INTRODUCTION
Concerns over the high prevalence of drug resistance and its impact on morbidity and mortality due to falciparum malaria1,2 have led many countries to adopt the strategy of combination therapy both to improve clinical response to malaria treatment and to delay the spread of antimalarial drug resistance.3,4 Following the demonstration that vector control measures can substantially reduce malaria transmission,5,6 malaria morbidity,7 and all-cause child mortality,8–11 it has been suggested that insecticide-treated materials (ITMs) may also curb the spread of antimalarial drug resistance.12 However, theoretical considerations also suggest that ITMs could favor the spread of drug resistance in the parasite population. If ITMs impair the acquisition of immunity to malaria,13 they might increase the proportion of malaria episodes that are symptomatic and hence the proportion of infections that encounter an antimalarial drug. Drug pressure is generally agreed to be the single most important factor in the spread of drug resistance.14,15 In addition, if the genetic basis of drug resistance is polygenic, reducing transmission intensity might reduce recombination breakdown of the loci encoding for resistance and hence favor the spread of resistance.16–19 Empirical data addressing this issue are limited. The slow spread of antimalarial resistance in Malaysia and Zimbabwe, countries with insecticide spraying programs, has been contrasted with the rapid development of resistance to chloroquine (CQ) observed in Malawi where there was no vector control program.12 More recently a study in Uganda that compared localities naturally exposed to low, intermediate, or high transmission concluded that resistance may spread fastest in areas of low or high transmission.20 Only two studies have directly examined the impact of vector control on drug resistance. A reduction in the frequency of mutant alleles (codons 51, 59, and 108) of the dhfr gene was observed 2 years after the introduction of treated bed nets in Tanzania.21 In Zimbabwe, indoor house-spraying was associated with reductions in clinical failure rates and the prevalence of alleles conferring resistance to CQ; pfmdr1 (codons 86 and 1246) and pfcrt (codon 76).22 Further, well-controlled studies, investigating sites with vector control interventions in place and comparable sites without interventions, are needed to address this question more thoroughly. We report a study from Burkina Faso, West Africa, which investigated whether long-term use of insecticide-treated curtains (ITCs) had affected the prevalence of genetic markers of resistance to CQ and sulfadoxine-pyrimethamine (SP) or had an impact on the efficacy of CQ when used for treatment of uncomplicated malaria in children.
MATERIALS
Study site.
The study was performed in Burkina Faso in 2002, in Ziniaré and Boussé health districts located north and northwest of the capital Ouagadougou. In the absence of vector control, malaria transmission is intense, with an average entomological inoculation rate (EIR) of 100–300 infective bites person/year, and highly seasonal. The main malaria vector is Anopheles gambiae s.l.6 At least 90% of children aged 6–59 months are parasitemic at the peak of malaria transmission.23 Almost all of the population in these districts belong to the Mossi ethnic group, the largest ethnic group in Burkina Faso. At the time of the study, CQ remained the first-line drug for the treatment of uncomplicated malaria, with sulfadoxine-pyrimethamine (SP), used rarely, as the second-line drug. CQ resistance was first observed in Burkina Faso in 1987,24 but in studies between 1998 and 2001 clinical failure rates were low to moderate (6–19%).25
Selection of the study villages.
The study was conducted in 18 villages with functioning health facilities. Nine of the villages were selected from among 158 villages, covering an area of about 1000 km2, which received ITCs in 1994 or 1996 as part of a trial of the impact of ITCs on child mortality.11,26 Curtains were retreated annually with permethrin11 in all 158 villages until 2001 and the EIR in this area declined to less than 1 infective bite per person per month between 1994–2000 (Ilboudo-Sanogo E, unpublished data).27 Two ITC villages with functioning heath centers were not included in the study owing to their small population size. In 2002, because of limited funds, curtains were retreated only in the 9 villages selected for inclusion in the CQ efficacy study. Nine “control” villages that had never benefited from systematic vector control measures were selected from the area surrounding the ITC area. Villages were chosen on the basis of their accessibility, subject to their being at least 5 km from the edge of the ITC area. The EIR in the area outside the ITC zone was of the order of 18–32 infective bites per person per month in 1998–2000 (Ilboudo-Sanogo E, unpublished data). The location of each ITC and non-ITC village is shown in Figure 1.
Recruitment of children.
Children aged 6–59 months who sought care at the health facilities in the study villages between August and November 2002 with uncomplicated symptomatic malaria were screened for potential eligibility and enrolled in an in vivo study of CQ efficacy, after obtaining written, informed consent from their parent(s). A modified version of the standard World Health Organization (WHO) in vivo method for assessing therapeutic efficacy of antimalarial drugs28 was implemented. Inclusion and exclusion criteria were the same as those described in the WHO protocol,28 except that the eligible asexual parasite density range was extended to 1000–150000 parasites per μL and the axillary temperature range to ≥ 37.5°C and ≤ 40°C. At presentation (Day 0), before CQ treatment, 250–500 μL of finger prick blood were taken to prepare thick and thin blood films for malaria diagnosis, to measure packed-cell volume (PCV), and to prepare filter paper blood spots for the detection of genetic markers of resistance to CQ and SP. A first dose of CQ was then administered and an appointment made for the next day (Day 1). Blood samples were transferred daily to the laboratory of the Centre National de Recherche et Formation sur le Paludisme (CNRFP) in Ouagadougou. Microscopic diagnosis of malaria and measurement of PCV were performed the same day and the results returned to the field the next morning. Children whose eligibility had been confirmed were then formally enrolled and received a second dose of CQ (Day 1). Appointments were made with the caretaker for further visits on days 2, 3, 7, and 14 for treatment or monitoring of treatment outcomes. Caretakers were also advised to bring children to the health center at any time between these scheduled visits if the child’s condition did not improve. Thick and thin blood films and filter paper blood spots were prepared on days 3, 7, and 14 and at unscheduled visits.
In September 2002, a sample of asymptomatic children aged 6–59 months, selected at random from census lists, was enrolled in a cross-sectional survey after obtaining parental consent. Thick and thin blood films and filter paper blood spots were collected to detect molecular markers of resistance to CQ.
METHODS
Antimalarial treatment.
A standard treatment with CQ (tablets of 100 mg base; SmithKline Beecham) was administered (25 mg/kg body weight) over 3 days to children with uncomplicated malaria. Treatment was administered at the clinic under the supervision of a nurse. Children were watched for 30 minutes after taking CQ. In the event of persistent vomiting, quinine was administered by the intramuscular route and the child was withdrawn from the study. A single dose of SP (1.25 and 25 mg/kg body weight of pyrimethamine and sulfadoxine, respectively) was provided as the second-line drug in the event of clinical failure or failure to clear parasites on day 14.
