INTRODUCTION
Malaria is a parasitic disease in approximately 100 countries in tropical and subtropical regions. The World Health Organization estimates that 1.5 billion people are at risk for developing this disease. Approximately 500 million clinical cases of malaria and one million deaths from this disease occur each year and most are caused by Plasmodium falciparum.1,2 Most cases of malaria caused by P. vivax, another malaria parasite of epidemiologic importance, occur in the Middle East, Asia, the western Pacific region, and Central and South America.3
In contrast to other infectious diseases, acquisition of immunity to malaria is a slow and complex process. Sterile immunity to malaria is never achieved, although in areas highly endemic for this disease, individuals repeatedly exposed to malaria infections progressively develop clinical immunity. This results in them being asymptomatic even if they remain infected.4 Additionally, immune mechanisms activated by the infection are able to block parasite transmission from the patient to the mosquito vector in what is called transmission-blocking immunity.5 This immunity may play an important role in decreasing the spread of malaria, particularly in areas of low transmission intensity.6
The overall protective immunity in malaria is mediated by a combination of antibodies, T cells, cytokines, and several other mechanisms that act at different levels of the parasite life cycle; some of these mechanisms appear to be more relevant for blocking of certain parasite stages.7,8 Sera from individuals naturally exposed to P. vivax and having acute infections contain antibodies to gametocytes capable of blocking the parasite fertilization process and the ookinete invasion of the mosquito midgut cells, thus preventing further parasite development.6,9 However, this blocking process is complex and appears to be dependent on antibody concentration. Approximately half of the sera collected in Kataragama, a P. vivax–endemic region of Sri Lanka, and analyzed for their transmission-blocking activity suppressed gametocyte infectivity; but 12% showed enhancement of malarial transmission.9
In the Western Hemisphere, P. vivax is present in 20 countries from the southern coast of Mexico to the Brazilian forests, where it is the most prevalent species.3 Difficulties in establishing laboratory colonies of susceptible Anopheles mosquitoes, as well as the limited availability of gametocyte-containing blood samples, have resulted in limited numbers of studies on P. vivax transmission-blocking immunity in this region. Studies by Ramsey and others have shown high transmission-blocking activity in individuals who had more than one P. vivax clinical attack in malaria-endemic regions of Mexico,10 but there are no reports of transmission-blocking studies in the other regions of the Americas.
We have benefited from access to patients infected with P. vivax in a malaria-endemic area of Colombia and availability of a well-established Anopheles albimanus colony to study the prevalence of transmission-blocking responses of patient sera on autologous parasites.
MATERIALS AND METHODS
Study area and population.
The study was conducted on the Pacific coast of Colombia where both P. vivax and P. falciparum are transmitted in approximately similar proportions throughout the year. In this region, there are peaks in malaria transmission between April and May and between September and October (annual parasite incidence = 6–10 confirmed malaria cases/1,000 people/year).11 We selected individuals living in urban and rural communities from Buenaventura, the main seaport in this region (Figure 1). It is located in a tropical rain forest area with an annual rainfall ranging from 6,000 to 9,000 mm, a relative humidity of approximately 85%, and a mean temperature of approximately 28°C. These communities in this region are composed of mixed ethnic groups: 70% of African origin and 30% of Spanish-Amerindian (Mestizo) origin.12 Most are involved in agricultural logging, fishing, and mining activities.
The study protocol was reviewed and approved by the Institutional Review Board of the Universidad del Valle and the International Clinical Studies Review Committee of the National Institute of Allergy and Infectious Diseases, National Institutes of Health (Bethesda, MD). Individuals were recruited from malaria diagnostic outpatient clinics in Buenaventura. They were eligible if they were ≥ 18 years old, had a mono-infection with P. vivax as determined by thick or thin blood smears, and gave written informed consent.
Participants responded to a questionnaire that included demographic and clinical information about current and previous malaria episodes. Samples of blood were obtained from P. vivax–infected patients and donors were immediately treated with curative doses of antimalarial drugs according to the standard therapeutic protocol recommended by the Colombian Ministry of Social Protection: primaquine, 15 mg/day for 14 days, plus chloroquine phosphate, 600 mg on the first day and 450 mg/day for three days. Parasitemia and gametocyte density were determined from a thick blood smear by light microcopy with an oil-immersion lens at a magnification of 1,000×. Asexual- and sexual-stage parasite densities were counted per 100 leukocytes and expressed as either parasites or gametocytes per microliter assuming a leukocyte count of 8,000/μL.13
Blood samples.
