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Standard malaria diagnosis is done by microscopic examination of stained blood films.1 This has undergone very little change since Laveran’s original discovery of the malaria parasite and improvement in staining techniques by Romanowsky in the late 1800s. More than a century later, microscopic detection and identification of Plasmodium species in Giemsa-stained thick (screen for presence of parasites) and thin blood smears (confirmation of species) continues to be the gold standard for the laboratory diagnosis of malaria. The wide acceptance of this technique can be attributed to its simplicity, low cost, and ability to identify infecting species and quantify parasitemia, both of which are required for selecting appropriate antimalarial drugs and monitoring their effect. It is, however, time-consuming, labor-intensive, and requires considerable expertise.2,3 The most important shortcoming of microscopy is its relatively low sensitivity especially at low parasite levels where the likelihood of parasite detection and correct species diagnosis depends on parasite density.4,5 Although experienced microscopists can detect up to 20 parasites/µL,2 routine diagnostic laboratories have a much lower sensitivity of detection (500 parasites/µL, 0.01% of erythrocytes infected).3 This has probably resulted in underestimation of malaria infection rates, especially those with low parasitemia and asymptomatic malaria. Use of the polymerase chain reaction (PCR), a highly sensitive and specific technique for the detection of all species of malaria parasite in whole blood, has been and continues to be extensively used to diagnose malaria, follow patients’ response to therapy,4 and identify drug resistance.5
We undertook this study to detect malaria infection retrospectively using stored serum samples. Serum, rather than the hitherto standard whole blood as a PCR specimen, is more readily available as it is commonly stored. For example, serum samples from patients with human immunodeficiency virus (HIV)/acquired immunodeficiency syndrome are routinely archived for future investigations. Because of a wide geographic overlap of HIV and malaria, co-infection rates are high.6 This interaction has major public health significance because when both pathogens infect the same host, transmission, clinical manifestations, and treatment outcomes of both diseases are impacted. A quick and cost-effective method of determining co-infection rates would be to detect Plasmodium DNA from stored serum samples collected from HIV-positive patients in the course of routine patient care or other research studies. There have been no studies done on detection of P. vivax from serum and there are no data on the stability of parasite DNA stored for prolonged periods of time. We undertook this study to test the hypothesis that P. vivax and P. falciparum DNA can be detected from serum samples collected from malaria patients that has been stored for more than two years.
The details of patient selection and enrollment have been reported.7 Briefly, 2–4 mL of blood was collected from microscopy-confirmed malaria patients in Iquitos, Peru. Serum was separated after allowing the blood to clot and stored at –20°C for an average period of 6 weeks before being transported to the United States on dry ice. The samples were then kept in a –80°C freezer for an average of 30 months before parasite DNA extraction and detection by PCR. DNA was extracted from 200 µL of serum sample using the QIAamp DNA Blood Mini Kit (catalog no. 51106; Qiagen, Valencia, CA) according to the blood and body fluid protocol. DNA extracted from 200 µL of heparinized whole blood samples from known smear-positive P. vivax and P. falciparum malaria patients was used as positive controls. Similarly, DNA from 200 µL of serum from field-collected smear-negative patients was used as a negative control.
Nested PCR was done using a modification of the technique originally described by Snounou and others with primers targeting the Plasmodium spp. 18S small subunit ribosomal RNA genes.8 The first PCR was performed in a total volume of 50 µL containing 5 µL of extracted DNA, 25 µL of HotStar Taq Master Mix (catalog no. 203443; Qiagen), and forward and reverse primers (0.2 µM). The nested species-specific PCR was performed in a total volume of 25 µL containing 1 µL of PCR product. Cycling conditions were same for both PCR cycling procedures: incubation at 95°C for 15 minutes, followed by 35 cycles at 95°C for 30 seconds and 58°C for 1 minute, with a final incubation at 72°C for 1 minute. Primer sequences were as follows: for the first round of PCR, 5'-TTAAAATTGTTGCAGTTAAAACG-3' (sense) and 5'-CCTGTTGTTGCCTTAAACTTC-3' (anti-sense); for the second round of PCR, P. falciparum primers 5'-TTAAACTGGTTTGGGAAAACCAAATATATT-3' (sense) and 5'-ACACAATGAACTCAATCATGACTACC-CGTC-3' (antisense); P. vivax primers 5'-CGCTTCTAGCT-TAATCCACATAACTGATAC-3' (sense) and 5'-ACTTCCAAGCCGAAGCAAAGAAAGTCCTTA-3' (antisense). The presence of amplification products was detected by ethidium bromide staining after agarose gel (1.8%) electrophoresis. All DNA samples were also amplified using primers for the human rRNA gene p53 to confirm the presence of amplifiable human DNA.9
Stored serum samples were available for 24 patients with a malaria diagnosis based on peripheral blood smear examination by the Ministry of Health (hospital or health post) technicians. Microscopy based on thick smear examination showed malaria caused by P. falciparum in 3 patients and P. vivax in the remaining 21 patients, with parasitemia levels ranging from 1 to 200 parasites per high-power field (HPF). The following semi-quantitative system was used to estimate parasitemia: < 1+, <1 parasite/100 HPF; +, 1–< 2parasites/HPF; ++, 2–20 parasites/HPF; +++, 21–200 parasites/HPF; ++++, > 200 parasites/HPF. As shown in Table 1
, PCR confirmed the presence of parasite DNA in all but 4 (2, 5, 7, and 8) samples (20 of 24, sensitivity = 83%). On reviewing peripheral blood smear for sample 2, the study microscopist found it to be negative for malaria and upon excluding it the PCR sensitivity increased to 87% (20 of 23). Samples 5, 7, and 8 were reported positive for P. vivax, both initially and on review, with parasitemias ranging from 1 to 20 parasites/HPF. Amplifiable human DNA, tested using primers for the p53 gene, was found in all but 1 (23 of 24) serum sample from malaria patients and all (8 of 8) malaria-negative controls, giving a detection rate of 97% (31 of 32). All 8 of 8 malaria-negative field collected samples were negative by PCR, giving the test a specificity of 100%. Sample 8, which did not have amplifiable human DNA, was also negative for parasite DNA by PCR. The second PCR using species-specific primers confirmed microscopy findings. Of 20 P. vivax smear-positive samples, 17 were confirmed by PCR. Of the other 3 samples reported as P. falciparum by smear examination, 2 had P. falciparum and the third was shown by PCR to have mixed (P. vivax and P. falciparum) infection. The study microscopist correctly identified the mixed infection missed by the health post technician. Results from a representative sample using DNA extracted from serum from patients with P. vivax, P. falciparum and mixed infection are shown in Figure 1. The PCR amplification product of 1,100 basepairs was seen in all 3 (Figure 1A), and 205-basepair and 120-basepair products were seen in patients with P. falciparum (Figure 1B) and P. vivax (Figure C) malaria, respectively. Patients with mixed infections had both PCR products.
