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    Figure 1.

    A, Polymerase chain reaction (PCR) amplification products using Plasmodium genus-specific primers. Lanes 1 and 2, DNA extracted from whole blood and serum of a P. vivax malaria patient; lane 3, DNA extracted from P. falciparum (3D7 strain) in vitro culture; lanes 4 and 5, DNA extracted from serum of patient with P. falciparum malaria and mixed infection, respectively; lane 6, DNA extracted from serum of a malaria-negative control; lane 7, DNA template from a negative control. B, PCR amplification for the small subunit ribosomal RNA (ss rRNA) gene of P. vivax. Lane 1, DNA extracted from whole blood of patient with P. vivax malaria; lanes 2 and 3, DNA extracted from serum of patient with P. vivax and mixed infection, respectively; lane 4, DNA extracted from serum of a malaria-negative control; lane 5, DNA template from a negative control. C, PCR amplification for the ss rRNA gene of P. falciparum. Lane 1, DNA extracted from P. falciparum (3D7 strain) in vitro culture; lanes 2 and 3, DNA extracted from serum of patient with P. falciparum and mixed infection, respectively; lane 4, DNA extracted from serum of a malaria-negative control; lane 5, DNA template from a negative control.

  • 1

    Warhurst DC, Williams JE, 1996. ACP Broadsheet no 148. July 1996. Laboratory diagnosis of malaria. J Clin Pathol 49 :533–538.

  • 2

    Jonkman A, Chibwe RA, Khoromana CO, Liabunya UL, Chaponda ME, Kandiero GE, Molyneux ME, Taylor TE, 1995. Cost-saving through microscopy-based versus presumptive diagnosis of malaria in adult outpatients in Malawi. Bull World Health Organ 73 :223–227.

    • Search Google Scholar
    • Export Citation
  • 3

    Milne LM, Kyi MS, Chiodini PL, Warhurst DC, 1994. Accuracy of routine laboratory diagnosis of malaria in the United Kingdom. J Clin Pathol 47 :740–742.

    • Search Google Scholar
    • Export Citation
  • 4

    Roshanravan B, Kari E, Gilman RH, Cabrera L, Lee E, Metcalfe J, Calderon M, Lescano AG, Montenegro-James S, Calampa C, Vinetz JM, 2003. Endemic malaria in the Peruvian Amazon region of Iquitos. Am J Trop Med Hyg 69 :45–52.

    • Search Google Scholar
    • Export Citation
  • 5

    Moody A, 2002. Rapid diagnostic tests for malaria parasites. Clin Microbiol Rev 15 :66–78.

  • 6

    Whitworth JA, Hewitt KA, 2005. Effect of malaria on HIV-1 progression and transmission. Lancet 365 :196–197.

  • 7

    Bharti AR, Chuquiyauri R, Brouwer KC, Stancil J, Lin J, Llanos-Cuentas A, Vinetz JM, 2006. Experimental infection of the neotropical malaria vector Anopheles darlingi by human patient-derived Plasmodium vivax in the Peruvian Amazon. Am J Trop Med Hyg 75 :610–616.

    • Search Google Scholar
    • Export Citation
  • 8

    Snounou G, Viriyakosol S, Zhu XP, Jarra W, Pinheiro L, do Rosario VE, Thaithong S, Brown KN, 1993. High sensitivity of detection of human malaria parasites by the use of nested polymerase chain reaction. Mol Biochem Parasitol 61 :315–320.

    • Search Google Scholar
    • Export Citation
  • 9

    Shi SR, Datar R, Liu C, Wu L, Zhang Z, Cote RJ, Taylor CR, 2004. DNA extraction from archival formalin-fixed, paraffin-embedded tissues: heat-induced retrieval in alkaline solution. Histochem Cell Biol 122 :211–218.

    • Search Google Scholar
    • Export Citation
  • 10

    Gal S, Fidler C, Turner S, Lo YM, Roberts DJ, Wainscoat JS, 2001. Detection of Plasmodium falciparum DNA in plasma. Ann N Y Acad Sci 945 :234–238.

