https://orcid.org/0000-0003-1931-1442
Figures
Abstract
Background
Duffy Binding Protein (PvDBP) binding to the Duffy antigen receptor for chemokine (DARC) is essential for Plasmodium vivax invasion of human reticulocytes. PvDBP copy number variation (CNV) might increase parasite invasion and thus parasitemia. We examined the spatial distribution of PvDBP CNVs and DARC genotypes and their association with parasitemia in P. vivax endemic settings in Ethiopia.
Methodology/Principal findings
P. vivax isolates (n = 435) collected from five P. vivax endemic settings in Ethiopia were genotyped by amplifying the GATA1 transcription factor-binding site of the Duffy blood group and the CNV of PvDBP was quantified. Parasitemia was determined using 18S-based qPCR. The majority of participants were Duffy positive (96.8%, 421/435). Of the few Duffy negative individuals, most (n = 8) were detected from one site (Gondar). Multiple copies of PvDBP were detected in 83% (363/435) isolates with significant differences between sites (range 60%-94%). Both heterozygous (p = 0.005) and homozygous (p = 0.006) patients were more likely to have been infected by parasites with multiple PvDBP copies than Duffy negatives. Parasitemia was higher among the Duffy positives (median 17,218 parasites/µL; interquartile range [IQR] 2,895–104,489) than Duffy negatives (170; 78–24,132, p = 0.004) as well as in infections with 2 to 3 PvDBP copies (20,468; 3,649–110,632, p = 0.001) and more than 3 PvDBP copies (17,139; 2,831–95,946, p = 0.004) than single copy (5,673; 249–76,605).
Conclusions/Significance
A high proportion of P. vivax infection was observed in Duffy positives in this study, yet few Duffy negatives were found infected with P. vivax. The significant prevalence of multi-copy PvDBP observed among Ethiopian P. vivax isolates explains the high prevalence and parasitemia observed in clinical cases. This suggests that vivax malaria is a public health concern in the country where the Duffy positive population predominates. Investigating the relative contribution to the maintenance of the infectious reservoir of infections with different genotyping backgrounds (both host and parasite) might be required.
Author summary
P. vivax establishes infection in humans using its DBP which binds to the DARC receptor on host erythrocytes. A single point mutation in the DARC promoter region results in Duffy negative phenotype. This phenotype results in lower infection success of P. vivax and can explain the heterogeneity in the global geographic distribution of P. vivax. Here, we investigated the association between Duffy genotypes, CNV of PvDBP, and level of parasitemia using 435 P. vivax isolates from five different endemicities in Ethiopia. The results revealed that the majority of P. vivax infections do occur in Duffy positive individuals. This explains the high prevalence of P. vivax in Ethiopia. The high prevalence of multi-copy PvDBP along with the high rate of Duffy positivity explains the transmission success of P. vivax in the country, where the majority of the population are Duffy positive. The lower parasitemia levels among the Duffy-negatives, despite small samples, may signify an “undetected silent reservoir”.
Citation: Nasir Y, Molla E, Habtamu G, Sisay S, Ejigu LA, Kassa FA, et al. (2025) Spatial distribution of Plasmodium vivax Duffy Binding Protein copy number variation and Duffy genotype, and their association with parasitemia in Ethiopia. PLoS Negl Trop Dis 19(2): e0012837. https://doi.org/10.1371/journal.pntd.0012837
Editor: Ananias A. Escalante, Temple University, UNITED STATES OF AMERICA
Received: July 14, 2024; Accepted: January 13, 2025; Published: February 13, 2025
Copyright: © 2025 Nasir et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: The data is directly available using (https://doi.org/10.5061/dryad.7pvmcvf40) and the R codes are available using the sample GitHub link provided before (https://github.com/solsisay/Plasmodium-vivax-Duffy-binding-protein-copy-number-variation-and-Duffy-genotype).
Funding: FGT was supported by the Bill and Melinda Gates Foundation (ACHIDES; INV-005898, EMAGEN; INV-035257, and HAMMS; INV-048214) and the Wellcome Trust Early Career Award (UNS141457). EM and YN were supported by the Armauer Hansen Research Institute through its core funding from NORAD and SIDA. FGT received a salary from Bill and Melinda Gates Foundation and Wellcome Trust. The funders had no role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Malaria remains a significant public health problem globally with Africa bearing the major burden in terms of the number of cases and deaths, mainly attributable to Plasmodium falciparum [1]. Outside of Africa, P. vivax is the predominant human Plasmodium species responsible for more than half of malaria-associated cases, particularly in Afghanistan, India, and Pakistan [1]. Malaria in Africa has historically been linked to P. falciparum although recent reports demonstrated evidence of P. vivax transmission in different countries across the continent [2–4]. The infrequent presence of vivax malaria in western and central Africa is likely attributed to high Duffy-negativity among these populations (88–100%) [5]. However, reports confirmed that P. vivax is seen in Africa in areas where Duffy positive and negative individuals live together and in areas where Duffy negative population is predominant [6]. Uniquely, countries in the Horn of Africa such as Ethiopia [7], Eritrea [8], Djibouti [9], Somalia [10], and Sudan [11], are the most affected. In Ethiopia, P. vivax accounts for approximately 40% of clinical malaria cases [12] with varying innate susceptibility among populations to patent infection with vivax malaria [13,14].