Seventeen milliliters of whole blood was collected by venipuncture into dry Vacutainer® tubes (Becton Dickinson, Franklin Lakes, NJ) from each P. vivax–infected patient. Serum samples were obtained from blood by centrifugation at 800 × g for 10 minutes, and infected erythrocytes were collected into heparinized Vacutainer® tubes and immediately fractionated by centrifugation at 500 × g for 5 minutes at 37°C. Infected red blood cells were suspended in either patient (autologous) or normal human AB serum and used within one hour to feed mosquitoes by a membrane feeding assay (MFA). Serum aliquots were stored at −70°C and used to detect parasite-specific antibodies by an immunofluorescent antibody test (IFAT). Normal human serum consisted of a pool of human AB serum obtained from a Red Cross Blood Bank, aliquoted, and stored at −70°C.
Membrane feeding assays.
Parasite transmission-blocking activity of human serum samples was assessed by an MFA using An. albimanus mosquitoes reared in the Entomology Unit in Buenaventura. Two 400-μL aliquots of whole blood were centrifuged and their corresponding P. vivax–infected red blood cell fractions were washed with two volumes of serum-free RPMI 1640 medium (Sigma, St Louis, MO); blood samples were maintained at 37°C to prevent exflagellation prior to the MFA. One of these fractions was reconstituted with 200 μL of autologous serum and the other with the same volume of normal AB human serum (control) at a 50% hematocrit. Serum samples were not heat-inactivated. Reconstituted blood samples were immediately placed in the artificial membrane feeding device that was temperature regulated (37°C) and offered to batches of 110 laboratory-reared, 3–4-day-old female mosquitoes. After 30 minutes of feeding, unfed mosquitoes were removed from the cages. Seven to eight days after feeding, 40 mosquitoes were dissected and midguts were stained with 2% mercurochrome. The number of oocysts per mosquito midgut was counted for autologous and control serum. Uninfected control batches and their corresponding autologous assays were not included in the analysis.
Immunofluorescence antibody test.
Specific antibody titers to blood-stage parasites were determined by an IFAT using air-dried smears of P. vivax parasites obtained from infected patients containing both sexual and asexual blood forms.6 Sera samples diluted in phosphate-buffered saline (PBS) were tested in two-fold serial dilutions from 1:100 to 1:3,200 and incubated for 30 minutes at room temperature (20–24°C). Slides were washed with PBS 1× and a 1:100 dilution of goat anti-human IgG fluorescein conjugate (Jackson Immuno-Research, West Grove, PA) in Evans blue solution (diluted 1:5,000 in PBS) was added and incubated for 30 minutes. Slides were then washed, overlaid with glass cover slips using 30% glycerol, examined by ultraviolet microscopy, and scored for the presence or absence of fluorescent parasites in the test samples versus control sera.
Potency of transmission-blocking activity.
The inhibitory potency of sera on oocyst development was assessed in the MFA by using serial dilutions of test sera. Sixteen serum samples were tested undiluted and at two-fold dilutions (up to 1:64) for transmission-blocking activity on autologous P. vivax parasites.
Statistical analysis.
A batch of mosquitoes was tested using two outcomes of interest: the proportion of infected mosquitoes and the arithmetic mean of oocyst counts. We used the Wilcoxon signed-rank test to assess if differences between paired estimates made in batches fed with autologous and control samples were statistically significant.
Differences between the two outcomes of interest were also used to estimate the magnitude of the transmission-blocking activity. Specifically, both the percentage reduction in the proportion of infected mosquitoes and the percentage reduction in mean oocyst counts/mosquito were estimated using the formula: [(Xc − Xa)/Xc)] × 100, where X is the proportion or arithmetic mean in control (c) and autologous (a) sera.14 The percentage reduction of infected mosquitoes and oocyst counts were both classified according to the following categories: < 0% (enhancing), 0–49%, 50–89%, and 90–100%.
The magnitude of transmission-blocking activity was determined using the median of the percentage reduction because of the extremely skewed distribution and percentage of individuals with transmission-blocking activity > 90%. Association of transmission-blocking activity with demographic and clinical variables was evaluated by stratification.
To evaluate the correlation between the intensity of the immune response and transmission-blocking activity, four levels of antibody titers were defined (≤ 1:100, 1:200, 1:400, and ≥ 1:800 dilutions) and their corresponding values of transmission-blocking activity were analyzed using the Spearman’s correlation coefficient.