Microscopic diagnosis of malaria.
Thick and thin blood films were stained with 3% Giemsa for 45 minutes. Asexual parasites of P. falciparum were counted against 400 white blood cells (WBC) by 2 independent laboratory technicians. The number of parasites per μL of blood was calculated assuming a WBC count of 8000/μL. In cases of discrepancy (positive versus negative parasites count or parasite densities differing by > 50%), blood smears were re-examined by a third laboratory technician. The two closest parasite counts (positive, negative, or difference between parasite counts ≤ 50%) were retained and the final parasite density was expressed as the arithmetic mean of the two counts.
Detection of mutations at the pfcrt-76 and pfmdr1-86 gene-loci.
DNA was extracted from filter paper blood spots using the Chelex method.29 DNA amplification was performed by nested PCR. PCR primers and conditions were as described by Sutherland et al.30 Restriction fragment length polymorphism by Apo I endonuclease restriction enzyme31 was performed to detect lysine (wild type) or threonine (mutant) at codon 76 of the pfcrt gene (K76T). Digested products were electrophoresed on a 3% agarose gel containing 0.5 μg/ml of ethidium bromide and DNA bands were visualized on a UV transilluminator. Sequence-specific oligonucleotide probing was performed for the detection of asparagine (wild type) or tyrosine (mutant) at codon 86 of the pfmdr1-86 gene (N86Y).30
Detection of mutations at the dhfr (51, 59 108) and dhps (437 and 540) gene loci.
The presence of point mutations at dhfr codons 51, 59, and 108 and at dhps codons 437 and 540 was examined in a random sample of pre-treatment specimens from symptomatic children. Mutations were screened by nested PCR amplification of a 594-bp fragment of the dhfr gene and a 711-bp fragment of the dhps gene. PCR mixtures were prepared for each gene separately. Sequence-specific oligonucleotide probing was performed on final PCR products to detect mutations at each of the specified loci of the dhfr and dhps gene. Details of PCR conditions and primers have been published elsewhere.32
Discrimination of recrudescence from new infection.
Merozoite surface protein 2 (msp2) gene polymorphisms were studied in pre-treatment and post-treatment samples of children with parasitemia between day 8 and 14. This gene has been extensively used as polymorphic marker in the field. Additional genotyping of msp1 appears to contribute little to the discrimination of new infections over genotyping msp2 alone; for example in one study in Tanzania msp1 genotyping improved the classification in 2.6% of the samples only (7/269).33 Block 3 of the repetitive region of the msp2 gene was amplified by nested PCR. Each PCR plate was prepared to contain samples from both ITC and non-ITC villages. The PCR primers and conditions used were the same as those described previously.34 Infections were scored according to a method described by Cattamanchi et al.35 An infection was classified as recrudescent if the sample obtained on the day of treatment failure had identical alleles to, or a subset of alleles of, those observed at baseline. When the failure day sample contained alleles observed in the day 0 sample plus new alleles representing < 50% of all alleles observed in the failure day sample, the infection was also scored as recrudescent. When ≥ 50% of all alleles observed in the failure day sample were different from those observed at baseline the infection was scored as “new”.
Data processing and statistical analyses.
Data were double-entered and verified using EPIINFO version 6.0 (Centers for Disease Control). Analyses were performed using STATA 8.0 (www.stata.com). Infections carrying a mixture of mutant and wild-type alleles of pfcrt-76 were classified as mutant. A similar procedure was adopted for pfmdr1-86. Clinical and parasitological outcomes were defined as in the WHO protocol.28 Clinical and parasitological failure and presence of genetic markers of resistance to CQ and SP were analyzed as binary outcomes, using Generalized Estimating Equations (GEE) with robust standard errors to account for intra-cluster (village) correlation. We performed further analyses to estimate the proportion of parasite strains carrying the Pfcrt-76 mutant allele, adjusting for multiplicity of infection, by maximizing the likelihood as described by Schneider et al. (2002).36
Sample size.
On average, each village contained about 240 children aged 6–59 months and we assumed that each child would experience at least 1 malaria episode per year. Considering the study eligibility criteria, we assumed that we would be able to enroll, treat, and follow-up about 50 children with uncomplicated malaria per village over the 4-month study period. The formula described by Hayes and Bennett37 was used to calculate the number of villages required in ITC and non-ITC areas. Assuming that clinical and parasitological failure rates were 15% (ranging from 10–20%) and 25% (ranging from 20–30%) respectively, 9 villages were required per group to provide the study with 80% power to detect, at the 5% significance level, a 50% difference in the clinical failure rate (i.e., decreased to 7.5% or increased to 22.5%), assuming a design effect of 1.5. This sample size provides similar power to detect a 40% difference in the parasitological failure rate.
Ethical approval.
Ethical approval to conduct the study was obtained from the Ministry of Health of Burkina Faso and the ethics committee of the London School of Hygiene and Tropical Medicine.
RESULTS
Study profile.
For the in vivo study of CQ efficacy, 2,029 children aged 6–59 months with presumptive malaria were screened for inclusion. Nine hundred and thirty-four (46%) of these children were not eligible and 6 were mistakenly considered ineligible. The proportions of ineligible children were similar in ITC and non-ITC villages (48% versus 44%, P = 0.17). Fifty-four (5%) of the 1089 children who were enrolled were excluded from subsequent analyses for various reasons, with similar proportions in the two groups of villages (P = 0.2): 26 losses to follow-up (13 in ITC villages), 6 refusals (5 in ITC villages), 8 treatments changed due to persistent vomiting (5 in ITC villages), 1 parallel treatment with quinine (in an ITC village), 9 with broken or lost slides (4 in ITC villages), and 4 uncompleted treatments (4 in ITC villages). Thus, 1035 children (497 in ITC villages) with uncomplicated malaria were enrolled, correctly treated, and followed-up for the in vivo study of CQ efficacy. Forty-three of these children (18 in ITC villages) were excluded from the analysis of the association of ITC use with clinical and parasitological failure because msp2 polymorphisms could not be determined to differentiate re-infection from recrudescence. Thus, the final analysis of clinical and parasitological failures included 992 children (479 in ITC villages). Age and sex distributions were similar in the two groups of villages (Table 1). At enrollment, the prevalence of high temperature (> 38.5°C and ≤ 40°C), the distribution of parasite densities and the prevalence of moderate anemia were similar in ITC and non-ITC villages. Approximately similar proportions of children were recruited in each group each month. The mean number of parasite clones per infection, as assessed by msp2 polymorphisms was 2.30 in ITC villages and 2.67 in non-ITC villages (P = 0.26).
