopy may miss very low density parasitemia infections, molecular assays with sensitivities of 20–50 parasites/mL allow detection of pre-patent infections in humans experimentally infected with malaria parasites. 29,32,42,46
Detection of pre-patent parasitemia and determination of the parasite growth rate can help to determine whether vaccine candidates affect liver- and/or erythrocyte-stage parasite growth. 42
There are many factors to consider when choosing from the diverse molecular methods reported for malaria. 5–23
First, pre-analytical handling steps (such as leukocyte filtering reported in some assays) should be minimized or highly controlled to reduce the risk of contamination. Second, as the sample volume decreases, the limit of detection increases. For instance, if 200 μL of blood is extracted, the lowest possible limit of detection is 5 parasites/mL because there can be at a minimum only 1 parasite/200 μL, whereas for 50 μL samples, the lowest limit of detection is 20 parasites/mL (or 1 parasite/50 μL). The actual limit of detection is further affected by the blood sample volume, the extracted nucleic acid volume, the volume added to the (RT)-PCR assay and the Poisson distribution. Low parasitemia samples can be difficult to detect using single-step DNA PCR approaches, 14
although reports vary. 8
Some RNA assays have improved sensitivities at low parasite densities because of the increased copy number relative to coding DNA. 22
Third, many assays do not include a multiplexed internal control, which is critical for ruling out PCR inhibitors in every sample.
To overcome these limitations, we developed a high sensitivity, internally controlled real-time RT-PCR assay for absolute P. falciparum quantification and demonstrated its use in a clinical trial. The high sensitivity of qRT-PCR from a 50 μL blood sample is possible because 18S rRNAs are abundant within individual parasites (1×104 copies/ring form). Thus, we obtain three pieces of evidence to support the test result: the malaria CT, the malaria PCR product Tm, and a reciprocal decrease in total competitor amplification. We include a competitive internal control RNA in every multiplexed reaction to detect target extraction/amplification failures caused by inhibitors, laboratory errors, and/or other Plasmodium infections.
Using a combination of purified RNA calibrators, synchronized (12-hr) ring-stage parasite standards in whole blood, and synchronous parasite cultures sampled throughout the 48-hr lifecycle, we measured 10,000 copies of the 18S rRNA per ring-stage parasite, in agreement with the conversion factor used for NASBA assays. 7
Although RT-PCR on aliquots of frozen whole blood were recently reported to yield ≤ 1-log increase in parasite detection compared with PCR alone, 23
we find a nearly 4-log increase in detection, probably because 18S rRNA are better preserved when lysed in guanidinium-based buffer before and stored at −80°C. To generate a standard curve for absolute quantification, we used RNA standards of known quantity added to extracted nucleic acids from whole blood of known erythrocyte density. As noted previously, we can convert log10
RNA copies to parasite density, allowing for absolute quantification of P. falciparum
from whole blood.
With this system, we validated the qRT-PCR for clinical trial use and used it in a demonstration human challenge trial of six volunteers. The assay was highly sensitive and specific across a wide range of parasitemia and performed favorably in accuracy, correlation, precision, analytical sensitivity, analytical specificity, and carryover evaluations. This assay detected sub-patent parasitemia 3.7 days on average before blood smears were positive, similar to that seen in other trials using other techniques. 42,45
Although it must be remembered that Poisson statistics affect the confidence limits for positive detection at parasite densities < 60 parasites/mL, we routinely detect samples containing as few as one parasite per 50 μL of whole blood (i.e., 20 parasites/mL). However, we did not test samples at a nominal concentration of 20 parasites/mL because parasites would be statistically absent from 37% of such samples. Because our RNA standards can be detected > 95% of the time at copy numbers below that expected for a single parasite in 50 μL of blood, we can quantitatively report parasite densities down to 20 parasites/mL, recognizing again that the Poisson distribution influences the actual outcome.
In P. falciparum
, the expression of the A-type 18S rRNA target during ring-stage development was critical for the interpretation of our assay. Here, we report that the A-type 18S rRNA is stably expressed throughout the ring stage of the P. falciparum
lifecycle. The 18S rRNA has not been previously profiled by quantitative molecular methods in P. falciparum
, but our data is generally consistent with earlier Northern blotting of murine P. berghei
, which showed that A-type 18S rRNA expression was maximal in the growing trophozoite stage and lower in the transcriptionally quiescent ring stage. 47
In P. falciparum
infections, only ring-stage parasites are detected in peripheral blood during the first 24 hrs after erythrocyte invasion. Although mature trophozoite and schizont forms are easily cultured in vitro
, such mature forms are not usually present in peripheral blood in vivo
because of cytoadherence during the latter half of the lifecycle. 48
Mature forms are only observed in the peripheral blood in cases of severe hyperparasitemia. For non-P. falciparum
species, sequestration does not occur and all lifecycle forms are present in peripheral blood, making quantification of other species more complicated. Thus, for the relevant ring-stage P. falciparum
parasites, the 18S rRNA signal is stably expressed in vitro
. Indirect evidence for stable ring-stage expression of the target in vivo
comes from the clinical trial data, which showed rising cyclical parasitemia in participants by qRT-PCR that was consistent with data from other centers using DNA-based quantification. Such agreement would be unlikely if the RNA target varied dramatically during in vivo
infections. Whether stable expression typifies in vivo
infections in the presence of sub-therapeutic anti-malarial drugs or in infections with mixed strains is unknown.
RNA-based assays for malaria allow for high sensitivity detection and quantification of parasites from a small sample. To reap the benefits of the high copy number 18S rRNA as a target, samples must be adequately preserved at the time of collection, which is easily accomplished using guanidinium-based lysis buffer and, possibly, even dried blood spots. 49–51
Given the need for reliable, controlled assays to support malaria clinical trials and eradication efforts, we recommend the use of multiplexed internal control targets and in vitro transcribed RNA standards for absolute quantification in all RNA- or total nucleic acid-based assays. Furthermore, a network for inter-laboratory exchange of well-characterized malaria comparator samples would be useful to help laboratories establish and maintain assay performance. Such quality control programs for malaria molecular diagnostics would benefit research and clinical efforts alike.