P. vivax invasion of human erythrocytes relies on a protein (Duffy Binding Protein, PvDBP) - receptor (Duffy antigen) interaction on the youngest red blood cells, reticulocytes. The Duffy (Fy) antigen is a protein encoded by the Duffy antigen receptor for the chemokines (darc) gene on chromosome 1 [15]. DARC is located on both normocytes and reticulocytes; hence this ligand-receptor interaction does not govern selective entry into reticulocytes [16]. P. vivax reticulocyte binding protein (PvDBP) interaction with transferrin receptor (CD71) is important for the initial recognition of reticulocytes and explains its strict preference [17,18]. For a long, it was believed that P. vivax cannot infect Duffy negative individuals, which is caused by a point mutation in the promoter region of the DARC gene, and this is the reason that P. vivax is rare or absent from most of Africa [5,19]. More recent studies confirmed that Duffy-negativity does not confer complete protection against P. vivax infections [6,20–24]. The point mutation in the promoter does not necessarily abolish the expression of the Duffy antigen [22,25]. A recent study demonstrated that a subset of Duffy negative erythroblasts transiently express DARC and can thus be invaded by P. vivax merozoites [26]. Hence, Duffy negativity is confirmed to not be Duffy null [27,28]. A study in Madagascar showed that P. vivax is capable of infecting human erythrocytes without the Duffy antigen [24]. Recently, a similar line of argument emerged to explain the unexpected prevalence of P. vivax in Africa.
Amplification of the PvDBP gene copy number has been reported from Ethiopia and elsewhere [29,30]. The parasite’s DBP duplication is an important strategy for reticulocyte invasion [30]. PvDBP duplication might allow for binding to an alternative lower affinity receptor in Duffy negative reticulocytes or supports successful infection in Duffy negative patients who may have low level expression of the DARC gene [31,32]. In addition, the multiplication of the PvDBP gene may allow P. vivax to evade the host anti-PvDBP immune system [30]. However, the association of DARC genotyping and PvDBP CNVs with P. vivax parasitemia is not clearly established. Findings varied from similar parasitemia between heterozygous and homozygous Duffy positive individuals [21,22,33,34] to higher P. vivax parasitemia in homozygous than heterozygous individuals [20,35]. In terms of PvDBP CNVs, results varied from a study in Ethiopia that did not find an association with parasitemia [36] to another study in Sudan that showed an association of PvDBP duplications with increases in parasitemia levels [21]. As Ethiopia is a co-endemic setting, the different biological features of P. vivax compared with P. falciparum such as the hypnozoite stage, evidence of P. vivax infections in Duffy negative population in addition to Duffy positives as well as the presence of vectors with wide ranges of feeding and breeding behavior [37] may challenge malaria control and elimination activities.
Ethiopia, with its diverse population of Duffy positive and Duffy negative population [5,38], may provide insights into the epidemiology of P. vivax. In this study, the distribution of CNV of PvDBP and DARC genotyping and their association with parasitemias were investigated using P. vivax isolates from five different endemic settings in Ethiopia.
Methods
Ethical approval
The study protocol was approved by the Armauer Hansen Research Institute and All Africa Leprosy Rehabilitation and Training Center Ethics Review Committee (PO/46/20). Before sample collection, informed written consent and/or assent were obtained from all participants and/or the parents/guardians of the children.
Study setting, sampling, and sample collection
This study was carried out in five P. vivax and P. falciparum co-endemic settings in Ethiopia (Adama, Arba Minch Zuria, Batu, Dilla, and Gondar Zuria) from July to October 2021. A total of 435 symptomatic (n = 374) and asymptomatic (n = 61, only from Dilla Town) participants infected with P. vivax mono- (n = 415) and mixed-P. vivax and P. falciparum (n = 20) infections were included in this study. From each site, 86 to 88 participants were enrolled. Blood samples were collected from symptomatic P. vivax patients to investigate the association of P. vivax parasitemia (expressed in parasite copies per mL of blood) with host Duffy genotypes and PvDBP copy numbers, with additional asymptomatic community members recruited from one of the study sites (Dilla) in the same period as with clinical cases.
Five-year national malaria surveillance data reported from each of the selected study sites between January 2018 and December 2022, captured by the District Health Information System 2 (DHIS2), were used to analyze the trend and proportions of P. falciparum and P. vivax diagnosed by rapid diagnostic test (RDT) or microscopy in the five study sites.