Clinical and parasitological response to treatment.
Overall, after correction for new infections, 164 (17%) children experienced clinical failure. Clinical failure rates varied from 2–33% (P < 0.001) across villages (Table 2). The risk of clinical failure was 14% in ITC villages compared with 19% in non-ITC villages (Table 3), (OR = 0.68; P = 0.17). Unadjusted clinical failures were 14% (67) in ITC villages and 19.5% (100) in non-ITC villages (data not shown). Age-stratified analyses showed similar patterns in younger (6–35 months) (OR = 0.69; P = 0.20) and in older children (36–59 months) (OR = 0.61; P = 0.30) (see Table 3). There was no evidence that odds of clinical failure varied with age (P = 0.76).
Five hundred and thirty-four (54%) children were classified as parasitological failures (after correction for new infections) with a range of 30–70% between villages (P < 0.001) (see Table 2). Older children experienced fewer parasitological failures than younger children (adjusted OR = 0.60; 95%CI: 0.40, 0.92; P = 0.02). The risk of parasitological failure was 49% in ITC villages compared with 58% in non-ITC villages (adjusted OR= 0.71; P = 0.15) (see Table 3). Uncorrected estimates were 52% (249) and 63% (323) respectively in ITC and non-ITC villages (data not shown). There was evidence that the association of ITCs with parasitological failure varied with age (P = 0.01). In younger children, the odds of parasitological failure were lower in ITC villages than non-ITC villages (OR = 0.57; 95%CI: 0.35, 0.91; P = 0.02) while in older children there was no evidence of a difference between the two groups of villages (OR = 1.14; P = 0.77) (see Table 3).
Prevalence of pfcrt-76 and pfmdr1-86 mutations in children with uncomplicated malaria.
Nine hundred and ninety-nine and 973 samples of the 1035 samples obtained pre-treatment from symptomatic children were successfully analyzed by PCR for the detection of pfcrt-76 and pfmdr1-86 alleles, respectively. Overall, 41% (409) of children harbored parasites carrying the pfcrt-76T allele, with a range of 23–57% across villages (P = 0.003) (Table 4). Similar proportions of children in ITC and non-ITC villages carried parasites with this mutation (Table 5): 43% and 40%, respectively (OR = 1.09; P = 0.65). The pfmdr1-86Y mutation was observed in 30% of children in the study area, with similar proportions in ITC villages (31%) and non-ITC villages (29%) (OR = 1.14; P = 0.54). After taking multiple infections into account, there was no indication that the proportion of parasite strains carrying the pfcrt-76 mutant allele differed between the two groups of villages (25% in ITC villages, 26% in non-ITC villages).
Analyses that accounted for clustering and adjusted for age, parasite density at enrollment, and for the use of ITCs, indicated a strong association of pfcrt-76T with pfmdr1-86Y (OR = 3.14; 95%CI: 2.38, 4.15; P < 0.001). Pfcrt-76T (OR = 5.0; 95%CI: 3.4, 7.2; P < 0.001), pfmdr1-86Y (OR = 1.7; 95%CI: 1.3, 2.3; P = 0.001), and pfcrt-76T/pfmdr1-86 (OR = 7.4; 95%CI: 5.9, 9.2; P < 0.001) genotypes were all associated with increased risk of parasitological failure. Pfcrt-76T (OR = 4.4; 95%CI: 2.7, 7.3; P < 0.001) and pfcrt-76T/pfmdr1-86 (OR = 6.4; 95%CI: 4.0, 10.3; P < 0.001) genotypes were also associated with clinical failure. There was no evidence that the pfmdr1-86Y mutation was associated with risk of clinical failure in the absence of pfcrt-76T (OR = 1.3; 95%CI: 0.7, 2.5; P = 0.5), though such an association cannot be ruled out.
Prevalence of pfcrt-76T and the pfmdr1-86Y in asymptomatic children.
Two hundred and thirty-one (84%) of 276 children seen in the cross-sectional survey were infected with P. falciparum: 110 (82%) and 121 (86%) in ITC and non-ITC villages, respectively. PCR amplification was successful in 197 (pfcrt-76) and 211 (pfmdr1-86) children. Age, sex, and parasite density distributions of children were similar in ITC and non-ITC villages. Similar proportions of asymptomatically infected children in ITC and non-ITC villages carried parasites with the pfcrt-76T allele (47% versus 46%; OR = 1.01; 95%CI: 0.56, 1.84; P = 0.97) and with the pfmdr1-86Y allele (36% versus 33%; OR = 1.05; 95%CI: 0.57, 1.95; P = 0.86). The pfcrt-76T mutant allele was strongly associated with pfmdr1-86Y (OR = 2.64; 95%CI: 1.35, 5.15; P = 0.004).
Prevalence of dhfr (51, 59 108) and dhps (437, 540) alleles in symptomatic children.
Of 509 randomly selected pre-treatment samples, 453 and 397 were successfully analyzed for dhfr (51, 59, and 108) and dhps (437 and 540) mutations, respectively. Wild-type alleles at all 3 loci of the dhfr gene were observed in 74% of children in ITC villages and in 73% of children from non-ITC villages (OR = 1.09; 95%CI: 0.67, 1.17; P = 0.74). The dhfr-108N mutation alone and the double mutation dhfr-108N-51I were observed in fewer than 5% of children in both groups of villages. Dhfr-108N-59R was observed in 9% of children in ITC villages and in 12% in non-ITC villages (OR = 0.72; 95%CI: 0.35, 1.48; P = 0.38), while the dhfr triple mutation (108N-51I-59R) was present in 12% of children from both ITC and non-ITC villages (OR = 0.96, 95%CI: 0.47, 1.94; P = 0.90). The proportion of infections with parasites carrying a mutation at dhps-437 only was high (≈58%) in children from both groups of villages. The dhps-540 mutation was observed in 12% and 9% of infections in ITC and non-ITC villages, respectively (OR = 1.43; 95%CI: 0.72, 2.91; P = 0.31). However, no infections with a mutation at both the dhps-437 and dhps-540 loci were observed in the study area.
DISCUSSION
We compared measures of CQ resistance in villages in Burkina Faso with and without ITC, in a setting where CQ was the first-line treatment of falciparum malaria cases, and the use of other antimalarial drugs, including SP, was rare.
