| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
Malaria transmission and mortality rates remain unchanged in endemic countries lacking adequate health care and malaria control despite the use of preventive measures and treatments against malaria.1 A major obstacle to effective malaria control is the lack of affordable and accurate malaria diagnostics and treatment, which has led to misuse and abuse of anti-malarial drugs and the development of drug resistance in parasites.
Microscopic examination of blood smears, the conventional method for P. falciparum detection, is currently being augmented with antigen- and PCR-based rapid diagnostic tests (RDTs) for blood. However, inaccurate microscopic evaluation of blood smears have resulted in misdiagnoses and misclassification of malaria severity.2,3 Blood taboos and increased risk of accidental infections due to needle pricks continue to impact malaria diagnosis negatively. In non-specialized laboratories,4 microscopic evaluation of blood smears is slow and may lead to late diagnoses and treatment, which contributes to high mortality rates.5
Rapid diagnostic tests (RDTs) or "dipstick" are currently being used to detect antigens of Plasmodium species in blood or plasma to supplement microscopic evaluation of blood smears to manage tropical febrile disease.6 The benefits of this approach include the rapid turnaround time and the ease of use, which allows inexperienced laboratory or clinical staff to make on-the-spot diagnoses in the absence of visible parasites.6 However, issues associated with cultural objections to the collection of blood in communities with blood taboos7,8 and increased risk of needle injuries and disease transmission must be addressed.9
Saliva has been used in surveillance of vaccine-preventable diseases, such as measles, mumps, and rubella,10,11 and for individual diagnosis of HIV infection12 by detecting antibodies against the target pathogen. Although P. falciparum HRP II antigen has been detected in erythrocytes, serum, plasma, cerebrospinal fluid, and urine,13,14 detection of parasite antigens in saliva of P. falciparum-infected humans has not been reported. The goal of this pilot study was to test the possibility of detecting malaria parasite antigen in saliva in malaria patients. The hypothesis is that P. falciparum histidine-rich protein II (PfHRP II) is detectable in saliva in patients with symptomatic malaria.
The study was conducted at the Korle-Bu Teaching Hospital’s Child Health Department, Accra, Ghana, after ethical approval by Morehouse School of Medicine and University of Ghana Medical School. Randomly collected samples (plasma and saliva) from children (22 months to 16 years) reporting to the Child Health Department’s diagnostic laboratory were retrospectively analyzed for this study. Malaria positive cases were confirmed by thick film slides. Parasitemia was evaluated on the number of parasites per field (+, 1–10 parasites/100 fields, ++, > 10 parasites/100 fields, +++, 1–10 parasites/field, and ++++ > 10 parasites/field) and at least 100 fields/slide were examined to rule out any negative thick film slide. Thirty thick film positive children and 10 negative children were enrolled. Red blood cells (infected and uninfected) and plasma were separated using Vacutainer Cell Preparation Tubes (CPT) with Sodium Citrate (Becton Dickinson, USA). Saliva was collected in sterile containers and aliquoted into microcentrifuge tubes and stored at –20°C. Saliva samples were centrifuged for 3 min at 14,000 rpm and the supernatants were analyzed by ELISA. Both saliva and plasma samples from the same patient were analyzed on the same plate, date, and conditions for PfHRP II antigen levels using a Malaria Antigen ELISA kit (CELISA, Cellabs, Australia). This kit measures HRP II production during growth and multiplication15 of P. falciparum at a specificity of 96% and sensitivity of 98% in whole blood or plasma and can detect P. falciparum parasites at a limit of detection of 0.001%16; thus incubation periods with reagents were the same for plasma and saliva for the same patient. The plasma samples were tested at a 1:2 dilution and all samples were run in duplicates by ELISA. The incubation period for primary and secondary antibodies with the samples was 1 hr each in a humid chamber and 15 min for enzyme development (substrate) in the dark at room temperature. The minimum limit of detection (cut-off level) of the kit was determined according to manufacturer’s instructions.