    • Search Google Scholar
    • Export Citation
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Polymerase Chain Reaction Detection of Plasmodium vivax and Plasmodium falciparum DNA from Stored Serum Samples: Implications for Retrospective Diagnosis of Malaria

Ajay R. BhartiDivision of Infectious Diseases, Department of Medicine, University of California San Diego, La Jolla, California; Instituto de Medicina Tropical Alexander von Humboldt, Universidad Peruana Cayetano Heredia, Lima, Peru; Department of International Health, Johns Hopkins Bloomberg School of Public Health, Baltimore, Maryland

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Kailash P. PatraDivision of Infectious Diseases, Department of Medicine, University of California San Diego, La Jolla, California; Instituto de Medicina Tropical Alexander von Humboldt, Universidad Peruana Cayetano Heredia, Lima, Peru; Department of International Health, Johns Hopkins Bloomberg School of Public Health, Baltimore, Maryland

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Raul ChuquiyauriDivision of Infectious Diseases, Department of Medicine, University of California San Diego, La Jolla, California; Instituto de Medicina Tropical Alexander von Humboldt, Universidad Peruana Cayetano Heredia, Lima, Peru; Department of International Health, Johns Hopkins Bloomberg School of Public Health, Baltimore, Maryland

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Margaret KosekDivision of Infectious Diseases, Department of Medicine, University of California San Diego, La Jolla, California; Instituto de Medicina Tropical Alexander von Humboldt, Universidad Peruana Cayetano Heredia, Lima, Peru; Department of International Health, Johns Hopkins Bloomberg School of Public Health, Baltimore, Maryland

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Robert H. GilmanDivision of Infectious Diseases, Department of Medicine, University of California San Diego, La Jolla, California; Instituto de Medicina Tropical Alexander von Humboldt, Universidad Peruana Cayetano Heredia, Lima, Peru; Department of International Health, Johns Hopkins Bloomberg School of Public Health, Baltimore, Maryland

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Alejandro Llanos-CuentasDivision of Infectious Diseases, Department of Medicine, University of California San Diego, La Jolla, California; Instituto de Medicina Tropical Alexander von Humboldt, Universidad Peruana Cayetano Heredia, Lima, Peru; Department of International Health, Johns Hopkins Bloomberg School of Public Health, Baltimore, Maryland

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Joseph M. VinetzDivision of Infectious Diseases, Department of Medicine, University of California San Diego, La Jolla, California; Instituto de Medicina Tropical Alexander von Humboldt, Universidad Peruana Cayetano Heredia, Lima, Peru; Department of International Health, Johns Hopkins Bloomberg School of Public Health, Baltimore, Maryland

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Polymerase chain reaction (PCR) detection of Plasmodium DNA is highly sensitive in diagnosing malaria. The specimen of choice for this assay has been whole blood samples from malaria patients. To retrospectively determine malaria infection rates in populations or cohorts for whom stored serum samples are available, we determined the ability of a nested PCR assay to detect Plasmodium DNA in stored serum samples. The PCR result was positive in 20 of 23 serum samples from patients with microscopy-confirmed malaria and negative in 8 of 8 healthy controls, resulting in a sensitivity of 87% and specificity of 100%. In all positive samples, species were correctly identified by PCR except for one case where a mixed infection was detected. The PCR is able to detect Plasmodium DNA in serum samples frozen up to 2.5 years and has the potential for the retrospective identification of malaria parasitemia in patient cohorts to determine potential interactions of malaria and other diseases such as human immunodeficiency virus/acquired immunodeficiency syndrome.

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.

Several important findings emerge from this study. First, both P. vivax and P. falciparum DNA are PCR-amplifiable from serum stored for up to approximately 30 months. Second, PCR can differentiate P. vivax and P. falciparum DNA from serum samples. Third, serum from patients with mixed infections can be successfully diagnosed by PCR. These findings are particularly important because P. vivax has lower levels of parasitemia than P. falciparum, yet PCR on serum appears to be an effective method to detect P. vivax.

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.

Table 1

PCR detection of Plasmodium DNA using two-year-old stored serum samples from microscopy-confirmed malaria patients*

Sample no.Field smear resultPCR resultHuman p53 resultNo. of days stored
* PCR = polymerase chain reaction; Pf = Plasmodium falciparum; Pv = P. vivax.
† Field smears results reported from the Ministry of Health health post. This sample was found to be negative on microscopy carried out by independent microscopists in the study laboratory.
1Pf++Pf+956
2Pv++†+955
3Pv+Pv+946
4Pv+++Pv+946
5Pv+++942
6Pv++Pv+942
7Pv+++942
8Pv++940
9Pv++Pv+940
10Pv++Pv+939
11Pf++Pf+935
12Pv<1+Pv+935
13Pv++Pv+932
14Pv++Pv+932
15Pv++Pv+932
16Pv+++Pv+931
17Pv++Pv+931
18Pv++Pv+928
19Pv++Pv+928
20Pv+Pv+927
21Pv+Pv+926
22Pv<1+Pv+923
23Pf+Pv, Pf+920
24Pv++Pv+920
Figure 1.
Figure 1.