We observed that the odds of clinical and parasitological failure after treatment with CQ were about 30% lower among children in ITC villages than among children in non-ITC villages. However, statistical analyses indicate that both these observed differences could have arisen by chance. In Zimbabwe22 indoor spraying was reported to be associated with an 80% (OR = 0.2; 95%CI: 0.08, 0.65) reduction in the odds of clinical failure. Our confidence intervals overlap with those of the study in Zimbabwe, which could not take account of between village variation, since only one village received the intervention, and did not consider factors other than house-spraying that could account for the observed reduction. We observed a reduction (OR = 0.57; P = 0.02) in parasitological failure rates in younger children (6–35 months) in ITC villages, consistent with the findings from Zimbabwe.
We detected similar proportions of infections (both symptomatic and asymptomatic), with parasites carrying the pfcrt-76T and pfmdr1-86Y alleles associated with resistance to CQ in ITC and non-ITC villages. About 40% of infections involved parasites with the pfcrt-76T mutation while about 30% involved parasites with the pfmdr1-86Y mutation. A similar proportion of infections with parasites carrying the pfmdr1-86Y allele (36%) was observed a few years earlier in an urban area in Burkina Faso,38 but the proportion of infections with the pfcrt-76T allele (61%) was much higher there. This is unsurprising as drug pressure is likely to be higher in urban than rural areas as a result of better access to antimalarial drugs. The proportion of infections harboring parasites with the pfcrt-76T allele is similar to that reported from rural communities in Mali (41%).31
We also observed similar proportions of infections in the two groups of villages with parasites carrying the mutations in the dhfr and dhps genes associated with SP resistance. For an area in which SP had been little used, we observed a surprisingly high proportion (58%) of infections with the dhps-437G mutation. As dhfr mutant alleles are rare, it seems unlikely that the high prevalence of dhps mutations is due to SP use. It may reflect instead the effect of widespread use of co-trimoxazole in the population.
We have presented our data as the proportion of infected individuals carrying parasites with drug resistance-associated mutations, which depends on both the proportion of mutant parasites in the parasite population and the number of parasite genotypes with which an individual is infected. We did not observe a major difference in the multiplicity of infections between ITC and non-ITC villages, a finding consistent with a previous report from the same area.39 Analyses taking multiple infections into account did not alter the study findings with respect to prevalence of the pfcrt-76T mutation in the two groups of villages. We believe therefore that our results are unlikely to be confounded to any major degree by differences in the number of genotypes per infection.
Reductions in the prevalence of genetic markers of resistance to CQ and SP associated with vector control measures have been reported from studies in Tanzania21 and Zimbabwe.22 In Tanzania, insecticide-treated bed nets were associated with an increase in the proportion of infected individuals with parasites with the wild-type alleles of dhfr (51, 59, 108) (P < 0.001). This finding, after only 2 years of intervention, was surprising given that the baseline frequencies of dhfr (51, 59, 108) mutant alleles were high (about 60% of triple mutations, after excluding mixed infections), and because SP was still in use in the area as first-line drug for malaria treatment despite waning efficacy. The Zimbabwean study reported a decrease in the proportions of infected individuals carrying parasites with mutant alleles at the pfcrt-76 (OR = 0.45; 95%CI: 0.22, 0.91), pfmdr1-86 (OR = 0.42; 95%CI: 0.21, 0.83), and pfmdr1-1246 (OR = 0.24; 95%CI: 0.09, 0.59) gene loci in a village that benefited from indoor spraying over a period of 4 years. In the current study ITCs were implemented, free of charge, in a larger number of villages (158 villages) and over a longer period. However, the confidence intervals around our point estimates do not preclude reductions in the odds of carriage of parasites with the pfcrt-76T allele of 20% in symptomatic and 44% in asymptomatic children from ITC villages. Thus both our results and those from Zimbabwe are compatible with reductions of 10–20% in the odds of an individual being infected with parasites carrying the pfcrt-76T mutation associated with vector control measures.
Taken together, our findings do not support the hypothesis that sustained use of ITCs would enhance the evolution of drug resistance in the P. falciparum population. Could this conclusion be flawed due to limitations in our study design? To answer this question we examined a number of factors with the potential to confound or interfere with our comparison of ITC and non-ITC villages. In 2003 we collected data on the socio-economic and health-seeking characteristics of the study villages. The two groups of villages were similar with respect to the distribution of assets (Diallo DA, unpublished data) and all households had easy physical access to a health facility located in their village. Due to the loss of reagents, we were unable to measure concentrations of CQ and SP in the blood or urine. Instead, we examined the availability and reported use of other antimalarials, including SP, and found no evidence of a difference between these communities. ITCs had not been taken up in non-ITC villages, nor had any other vector control programs taken place in these villages that might mask any impact of ITCs on drug resistance. The proportion of children who used a treated bed net was less than 2% in both groups of villages.
In 2001, because of financial constraints, only badly damaged curtains were replaced, and in 2002 only villages selected to be part of the CQ efficacy study had their curtains retreated. In addition, previous surveys in the area have shown that the quality of curtain usage declined between 1996 and 2000.23,26 Individuals without treated nets or curtains who live in protected communities have been reported to benefit from the indirect effect of treated nets or curtains,27,40,41 but it remains unclear what level of coverage must be achieved to obtain such protection. Thus, reduced use of ITCs in recent years may have led to a less marked impact on vector populations than seen in the early years of intervention, attenuating any impact of ITCs on drug resistance. A reduction in the effectiveness of ITCs during the 1–2 years leading up to the current study may partly explain the substantial rise in EIR observed in 2002: 54 and 87 infective bites/person/month in ITC and non-ITC villages, respectively (Ilboudo-Sanogo, unpublished data). Such a sharp rise in EIR was unexpected given that the 9 ITC study villages had curtains retreated in 2002, and that the other villages in the ITC area had curtains retreated the previous year and treated materials remain effective at least 1 year after treatment.11,42 Pyrethroid sensitivity tests (1% permethrin), carried out in 4 villages at the center of the ITC area in 2002, indicated a 100% anopheline mortality rate after 24 hours (Ilboudo-Sanogo, unpublished data). The 50% and 90% knock down times were 9 min and 32 min, respectively. Molecular analyses of the knockdown-resistance gene during the same periods did not detect the presence of pyrethroid-resistant vectors.