Of the 30 children testing positive for blood smear, 16 (53%) had detectable PfHRP II antigens in their plasma (Table 1
). Thirteen (43%) patients of the 30 positive blood smears were PfHRP II positive for saliva samples (Table 1). All patients that were PfHRP II positive for saliva were also positive for plasma. Three patients (P006, P008, and P011) were PfHRP II positive in plasma but negative for saliva samples. Surprisingly, P006 had a mean OD reading (0.144) that is slightly below the cut-off level of 0.161 compared with the other 2 (P008 and P011) PfHRP II negative saliva. This observation suggests that P006 may have PfHRP II in the saliva that is undetectable in the kit used for this study. The 10 negative blood smears were also negative for PfHRP II antigen in both plasma and saliva. In our study the minimum limit of detection (cut-off level) was an OD reading of 0.161, which was determined according to the manufacturer’s instructions. In addition, all 13 saliva specimens had lower titers (OD, 0.166–0.427) of PfHRP II with a mean of 0.209 ± 0.07. The sensitivity of PfHRP II detection test for plasma was 53% and 43% for saliva whereas specificity was 100% for both specimens when compared with blood smears.
|
Malaria detection and epidemiological surveys in developing countries often require collection of blood samples from severely anemic children and communities with blood taboos. In central Africa, blood is considered an essential constituent of the "vital force" and an object of greed, "devoured" by sorcerers.7 Therefore, collection of blood specimens—regardless of the volume for definitive or confirmatory diagnosis—is poorly accepted.7 Thus, a non-invasive approach will greatly enhance cooperation of patients.
PCR methods have been used to detect malaria parasites in the blood.8,20 Although PCR-based methods are more sensitive and specific than existing techniques, the process is lengthy and requires specialized, costly equipment and reagents, as well as laboratory conditions that are not possible in the field.21 Sensitivity of detection in saliva was not enhanced in this study due to limitations of the commercially available kit used, which is designed to detect higher levels of PfHRP II in whole blood or plasma than is found in saliva. Therefore, development of a kit or test that is sensitive enough to detect lower levels of the antigen present in saliva could be a more appropriate approach to malaria diagnostics and in epidemiological surveys, thus, substituting blood samples with saliva specimens.
The detection of PfHRP II in saliva offers a practical alternative to PfHRP II detection in blood for malaria diagnosis and offers some distinct advantages over blood. Collection of saliva is non-invasive, simple, safe, stress free, painless, and can be done by individuals with limited training, including patients. It will not require blood cell lysis that diminishes HRP II antigen availability and detection. No special equipment is needed for collection and it allows for multiple or serial collections outside of the hospital.
Detecting parasite antigens in saliva to determine presence or absence of parasites could be valuable for communities with blood taboos and reduce compliance problems associated with collection of blood.22,23 Furthermore, it will provide a cost-effective approach for the screening of large populations in epidemiological surveys while being affordable, rapid, non-invasive, and safe for patients and technicians in resource-poor environments.
Received December 19, 2007. Accepted for publication February 15, 2008.
Acknowledgments: The authors acknowledge Clement Adu-Gyamfi and Opuni Asiedu for technical assistance and Chelsea Glass, Nathan McGinnis, and September Hesse of Morehouse School of Medicine for participation in sample analysis and the patients and their guardians for providing permission to use samples. The authors are grateful to the laboratory staff of the Korle-Bu Teaching Hospitals’ Child Health Department.
Financial support: This investigation received financial support from WHO/UNDP/TDR Collaborative Research Grant (A00524) and National Institutes of Health grant numbers NIH-RCMI (RR03034), NIH-NIGM-MBRS (SO6GM08248), and NIH-FIC (R21TW006804-01).
* Address correspondence to Jonathan K. Stiles, 720 Westview Dr. SW, Atlanta, GA 30310. E-mail: jstiles{at}msm.edu 
Authors’ addresses: Nana O. Wilson, Morehouse School of Medicine, Department of Microbiology, Biochemistry and Immunology, BMSB Room 350, 720 Westview Dr. SW, Atlanta, GA 30310, Tel: 404-742-1765, Fax: 404-752-1179, E-mail: nwilson{at}msm.edu. Andrew A. Adjei, University of Ghana Medical School, Department of Pathology Accra, Ghana, Tel: +233-20-813-5979, Fax: +233-21-668286, E-mail: andrewadjei50{at}hotmail.com. Winston Anderson, Howard University, Department of Biology, Just Hall, 415 College St. NW, Washington, DC 20059, Tel: 202-806-6933, E-mail: wanderson{at}howard.edu. Stella Baidoo, Korle-Bu Teaching Hospital, Department of Hematology, Child Health Laboratory, Accra, Ghana, Tel: +233-20-832-7836, E-mail: nanakosua2004{at}yahoo.co.uk. Jonathan K. Stiles, Morehouse School of Medicine, Department of Microbiology, Biochemistry and Immunology, BMSB Room 349D, 720 Westview Dr. SW, Atlanta, GA 30310, Tel: 404-742-1586, Fax: 404-752-1179, E-mail: jstiles{at}msm.edu
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