Factors potentially associated with transmission-blocking activity > 90% were evaluated using a multiple logistic regression model to control for confounders. We included in the model demographic and clinical variables and previous malarial episodes. All statistical inferences were made using Stata 8.0 software (Stata Corporation, College Station, TX).
RESULTS
We performed 114 paired MFAs using blood from patients acutely infected with P. vivax. Among them, 88 (77.2%) of the control batches were able to infect mosquitoes and were used for subsequent analysis. No statistical differences were found between autologous and control batches in the average numbers of fed and surviving mosquitoes. A total of 3,520 mosquitoes were dissected from each of these two groups. From the 88 paired mosquito batches used to assess the transmission-blocking activity, 43.3% of mosquitoes fed with parasites in autologous sera and 66.6% of mosquitoes in control batches had oocysts (P < 0.001, by Wilcoxon signed-ranked test).
Figure 2 shows the distribution of arithmetic mean oocyst counts for both autologous and control groups. The mean oocyst count was 4.6/mosquito in autologous batches and 11.1/mosquito in control batches (P < 0.001, by Wilcoxon signed-ranked test). In control batches, the mean oocyst count varied according to the febrile status of the patients. The mean number of oocysts/mosquito in the mosquitoes fed with parasites from febrile patients was 7.3 and the percentage of infected mosquitoes was 57.9%. In patients without fever, the mean oocysts/mosquito was 12.8 and the percentage of infected mosquitoes was 70.7%.
Transmission-blocking activity, as measured by the percentage reduction in the proportion of infected mosquitoes and by the percentage reduction in the mean oocyst counts/ mosquito, is summarized in Table 1. High levels of transmission-blocking activity (90–100% reduction) were observed in 20 (22.7%) of 88 patients and 32 (36.4%) of 88 patients for reductions in infected mosquitoes and oocysts, respectively. Intermediate levels of transmission-blocking activity (50–89%) were more frequent for oocyst reduction (29%) than for reduction in the number of infected mosquitoes (17.0%). Conversely, in the range of low levels of transmission-blocking activity (0–49%), reductions in the percentage of infected mosquitoes (42.1%) were more frequently observed than reductions in mean oocyst counts (17%).
Univariate analysis of factors associated with transmission-blocking activity is summarized in Table 2. Greater transmission-blocking activity was consistently observed in 30–40-year-old non–Afro-Colombian individuals in comparison with Mestizos, with rural residence or occupation, prior malaria attacks, and fever (body temperature > 38°C) associated with increased antibody titers. The number of previous malaria episodes recorded during the informed clinical history ranged from 1 to 20. Multiple logistic regression analyses of factors associated with transmission-blocking activity ≥ 90% showed different results for positive mosquitoes and oocyst count reductions. After adjusting for demographic and clinical variables, a temperature > 38°C was the only factor significantly associated with a ≥ 90% relative reduction in the percentage of infected mosquitoes (odds ratio [OR] = 3.8, 95% confidence interval [CI] = 1.2–11.4). Factors associated with a comparable reduction in oocysts/mosquito were Afro-Colombian ethnicity (OR = 0.16, 95% CI = 0.1–0.5), previous malaria episodes (OR = 4.6, 95% CI = 1.1–19.1), and antibody titers ≥ 1:800 compared with those ≤ 1:100 (OR = 5.4, 95% CI = 1.7–17.5).
Antibody titers in test sera were determined by IFAT analysis using sexual and asexual stage parasites, and the results were compared with transmission-blocking activity of the same sera as defined by a reduction in oocysts/mosquito. Box plots of transmission-blocking activity according to serum antibody titers are shown in Figure 3. The medians of transmission-blocking activity for antibody titers < 1:100, 1: 200, 1:400, and >1:800 were 18.8%, 66.8%, 76.9%, and 94.9%, respectively. A statistically significant correlation between these variables supports the strong dependence of transmission-blocking activity on increased antibody titers (Spearman’s correlation coefficient = 0.4, P = 0.0001).
For the assessment of transmission-blocking potency, an additional 16 samples were tested at different concentrations. Results indicated that blocking effects of all sera rapidly decreased with dilutions > 1:4.
DISCUSSION
This study shows that sera from Colombian patients with acute P. vivax malarial infections are able to efficiently block parasite transmission to An. albimanus mosquitoes. Approximately two-thirds of the studied individuals displayed significant transmission-blocking activity (≥ 50%) and half had > 90% inhibition as determined by the reduction in mean oocysts/mosquito. Of those sera, 15.9% totally blocked the development of mosquito infection, and the remaining samples all showed a decrease in the number of infected mosquitoes per batch. Likewise, more than 90% relative reduction in the proportion of infected mosquitoes was found more frequently in patients with fever (> 38°C) at the time of the test.