Movements of humans and malaria vectors promote the spread of resistance by facilitating exchanges of parasite strains between different areas.43–45 There was no history of recent large-scale migration in our study areas. Data we collected by interview indicate that rates of population movement were low in both ITC and non-ITC areas (3% and 2% of residents having spent a night outside the village in the preceding 2 weeks). Malaria vectors are able to fly between villages separated by short distances.46 We tried to minimize the effect of parasite gene flow by selecting control villages located at least 5 kilometers from the edge of the ITC protected area. This should have reduced gene flow from the ITC area to our control villages. However, 5 ITC villages involved in the study were located on the edge of the ITC protected area and were therefore relatively susceptible to gene flow from outside the ITC area. Thus, the possibility that human and vector movements between protected and unprotected villages attenuated any impact of ITCs on drug resistance cannot be excluded. If this is the case in our setting, where very high coverage was achieved over a large area over a period of 6–8 years, it seems unlikely that implementation of ITMs at relatively low coverage, as in many program settings, will have a marked impact on the evolution of drug resistance.
Our study was not a randomized controlled trial, as such a study would now be considered unethical, but it was designed to obtain the maximum information available from observations made in an area where ITCs have been used for many years. While we cannot exclude the possibility that differences in the prevalence of resistant parasites were present between ITC and non-ITC villages before the intervention, we think this is unlikely as the two groups of villages were well matched by socio-economic and other variables and control villages were located all around the ITC area.
Given the relatively long half-life of CQ, it is possible that a 14-day in vivo study of CQ efficacy resulted in an underestimation of treatment failures. However, we do not believe that this has had a major role in the lack of a difference between these communities, as the prevalence of genetic markers of resistance to CQ at baseline was similar in ITC and non-ITC villages.
In summary, we believe that the results of this study are robust. These results show a tendency for the response to treatment with CQ to be better in children from ITC than non-ITC villages and the prevalence of molecular markers of resistance to be similar. Thus, we can be reasonably confident that widespread use of ITCs or ITNs will not facilitate the spread of resistant parasites and that such concerns should not hold back widespread deployment of these very effective malaria control interventions. After this study, national policy in Burkina Faso was changed, with arthemeter/lumefantrine adopted in February 2005 as the first-line drug for the management of uncomplicated malaria. The implementation of this policy is expected to start in 2006.
Baseline characteristics of children included in analyses of the association between ITCs and response to treatment with CQ*
| Non-ITC N = 513 | ITC N = 479 | |||
|---|---|---|---|---|
| % (n) | % (n) | P value | ||
| * ITC = insecticide-treated curtains; CQ = chloroquine. | ||||
| Age group | 6–11 months | 16.4 (84) | 17.9 (86) | |
| 12–23 months | 30.8 (158) | 27.4 (131) | ||
| 24–35 months | 23.8 (122) | 22.3 (107) | 0.64 | |
| 36–47 months | 16.7 (86) | 18.4 (88) | ||
| 48–59 months | 12.3 (63) | 14.0 (67) | ||
| Sex | Female | 44.6 (229) | 45.7 (219) | 0.73 |
| Temperature | > 38.5 & < = 40.0°C | 45.0 (231) | 46.6 (223) | 0.94 |
| Moderate anaemia | PCV < 25% | 14.2 (73) | 13.6 (65) | 0.76 |
| Parasite density | 1000–10000/μL | 29.0 (149) | 25.5 (122) | |
| 10001–75000/μL | 34.7 (178) | 35.1 (168) | 0.40 | |
| 75001–150000/μL | 35.3 (186) | 39.4 (189) | ||
| Recruitment period | August 2002 | 31.8 (163) | 32.2 (154) | |
| September 2002 | 34.5 (177) | 32.2 (154) | 0.19 | |
| October 2002 | 21.6 (111) | 26.4 (127) | ||
| November 2002 | 12.1 (62) | 9.2 (44) | ||
Response to treatment with CQ in ITC and non-ITC villages*
| Non-ITC | ITC | ||||
|---|---|---|---|---|---|
| Village | Clinical failure % (n/N) | Parasitologic failure % (n/N) | Village | Clinical failure % (n/N) | Parasitologic failure % (n/N) |
| * ITC = insecticide-treated curtains; CQ = chloroquine. | |||||
| Golmidou | 33 (18/54) | 69 (37/54) | Barkuitenga | 16 (9/57) | 36 (21/57) |
| Gonsé | 17 (10/59) | 59 (35/59) | Laway | 17 (9/52) | 60 (31/52) |
| Likenkelse | 21 (12/56) | 61 (34/56) | Donsé | 2 (1/56) | 50 (28/56) |
| Nagreongo | 16 (10/64) | 66 (42/64) | Loumbila | 26 (13/50) | 70 (35/50) |
| Seguedin | 29 (16/55) | 67 (37/55) | Nioniogo | 8 (5/67) | 39 (26/67) |
| Sawana | 5 (3/62) | 52 (32/62) | Yaoghin | 16 (9/57) | 56 (32/57) |
| Sourgoubila | 29 (18/62) | 60 (37/62) | Tiben | 17 (8/46) | 30 (14/46) |
| Yimiougou | 2 (1/56) | 34 (19/56) | Voaga | 9 (5/53) | 59 (31/53) |
| Zibako | 24 (11/45) | 56 (25/45) | Bendogo | 15 (6/41) | 44 (18/41) |
| Nedogo | |||||
| Total | 19 (99/513) | 58 (298/513) | Total | 14 (65/479) | 49 (236/479) |
Clinical and parasitological failure rates by age and village type treating re-infections as successful treatments
| Number of children | Failure rates | |||||||
|---|---|---|---|---|---|---|---|---|
| Age group (months) | Non-ITC | ITC | Non ITC % (n) | ITC % (n) | Adjusted OR (95% CI) | P value | ||
| The odds ratios were adjusted for age parasite density and pfmdr1-86 (for pfcrt-76) or pfcrt76 (for pfmdr1-86) using a GEE regression model and 95%CI were calculated using robust standard errors. | ||||||||
| Clinical failure | 6–35 | 364 | 324 | 22 (79) | 16 (51) | 0.69 (0.40, 1.21) | 0.20 | Test for interaction |