A, Polymerase chain reaction (PCR) amplification products using Plasmodium genus-specific primers. Lanes 1 and 2, DNA extracted from whole blood and serum of a P. vivax malaria patient; lane 3, DNA extracted from P. falciparum (3D7 strain) in vitro culture; lanes 4 and 5, DNA extracted from serum of patient with P. falciparum malaria and mixed infection, respectively; lane 6, DNA extracted from serum of a malaria-negative control; lane 7, DNA template from a negative control. B, PCR amplification for the small subunit ribosomal RNA (ss rRNA) gene of P. vivax. Lane 1, DNA extracted from whole blood of patient with P. vivax malaria; lanes 2 and 3, DNA extracted from serum of patient with P. vivax and mixed infection, respectively; lane 4, DNA extracted from serum of a malaria-negative control; lane 5, DNA template from a negative control. C, PCR amplification for the ss rRNA gene of P. falciparum. Lane 1, DNA extracted from P. falciparum (3D7 strain) in vitro culture; lanes 2 and 3, DNA extracted from serum of patient with P. falciparum and mixed infection, respectively; lane 4, DNA extracted from serum of a malaria-negative control; lane 5, DNA template from a negative control.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 77, 3; 10.4269/ajtmh.2007.77.444

*

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@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@ucsd.edu, kpatra@ucsd.edu, and jvinetz@ucsd.edu. Raul Chuquiyauri, Instituto de Medicina Tropical Alexander von Humboldt, Universidad Peruana Cayetano Heredia, Morona 448-452, Iquitos, Peru, E-mail: raulharo@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@jhsph.edu and rgilman@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@upch.edu.pe.

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).

REFERENCES

  • 1

    Warhurst DC, Williams JE, 1996. ACP Broadsheet no 148. July 1996. Laboratory diagnosis of malaria. J Clin Pathol 49 :533–538.

  • 2

    Jonkman A, Chibwe RA, Khoromana CO, Liabunya UL, Chaponda ME, Kandiero GE, Molyneux ME, Taylor TE, 1995. Cost-saving through microscopy-based versus presumptive diagnosis of malaria in adult outpatients in Malawi. Bull World Health Organ 73 :223–227.

    • Search Google Scholar
    • Export Citation
  • 3

    Milne LM, Kyi MS, Chiodini PL, Warhurst DC, 1994. Accuracy of routine laboratory diagnosis of malaria in the United Kingdom. J Clin Pathol 47 :740–742.

    • Search Google Scholar
    • Export Citation
  • 4

    Roshanravan B, Kari E, Gilman RH, Cabrera L, Lee E, Metcalfe J, Calderon M, Lescano AG, Montenegro-James S, Calampa C, Vinetz JM, 2003. Endemic malaria in the Peruvian Amazon region of Iquitos. Am J Trop Med Hyg 69 :45–52.

    • Search Google Scholar
    • Export Citation
  • 5

    Moody A, 2002. Rapid diagnostic tests for malaria parasites. Clin Microbiol Rev 15 :66–78.

  • 6

    Whitworth JA, Hewitt KA, 2005. Effect of malaria on HIV-1 progression and transmission. Lancet 365 :196–197.

  • 7

    Bharti AR, Chuquiyauri R, Brouwer KC, Stancil J, Lin J, Llanos-Cuentas A, Vinetz JM, 2006. Experimental infection of the neotropical malaria vector Anopheles darlingi by human patient-derived Plasmodium vivax in the Peruvian Amazon. Am J Trop Med Hyg 75 :610–616.

    • Search Google Scholar
    • Export Citation
  • 8

    Snounou G, Viriyakosol S, Zhu XP, Jarra W, Pinheiro L, do Rosario VE, Thaithong S, Brown KN, 1993. High sensitivity of detection of human malaria parasites by the use of nested polymerase chain reaction. Mol Biochem Parasitol 61 :315–320.

    • Search Google Scholar
    • Export Citation
  • 9

    Shi SR, Datar R, Liu C, Wu L, Zhang Z, Cote RJ, Taylor CR, 2004. DNA extraction from archival formalin-fixed, paraffin-embedded tissues: heat-induced retrieval in alkaline solution. Histochem Cell Biol 122 :211–218.

    • Search Google Scholar
    • Export Citation
  • 10

    Gal S, Fidler C, Turner S, Lo YM, Roberts DJ, Wainscoat JS, 2001. Detection of Plasmodium falciparum DNA in plasma. Ann N Y Acad Sci 945 :234–238.

    • Search Google Scholar
    • Export Citation
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