The results of this study are the first from a malaria-endemic region of South America. Although not completely comparable with results from other regions, they confirm the presence of transmission-blocking activity in areas endemic for P. vivax and are within ranges described in other regions. In Sri Lanka, the intensity of transmission-blocking activity is dependent on malaria endemicity.6,9 In Kataragama, Sri Lanka, a malaria-endemic region with an annual parasite incidence (API; the number of confirmed malaria cases/1,000 people/year) of 9.3 for P. vivax,15 only 22% of the studied sera induced total transmission-blocking activity.9 In Colombo, Sri Lanka, an area not endemic for malaria, area, complete transmission-blocking activity was observed in two-thirds of patients acutely infected with P. vivax.6 Consistent with these findings, the API in Buenaventura was 8.0, similar to that in Kataragama, as reported by the Colombian Ministry of Health; however, transmission-blocking activity was higher in our study (36%). In Mexico, high transmission-blocking activity was observed in individuals with malaria within the past six months, as determined by a reduction of sporozoite production by mosquitoes.10 In our study no correlation was observed between transmission-blocking activity and previous malaria episodes.
Although the mechanism for transmission-blocking activity in the present study was not determined, it directly correlated with antibody titers against P. vivax as determined by immunofluorescence. The role of monoclonal or polyclonal antibodies in suppression of parasite infectivity has been previously documented.15 In the studies carried out in Sri Lanka and Mexico, sera from individuals with low antibody titers produced less blocking activity and, in some cases, enhancement of transmission.9,10,16 This correlation is also consistent with a reduction of transmission-blocking activity when inhibitory sera were diluted and with previous studies.6,9,15 Moreover, in a recent Phase I clinical trial using the ookinete surface protein P. vivax Pvs25H, a correlation between antibody concentration and transmission-blocking activity was observed.17
Sera from the Afro-Colombian population studied induced significantly less transmission-blocking activity than sera from other ethnic groups, although no statistical differences could be identified with previous exposure to malaria, antibody titers, or body temperature. Afro-Colombians are a native population of the Pacific coast, whereas Mestizos are more recently established in this area. Similar observations have been reported and explained by continuous exposure to malaria infections that down-regulates the immune response against sexual-stage antigens.9 In our study, approximately 40% of sera from patients with fever (temperature > 38°C) produced a relative reduction in the proportion of infected mosquitoes ≥ 90%, while only 15.0% of those sera from patients without fever showed transmission-blocking activity ≥ 90%. Similarly, previous studies also found that paroxysms associated with fever during P. vivax malaria episodes induced a decrease in mosquito infectivity, which was probably due to decreased gametocyte viability.18,19 Our results are consistent with this hypothesis because we found that the percentage of infected mosquitoes in febrile patients was lower than the percentage found in patients without fever. This has been proposed to be a pro-inflammatory immune response mediated by several factors such as tumor necrosis factor-α and other toxic factors that kill gametocytes.20,21 Further studies on the characterization of these sera are being performed to clarify the mechanisms involved in transmission-blocking activity.
The overall blockage of parasite transmission to mosquitoes is a complex, multi-factorial process that involves several immune mechanisms and other host and parasite factors.22–24 This study has confirmed previous findings from other geographic regions regarding the potential of transmission-blocking immune responses in the decrease of parasite dissemination. It also provides the basis for a more detailed analysis of the mechanisms involved in transmission-blocking activity and their potential immune targets.