| 36–59 | 149 | 155 | 13 (20) | 9 (14) | 0.61 (0.24, 1.55) | 0.30 | ||
| 6–59 | 513 | 479 | 19 (99) | 14 (65) | 0.68 (0.39, 1.18) | 0.17 | P = 0.76 | |
| Parasitologic failure | 6–35 | 364 | 324 | 64 (234) | 50 (161) | 0.57 (0.35, 0.91) | 0.02 | Test for interaction |
| 36–59 | 149 | 155 | 43 (64) | 48 (75) | 1.14 (0.50, 2.30) | 0.77 | ||
| 6–59 | 513 | 479 | 58 (298) | 49 (236) | 0.71 (0.44, 1.13) | 0.15 | P = 0.01 | |
Prevalence (%) of molecular markers of CQ resistance in children with uncomplicated malaria before treatment*
| Non-ITC | ITC | ||||
|---|---|---|---|---|---|
| Village | Pfcrt-76T | Pfmdr1-86Y | Village | Pfcrt-76T | Pfmdr1-86Y |
| CQ = chloroquine; ITC = insectide-treated curtains. | |||||
| Golmidou | 36 (19/53) | 19 (10/53) | Barkuitenga | 47 (26/55) | 41 (23/56) |
| Gonsé | 44 (27/61) | 33 (19/58) | Laway | 36 (19/53) | 27 (14/52) |
| Likenkelse | 45 (26/58) | 28 (16/57) | Donsé | 40 (22/55) | 24 (13/54) |
| Nagreongo | 57 (36/63) | 48 (29/60) | Loumbila | 55 (28/51) | 47 (24/51) |
| Seguedin | 36 (20/56) | 28 (16/57) | Nioniogo | 48 (33/69) | 33 (20/60) |
| Sawana | 29 (17/59) | 20 (11/56) | Yaoghin | 35 (19/54) | 42 (22/53) |
| Sourgoubila | 42 (22/53) | 32 (17/54) | Tiben | 29 (15/52) | 18 (9/51) |
| Yimiougou | 23 (13/56) | 21 (11/53) | Voaga | 57 (30/53) | 25 (13/53) |
| Zibako | 41 (24/58) | 27 (15/55) | Bendogo | 33 (13/40) | 23 (9/40) |
| Nedogo | |||||
| Total | 40 (204/517) | 29 (144/503) | Total | 43 (205/482) | 31 (147/470) |
Prevalence of parasites with pfcrt-76T and pfmdr1-86Y mutations in children with uncomplicated malaria by village type (ITC, non-ITC)*
| % (n) with mutant parasites | ||||||||
|---|---|---|---|---|---|---|---|---|
| Number of children | Failure rates | |||||||
| Age group (months) | Non-ITC | ITC | Non-ITC | ITC | Adjusted OR (95% CI) | P value | ||
| ITC = insecticide-treated curtains. | ||||||||
| The odds ratios were adjusted for age parasite density and pfmdr1-86 (for pfcrt-76) or pfcrt76 (for pfmdr1-86) using a GEE regression model and 95%CI were calculated using robust standard errors. | ||||||||
| Pfcrt-76T | 6–35 | 365 | 324 | 41 (150) | 40 (130) | 0.91 (0.62, 1.34) | 0.64 | Test for interaction |
| 36–59 | 152 | 158 | 36 (54) | 48 (75) | 1.56 (0.92, 2.63) | 0.10 | ||
| 6–59 | 517 | 482 | 40 (204) | 43 (205) | 1.09 (0.80, 1.50) | 0.65 | P = 0.13 | |
| Pfmdr1-86Y | 6–35 | 352 | 319 | 31 (106) | 33 (105) | 1.19 (0.76, 1.87) | 0.45 | Test for interaction |
| 36–59 | 151 | 151 | 25 (38) | 28 (42) | 1.02 (0.55, 1.89) | 0.96 | ||
| 6–59 | 503 | 470 | 29 (144) | 31 (147) | 1.14 (0.75, 1.72) | 0.54 | P = 0.86 | |
The white area in the center of the map represents the ITC protected area with the location of the nine villages involved in the in vitro study. Villages outside the white area represent the nine non-ITC villages.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 76, 2; 10.4269/ajtmh.2007.76.237
The white area in the center of the map represents the ITC protected area with the location of the nine villages involved in the in vitro study. Villages outside the white area represent the nine non-ITC villages.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 76, 2; 10.4269/ajtmh.2007.76.237
The white area in the center of the map represents the ITC protected area with the location of the nine villages involved in the in vitro study. Villages outside the white area represent the nine non-ITC villages.
Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 76, 2; 10.4269/ajtmh.2007.76.237
Address correspondence to Diadier Diallo, Centre National de Recherche et de Formation Sur Le Paludisme (CNRFP), Avenue de l’Oubritenga, 01 BP 2208 Ouagadougou 01, Burkina Faso. E-mail: ddiallo.cnlp@fasonet.bf
Authors’ addresses: Diadier A. Diallo, Issa Nebié, Amadou T. Konaté, and Edith Ilboudo-Sanogo, Centre National de Recherche et de Formation Sur Le Paludisme (CNRFP), Avenue de l’Oubritenga, 01 BP 2208 Ouagadougou 01, Burkina Faso, Telephone: +226 50 32 46 95, Fax +226 50 31 04 77, E-mails: ddiallo.cnlp@fasonet.bf, and issanebie.cnlp@fasonet.bf, and a.konate.cnlp@fasonet.bf. Colin Sutherland, Rosalynn Ord, Hirva Pota, Cally Roper, Brian M. Greenwood, and Simon N. Cousens, London School of Hygiene and Tropical Medicine (LSHTM), Keppel Street, London WC1E 7HT, United Kingdom, Telephone: +44 (0)20 7636 8636, Fax: +44 (0)20 7436 5389, E-mails: colin.sutherland@lshtm.ac.uk, rosalynn.ord@lshtm.ac.uk, hirva.pota@lshtm.ac.uk, cally.roper@lshtm.ac.uk, edith.cnlp@fasonet.bf, brian.greenwood@lshtm.ac.uk, and simon.cousens@lshtm.ac.uk.
Acknowledgments: The authors are most grateful to the population of the study villages and to the medical teams of Ziniaré, Boussé, Paul VI, secteur 30 and Kaya districts of the Ministry of Health of Burkina Faso. We are grateful to the Director of CNRFP, to the CNRFP staff, to the Gates Malaria Partnership staff, to Rachel Hallett and Anna Randall. The authors thank Andrew Thomson for his assistance in estimating the proportion of parasite strains carrying mutant alleles. This investigation received financial support from the Gates Malaria Partnership. ITC coverage was maintained from 1994–2002 thanks to the UNDP/World Bank/WHO Special Programme for Research and Training in Tropical Diseases (TDR), the European Commission (INCO-DC, Directorate General XII), the Danish Agency for International Development and the Ministry for University and Scientific Research of Italy. It formed part of a programme of activities run by CNRFP, under the bilateral co-operation agreement between Burkina Faso and the Italian Direzione Generale per la Cooperazione allo Sviluppo, Ministry of Foreign Affairs. The principal investigator received partial support from the Multilateral initiative on malaria/TDR.