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To date, PCR has typically been done using DNA extracted from whole blood. Plasmodium falciparum DNA from serum from microscopy-confirmed patients has previously been detected by PCR.10 However, in contrast to P. falciparum, para-sitemia with P. vivax is generally low (
1%), so that findings of the present study show that even P. vivax DNA can be detected in serum samples. Further retrospective diagnosis of mixed infection is feasible, as shown here, and may be of clinical importance when severe malaria complicates P. vivax malaria. The PCR assay also correctly diagnosed a mixed infection in one patient that was reported caused by P. vivax alone.
It is important to point out that serum was collected under usual field conditions and DNA extracted in a routine manner. This highlights the point that retrospective analysis for Plasmodium DNA from stored sera is possible. The presence of amplifiable parasite DNA in stored serum samples is required for successful diagnosis of malaria by PCR. We were surprised to be able to detect parasite DNA in most of the samples from malaria patients and amplifiable human DNA from all but one sample. We attribute the three non-concordant smear positive/PCR negative cases to either degradation of parasite DNA (as in sample 8) or low parasitemia combined with degradation of parasite DNA (as in samples 5 and 7).
Although the number of samples reported here are small, the high degree of both sensitivity and specificity is encouraging. Larger studies of both P. falciparum and P. vivax malaria in endemic regions, involving both symptomatic and asymptomatic parasitemic patients under varying conditions of transmission intensity (i.e., holoendemic versus seasonal sporadic malaria) will enhance the generalizability of the present findings.
Detection of Plasmodium DNA in serum samples by PCR may have an important application in retrospective diagnosis of malaria infection in specimen banks of cohort studies, such as in determining malaria co-infection in HIV-seropositive populations. Since the HIV epidemic, the practice of saving serum samples for clinical care and future research projects makes this diagnosis possible.
Received May 5, 2007. Accepted for publication June 5, 2007.
Acknowledgments: We thank our field team (Dr. Eddy Segura, Sonia Torres Andrade, and Nahir Chuquipiodo) for assistance; Carlos Pacheco and Flor Pacheco for technical help in Iquitos; and patients from the city of Iquitos and the villages of Varillal and Santo Tomas for their participation.
Financial support: This study was supported by an American Society of Tropical Medicine and Hygiene–Ellison Postdoctoral Fellowship in Tropical Medicine (Ajay R. Bharti), National Institutes of Health (NIH) institutional training grant T32A107036-26 (on which Ajay R. Bharti was supported), a Doris Duke Charitable Foundation Innovations in Clinical Research Program grant (Joseph M. Vinetz), National Institutes of Allergy and Infectious Diseases/NIH grant K24AI068903 (Jospeph M. Vinetz), and NIH Fogarty International Center Global Infectious Diseases Training grant 5D43TW007120 (Joseph M. Vinetz).
* Address correspondence to Joseph M. Vinetz, Division of Infectious Diseases, Department of Medicine, University of California San Diego, School of Medicine, 9500 Gilman Drive, 0741, George Palade Laboratories Room 125, La Jolla, CA 92093-0741. E-mail: jvinetz{at}ucsd.edu 
Authors’ addresses: Ajay R. Bharti, Kailash Patra, and Joseph M. Vinetz, Division of Infectious Diseases, Department of Medicine, University of California San Diego, 9500 Gilman Drive, 0741, George Palade Laboratories Room 105, La Jolla, CA 92093-0741, E-mails: abharti{at}ucsd.edu, kpatra{at}ucsd.edu, and jvinetz{at}ucsd.edu. Raul Chuquiyauri, Instituto de Medicina Tropical Alexander von Humboldt, Universidad Peruana Cayetano Heredia, Morona 448-452, Iquitos, Peru, E-mail: raulharo{at}yahoo.com. Margaret Kosek and Robert H. Gilman, Department of International Health, Johns Hopkins Bloomberg School of Public Health, 615 North Wolfe Street, Room #W5515, Baltimore, MD 21205, E-mails: mkosek{at}jhsph.edu and rgilman{at}jhsph.edu. Alejandro Llanos-Cuentas, Instituto de Medicina Tropical Alexander von Humboldt, Universidad Peruana Cayetano Heredia, Avenida Honorio Delgado 430, Urbanización Ingenieria, San Martin de Porres, Lima 31, Peru, E-mail: allanos{at}upch.edu.pe.
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