Transmission-blocking (TB) activity as a reduction in percentage of infected mosquitoes and mean oocyst counts/mosquito among individuals acutely infected with Plasmodium vivax in Buenaventura, Colombia
| No. (%) of patients in each TB category | ||||
|---|---|---|---|---|
| TB categories* | Infected mosquitoes | Oocyst/mosquito | ||
| % Reduction | n† | % | n† | % |
| * Categories of transmission-blocking activity to classify the percentage reduction of infected mosquitoes and oocyst counts. | ||||
| † Total number of control positive batches of mosquitoes of the 114 paired membrane feeding assays performed. | ||||
| 90–100 | 20 | 23 | 32 | 36 |
| 50–89 | 15 | 17 | 26 | 30 |
| 0–49 | 37 | 42 | 15 | 17 |
| < 0 (enhancing) | 16 | 18 | 15 | 17 |
| Total | 88 | 100 | 88 | 100 |
Demographic and clinical factors of infected individuals according to transmission-blocking (TB) activity
| % Infected mosquitoes | % Oocyst count | ||||||||
|---|---|---|---|---|---|---|---|---|---|
| Variables | n* | Median† | P | ≥ 90‡ | P | Median† | P | ≥ 90‡ | P |
| * Total number of participants in 88; some variables had missing values. | |||||||||
| † Median of transmission-blocking activity as % reduction in either % infected mosquitoes or oocyst counts/mosquito and P values determined by Kruskal-Wallis test. | |||||||||
| ‡ Percentage of individuals with > 90% reduction in either % of infected mosquitoes or oocyst count/mosquito P values determined by Fisher exact test. | |||||||||
| § Asexual- and sexual-stage parasite densities per 100 leukocytes. Parasite and gametocytes results are expressed in microliters assuming a leukocyte count of 8,000/μL. | |||||||||
| Age, years | |||||||||
| < 30 | 40 | 34.6 | 20.0 | 80.4 | 40.0 | ||||
| 30–39 | 14 | 52.4 | 35.7 | 87.5 | 42.9 | ||||
| ≥ 40 | 30 | 20.0 | 20.0 | 53.1 | 30.0 | ||||
| 0.23 | 0.44 | 0.17 | 0.62 | ||||||
| Ethnicity | |||||||||
| Afro-Colombians | 32 | 16.4 | 12.5 | 49.3 | 15.6 | ||||
| Mestizos | 56 | 52.0 | 27.8 | 88.6 | 48.2 | ||||
| 0.01 | 0.11 | < 0.01 | < 0.01 | ||||||
| Residence | |||||||||
| Urban | 48 | 29.0 | 16.7 | 67.4 | 31.3 | ||||
| Rural | 40 | 37.0 | 30.0 | 81.7 | 42.5 | ||||
| 0.95 | 0.20 | 0.28 | 0.37 | ||||||
| Place of work | |||||||||
| Urban | 45 | 31.4 | 17.8 | 64.3 | 31.1 | ||||
| Rural | 43 | 33.3 | 27.9 | 83.1 | 41.9 | ||||
| 0.86 | 0.31 | 0.20 | 0.38 | ||||||
| Previous malaria episodes | |||||||||
| None | 17 | 25.0 | 11.8 | 52.8 | 23.5 | ||||
| > 1 | 69 | 33.3 | 24.6 | 77.7 | 39.1 | ||||
| 0.41 | 0.34 | 0.17 | 0.27 | ||||||
| Days with symptoms | |||||||||
| ≤ 3 | 44 | 38.3 | 22.7 | 79.7 | 38.6 | ||||
| 4–7 | 28 | 16.9 | 17.9 | 61.9 | 32.1 | ||||
| > 7 | 14 | 33.3 | 28.6 | 75.6 | 35.7 | ||||
| 0.68 | 0.69 | 0.39 | 0.91 | ||||||
| Parasitemia/μL§ | |||||||||
| < 10,000 | 49 | 33.3 | 22.5 | 79.2 | 36.7 | ||||
| ≥ 10,000 | 39 | 27.3 | 23.1 | 61.4 | 35.9 | ||||
| 0.82 | 1.00 | 0.63 | 1.00 | ||||||
| Gametocytemia/μL§ | |||||||||
| < 3,000 | 45 | 33.3 | 20.0 | 77.7 | 35.6 | ||||
| ≥ 3,000 | 43 | 27.3 | 25.6 | 71.2 | 37.2 | ||||
| 0.76 | 0.62 | 0.71 | 1.00 | ||||||
| Fever (> 38.0°C) | |||||||||
| No | 60 | 29.3 | 15.0 | 76.1 | 31.7 | ||||
| Yes | 22 | 42.4 | 40.9 | 71.1 | 45.5 | ||||
| 0.48 | 0.02 | 0.77 | 0.30 | ||||||
| Antibody titers | |||||||||
| ≤ 1:100 | 21 | 17.6 | 14.3 | 18.8 | 19.1 | ||||
| 1:200 | 16 | 24.4 | 12.5 | 66.8 | 18.8 | ||||
| 1:400 | 26 | 25.8 | 23.1 | 76.9 | 38.5 | ||||
| ≥ 1:800 | 25 | 77.8 | 36.0 | 94.9 | 60.0 | ||||
| 0.04 | 0.26 | < 0.01 | 0.01 | ||||||
Map of the Colombian Pacific coast showing Buenaventura and surrounding rural communities where blood samples were collected.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 73, 5_suppl; 10.4269/ajtmh.2005.73.5_suppl.0730038
Map of the Colombian Pacific coast showing Buenaventura and surrounding rural communities where blood samples were collected.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 73, 5_suppl; 10.4269/ajtmh.2005.73.5_suppl.0730038
Map of the Colombian Pacific coast showing Buenaventura and surrounding rural communities where blood samples were collected.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 73, 5_suppl; 10.4269/ajtmh.2005.73.5_suppl.0730038