Financial support: The study received financial support from the Gates Malaria Partnership, which is supported by the Bill and Melinda Gates Foundation. The principal investigator received financial support from Gates Malaria Partnership and the Multilateral Initiative on Malaria of UNDP/World Bank/WHO/TDR for his PhD training at the London School of Hygiene and Tropical Medicine, United Kingdom.
REFERENCES
- 1↑
Trape JF, 2001. The public health impact of chloroquine resistance in Africa. Am J Trop Med Hyg 64 :12–17.
- 2↑
Meerman L, Ord R, Teun Bousema J, van Niekerk M, Osman E, Hallett R, Pinder M, Walraven G, Sutherland CJ, 2005. Carriage of chloroquine-resistant parasites and delay of effective treatment increase the risk of severe malaria in Gambian children. J Infect Dis 192 :1651–1657.
- 3↑
WHO, 2001. Antimalarial drug combination therapy. Report of a WHO technical consultation. WHO/CDS/RBM/2001.35
- 5↑
Lindsay SW, Snow RW, Broomfield GL, Janneh MS, Wirtz RA, Greenwood BM, 1989. Impact of permethrin-treated bednets on malaria transmission by the Anopheles gambiae complex in The Gambia. Med Vet Entomol 3 :263–271.
- 6↑
Cuzin-Ouattara N, Van den Broek AH, Habluetzel A, Diabate A, Sanogo Ilboudo E, Diallo DA, Cousens SN, Esposito F, 1999. Wide-scale installation of insecticide-treated curtains confers high levels of protection against malaria transmission in a hyperendemic area of Burkina Faso. Trans R Soc Trop Med Hyg 93 :473–479.
- 7↑
Lengeler C, 2000. Insecticide-treated bednets and curtains for preventing malaria. Cochrane Database Syst Rev 2 :Cd000363.
- 8↑
D’Alessandro U, Olaleye BO, McGuire W, Langerock P, Bennett S, Aikins MK, Thomson MC, Cham MK, Cham BA, Greenwood BM, 1995. Mortality and morbidity from malaria in Gambian children after introduction of an impregnated bed-net programme. Lancet 345 :479–483.
- 9
Nevill CG, Some ES, Mung’ala VO, Mutemi W, New L, Marsh K, Lengeler C, Snow RW, 1996. Insecticide-treated bednets reduce mortality and severe morbidity from malaria among children on the Kenyan coast. Trop Med Int Health 1 :139–146.
- 10
Binka FN, Kubaje A, Adjuik M, Williams LA, Lengeler C, Maude GH, Armah GE, Kajihara B, Adiamah JH, Smith PG, 1996. Impact of permethrin impregnated bednets on child mortality in Kassena-Nankana district, Ghana: a randomized controlled trial. Trop Med Int Health 1 :147–154.
- 11↑
Habluetzel A, Diallo DA, Esposito F, Lamizana L, Pagnoni F, Lengeler C, Traore C, Cousens SN, 1997. Do insecticide-treated curtains reduce all-cause child mortality in Burkina Faso? Trop Med Int Health 2 :855–862.
- 12↑
Molyneux DH, Floyd K, Barnish G, Fevre EM, 1999. Transmission control and drug resistance in malaria: a crucial interaction. Parasitol Today 15 :238–240.
- 13↑
Snow RW, Bastos de Azevedo I, Lowe BS, Kabiru EW, Nevill CG, Mwankusye S, Kassiga G, Marsh K, Teuscher T, 1994. Severe childhood malaria in two areas of markedly different falciparum transmission in east Africa. Acta Trop 57 :289–300.
- 14↑
Payne D, 1988. Did medicated salt hasten the spread of chloroquine resistance in Plasmodium falciparum. Parasitol Today 4 :112–115.
- 15↑
White NJ, 1999. Delaying antimalarial drug resistance with combination chemotherapy. Parassitologia 41 :301–308.
- 16↑
Curtis CF, Otoo LN, 1986. A simple model of the build-up of resistance to mixtures of anti-malarial drug. Trans R Soc Trop Med Hyg 80 :889–892.
- 17
Paul RE, Packer MJ, Walmsley M, Lagog M, Ranford Cartwright LC, Paru R, Day KP, 1995. Mating patterns in malaria parasite populations of Papua New Guinea. Science 269 :1709–1711.
- 18
Dye C, Williams BG, 1997. Multigenic drug resistance among inbred malaria parasites. Proc R Soc Lond B Biol Sci 264 :61–67.
- 19↑
Hastings IM, D’Alessandro U, 2000. Modelling a predictable disaster: the rise and spread of drug-resistantmalaria. Parasitol Today 16 :340–347.
- 20↑
Talisuna AO, Langi P, Mutabingwa TK, Van Marck E, Speybroeck N, Egwang TG, Watkins WW, Hastings IM, D’Alessandro U, 2003. Intensity of transmission and spread of gene mutations linked to chloroquine and sulphadoxine-pyrimethamine resistance in falciparum malaria. Int J Parasitol 33 :1051–1058.
- 21↑
Alifrangis M, Lemnge MM, Ronn AM, Segeja MD, Magesa SM, Khalil IF, Bygbjerg IC, 2003. Increasing prevalence of wild-types in the dihydrofolate reductase gene of Plasmodium falciparum in an area with high levels of sulfadoxine/pyrimethamine resistance after introduction of treated bed nets. Am J Trop Med Hyg 69 :238–243.
- 22↑
Mharakurwa S, Mutambu SL, Mudyiradima R, Chimbadzwa T, Chandiwana SK, Day KP, 2004. Association of house spraying with suppressed levels of drug resistance in Zimbabwe. Malaria J 3 :1–9.
- 23↑
Habluetzel A, Cuzin N, Diallo DA, Nebie I, Belem S, Cousens SN, Esposito F, 1999. Insecticide-treated curtains reduce the prevalence and intensity of malaria infection in Burkina Faso. Trop Med Int Health 4 :557–564.
- 24↑
Guiguemde TR, Aouba A, Ouedraogo JB, Lamizana L, 1994. Ten-year surveillance of drug-resistant malaria in Burkina Faso (1982–1991). Am J Trop Med Hyg 50 :699–704.
- 25↑
Tinto H, Zoungrana EB, Coulibaly CO, Ouedraogo JB, Traore M, Guiguemde TR, Van Marck E, D’Alessandro U, 2002. Chloroquine and pyrimethamine efficacy for uncomplicated malaria treatment and hematological recovery in children in Bobo-Dioulasso, Burkina Faso during a 3-year period 1998–2000. Trop Med Int Health 7 :925–930.
- 26↑
Diallo DA, Cousens SN, Cuzin-Ouattara N, Nebie I, Ilboudo-Sanogo E, Esposito F, 2004. Child mortality in a West African population protected with insecticide-treated curtains for a period of up to 6 years. Bull World Health Org 82 :85–91.
- 27↑
Ilboudo Sanogo E, Cuzin-Ouattara N, Diallo DA, Cousens SN, Esposito F, Habluetzel A, Sanon S, Ouedraogo AP, 2001. Insecticide-treated materials, mosquito adaptation and mass effect: entomological observations after five years of vector control in Burkina Faso. Trans R Soc Trop Med Hyg 95 :353–360.
- 28↑
WHO, 1996. Assessment of therapeutic efficacy of antimalarial drugs for uncomplicated falciparum malaria in areas with intense transmission. WHO/MAL/1996.1077.
- 29↑
Plowe CV, Wellems TE, 1995. Molecular approaches to the spreading problem of drug resistant malaria. Adv Exp Med Biol 390 :197–209.
- 30↑
Sutherland CJ, Alloueche A, Curtis J, Drakeley CJ, Ord R, Duraisingh M, Greenwood BM, Pinder M, Warhurst D, Targett GA, 2002. Gambian children successfully treated with chloroquine can harbor and transmit Plasmodium falciparum game-tocytes carrying resistance genes. Am J Trop Med Hyg 67 :578–585.
- 31↑
Djimdé A, Doumbo OK, Cortese JF, Kayentao K, Doumbo S, Diourte Y, Dicko A, Su XZ, Nomura T, Fidock DA, Wellems TE, Plowe CV, Coulibaly D, 2001. A molecular marker for chloroquine-resistant falciparum malaria. N Engl J Med 344 :257–263.
- 32↑
Pearce RJ, Drakeley C, Chandramohan D, Mosha F, Roper C, 2003. Molecular determination of point mutation haplotypes in the dihydrofolate reductase and dihydropteroate synthase of Plasmodium falciparum in three districts of northern Tanzania. Antimicrob Agents Chemother 47 :1347–1354.
- 33↑
Felger I, Irion A, Steiger S, Beck HP, 1999. Genotypes of merozoite surface protein 2 of Plasmodium falciparum in Tanzania. Trans R Soc Trop Med Hyg 93 (Suppl 4):3–9.
- 34↑
Snounou G, Zhu X, Siripoon N, Jarra W, Thaithong S, Brown KN, Viriyakosol S, 1999. Biased distribution of msp1 and msp2 allelic variants in Plasmodium falciparum populations in Thailand. Trans R Soc Trop Med Hyg 94 :369–374.
- 35↑
Cattamanchi A, Kyabayinze D, Hubbard A, Rosenthal P, Dorsey G, 2003. Distinguishing recrudescence from reinfection in a longitudinal antimalarial drug efficacy study: comparison of results based on genotyping of MSP-1, MSP-2 and GLURP. Am J Trop Med Hyg 68 :133–139.
- 36↑
Schneider AG, Premji Z, Felger I, Smith T, Abdulla S, Beck H-P, Mshinda H, 2002. A point mutation in codon 76 of pfcrt of P. falciparum is positively selected for by Chloroquine treatment in Tanzania. Infect Genet Evol 1 :183–189.
- 37↑
Hayes R, Bennett S, 1999. Simple sample size calculation for cluster-randomised trials. Int J Epidemiol 28 :319–326.
- 38↑
Tinto H, Ouedraogo JB, Erhart A, Van Overmeir C, Dujardin JC, Van Marck E, Guiguemde TR, D’Alessandro U, 2003. Relationship between the Pfcrt T76 and the Pfmdr-1 Y86 mutations in Plasmodium falciparum and in vitro/in vivo chloroquine resistance in Burkina Faso, West Africa. Infect Genet Evol 3 :287–292.
- 39↑
Chiucchiuini A, Babiker H, Ranford-Cartwright L, Cuzin-Ouattara N, Nebie I, Cousens SN, Walliker D, Esposito F, 2001. Insecticide treated curtains and allelic polymorphism of Plasmodium falciparum genes in a rural area of Burkina Faso (west Africa). Parasitologia 43 (Suppl 1):7–10.
- 40↑
Curtis CF, Jana-Kara B, Maxwell CA, 2003. Insecticide treated nets: impact on vector populations and relevance of initial intensity of transmission and pyrethroid resistance. J Vector Borne Dis 40 :1–8.
- 41↑
Hawley WA, Phillips-Howard PA, ter Kuile FO, Terlouw DJ, Vulule JM, Ombok M, Nahlen BL, Gimnig JE, Kariuki SK, Kolczak MS, Hightower AW, 2003. Community-wide effects of permethrin-treated bed nets on child mortality and malaria morbidity in western Kenya. Am J Trop Med Hyg 68 :121–127.
- 42↑
Maxwell CA, Msuya E, Sudi M, Njunwa KJ, Carneiro IA, Curtis CF, 2002. Effect of community-wide use of insecticide-treated nets for 3–4 years on malarial morbidity in Tanzania. Trop Med Int Health 7 :1003–1008.
- 43↑
Warsame M, Wernsdorfer WH, Huldt G, Bjorkman A, 1995. An epidemic of Plasmodium falciparum malaria in Balcad, Somalia, and its causation. Trans R Soc Trop Med Hyg 89 :142–145.
- 44
Cortese JF, Caraballo A, Contreras CE, Plowe CV, 2002. Origin and dissemination of Plasmodium falciparum drug-resistance mutations in South America. J Infect Dis 186 :999–1006.
- 45↑
Roper C, Pearce R, Nair S, Sharp B, Nosten F, Anderson T, 2004. Intercontinental spread of pyrimethamine-resistant malaria. Science 305 :1124.
- 46↑
Thomson MC, Connor SJ, Quinones ML, Jawara M, Todd J, Greenwood BM, 1995. Movement of Anopheles gambiae s.l. malaria vectors between villages in The Gambia. Med Vet Entomol 9 :413–419.