Box plot showing the upper and lower quartiles of mean oocyst counts per mosquito. The central line is the median. The ends of the whiskers are 1.5 interquartile distances above the upper quartile and below the lower quartile.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 73, 5_suppl; 10.4269/ajtmh.2005.73.5_suppl.0730038
Box plot showing the upper and lower quartiles of mean oocyst counts per mosquito. The central line is the median. The ends of the whiskers are 1.5 interquartile distances above the upper quartile and below the lower quartile.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 73, 5_suppl; 10.4269/ajtmh.2005.73.5_suppl.0730038
Box plot showing the upper and lower quartiles of mean oocyst counts per mosquito. The central line is the median. The ends of the whiskers are 1.5 interquartile distances above the upper quartile and below the lower quartile.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 73, 5_suppl; 10.4269/ajtmh.2005.73.5_suppl.0730038
Box plot showing the upper and lower quartiles of transmission blocking activity as percent reduction in mean oocyst counts per mosquito. The central line is the median. The ends of the whiskers are 1.5 interquartile distances above the upper quartile and below the lower quartile. Samples are stratified by antibody titers to Plasmodium vivax in an immunofluorescent antibody test.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 73, 5_suppl; 10.4269/ajtmh.2005.73.5_suppl.0730038
Box plot showing the upper and lower quartiles of transmission blocking activity as percent reduction in mean oocyst counts per mosquito. The central line is the median. The ends of the whiskers are 1.5 interquartile distances above the upper quartile and below the lower quartile. Samples are stratified by antibody titers to Plasmodium vivax in an immunofluorescent antibody test.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 73, 5_suppl; 10.4269/ajtmh.2005.73.5_suppl.0730038
Box plot showing the upper and lower quartiles of transmission blocking activity as percent reduction in mean oocyst counts per mosquito. The central line is the median. The ends of the whiskers are 1.5 interquartile distances above the upper quartile and below the lower quartile. Samples are stratified by antibody titers to Plasmodium vivax in an immunofluorescent antibody test.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 73, 5_suppl; 10.4269/ajtmh.2005.73.5_suppl.0730038
Address correspondence to Myriam Arévalo-Herrera, Instituto de Inmunología, Edificio de Microbiología, Tercer Piso, Facultad de Salud, Universidad del Valle, Sede San Fernando, AA 25574, Cali, Colombia. E-mail: marevalo@inmuno.org
Authors’ addresses: Myriam Arévalo-Herrera, Yezid Solarte, Felipe Zamora, Maria Fernanda Yasnot, Leonardo Rocha, and Sócrates Herrera, Instituto de Inmunología, Edificio de Microbiología, Tercer Piso, Facultad de Salud, Universidad del Valle, Sede San Fernando, AA 25574, Cali, Colombia, Telephone: 57-2-558-1931, Fax: 57-2-557-0449, E-mail: marevalo@inmuno.org and Malaria Vaccine and Drug Development Center, Carrera 35 No 4A-53, AA 26020, Cali, Colombia, Telephone: 57-2-5583937, Fax: 57-2-5560141. Fabián Mendez, School of Public Health, Universidad del Valle, Sede San Fernando, AA 26020, Cali, Telephone: 57-2 558-1931 extension 107, Fax: 57-2-558-1931. Carole Long and Louis H. Miller, Malaria Vaccine Development Branch, Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD 20892, Telephone: 301-4352177, Fax: 301-435-2177, E-mail: lmiller@niaid.nih.gov.
Acknowledgments: We thank the community of Buenaventura for providing infected samples for this study, and Einer Celório and Sandra Preciado (Entomology Unit, Malaria Vaccine Development Branch) for technical assistance during these experiments.
Financial support: This work was supported by the National Institute of Allergy and Infectious Diseases through Tropical Medicine Research Centers grant no. 49486 and the World Health Organization/ Tropical Diseases Research Special Program (Research Capability Strengthening contract no. MVDC 991006).
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