Finger prick blood samples (approximately 0.5 mL) collected into EDTA microtainer tubes were used to diagnose malaria using an RDT (SD Bioline Malaria Ag Pf/Pv HRP2/LDH, Standard Diagnostics Inc., South Korea) and microscopic examination of Giemsa-stained blood films, and to prepare dried blood spots (DBS) on Whatman 3 MM filter paper. The DBS samples were allowed to air dry before being kept in zip-locked bags with self-indicating silica gel desiccant beads. The leftover blood in the EDTA tubes stored and shipped at −20 °C was used to extract genomic DNA using the MagMAX magnetic bead-based platform on the Kingfisher Flex robotic extractor (ThermoFisher Scientific) following the manufacturer’s protocol [39].
Plasmodium parasite species confirmation and quantification
Parasite species confirmation and quantification were done in a multiplex qPCR assay that targeted the 18 S rRNA genes for P. vivax and P. falciparum using primer and probe sequences described earlier [40,41] and TaqMan Fast Advanced Master Mix (Applied Biosystems) on a Bio-Rad CFX96 Real-Time PCR thermocycler (Bio-Rad Laboratories, Inc.). P. vivax copy numbers were quantified using serial dilutions (107 to 104 copies per mL) of recombinant plasmids that contained the amplicon, in duplicate per reaction plate.
DARC genotyping and copy number variation analysis of PvDBP genes
A qPCR-based TaqMan assay was used to analyze a point mutation in the darc gene GATA-1 transcription factor binding region, using previously reported primers and probes [22]. Previously confirmed samples were used as positive controls and molecular grade water as a negative control. The SYBR Green qPCR detection technique was used to determine the PvDBP CNV [30]. The CNV of the PvDBP gene was quantified relative to the single-copy β-tubulin gene (housekeeping gene). Briefly, PCRs were carried out in a total reaction mixture of 20 µL that contained 10 μL 2 X GoTaq qPCR Master Mix, 0.5 µL of each primer (at a concentration of 200 nM), and 2 μL of DNA extract. To detect the specificity of PCR amplification, a melting curve analysis was conducted between the temperature 65 °C and 95 °C with 0.5 °C increments. The PvDBP gene copy numbers were estimated by using synthetic gene (P. vivax β-tubulin and PvDBP) combined in varying ratios ranging from 1:1 to 1:6 (one-to-one copy of β-tubulin and one-to-six copies of PvDBP) as described before [42]. The difference in cycle threshold (ct) values between the first mix and 2 to 6 were equivalent to 1, 1.6, 2, 2.3, 2.8 respectively, (i.e., log2^x where x is the ratio of the mix). To determine PvDBP copy number, the equation that was previously employed for PvDBP copy number estimation was utilized as follows: N = 2^-ΔΔCt, where ΔΔCt = (Ct pvβ-t cal- Ct PvDBP cal) - (Ctpvβ-t - CtPvDBP). The Ctpvβ-t and CtPvDBP are threshold cycle values for the P. vivax β-tubulin and PvDBP genes respectively, whereas Ctcal is an average difference between Ctpvβ-tubulin and CtPvDBP obtained for the positive control. Then, PvDBP gene copy numbers were categorized into single copy (isolates with PvDBP estimates of ≤ 0.30), two to three copies (isolates with PvDBP estimates between 0.30 and ≤ 1.01), and more than three copies (PvDBP estimates > 1.01) [36].
Statistical analysis
Statistical analyses were performed using STATA (version 17.0, Stata Corp., TX, USA) and R (version 4.3.3, the R Foundation for Statistical Computing). The R codes are available using the sample GitHub link (https://github.com/solsisay/Plasmodium-vivax-Duffy-binding-protein-copy-number-variation-and-Duffy-genotype) and the data is directly available using https://doi.org/10.5061/dryad.7pvmcvf40 [43,44]. The sample size was calculated using a single population proportion formula assuming duplication of PvDBP to be 56% based on previous studies conducted in Ethiopia [31], 95% CI, 5% marginal error, and 15% non-response rate. Nonparametric tests were used for pairwise comparisons between variables. The Wilcoxon rank sum test was used to compare parasitemia between symptomatic and asymptomatic cases. The Kruskal-Wallis test was used to compare median parasitemia differences among age groups, Duffy genotypes, and PvDBP gene copy numbers. The Dunnett test was employed to test differences between the median parasitemias with variables such as age group, Duffy status, and PvDBP gene copy numbers. Continuous variables were presented as median and interquartile range (IQR). Ordinal logistic regression model was utilized to assess factors associated with PvDBP copy number. A scatter plot was used to determine the linearity of predictors and the target variable (S1–S3 Figs). A generalized additive model (GAM) with gamma link function that accounted for non-linearity was used to examine the association of parasitemia with DARC genotypes, PvDBP gene copy numbers, and age of the participants.
Results
P. vivax contributes substantially to the malaria burden in Ethiopia between 2018 and 2022
All the five study sites, with variable altitudes, are co-endemic for the two major species, P. falciparum and P. vivax. Overall, P. vivax contributed to 39% of malaria infections, detected by RDT and/or microscopy, reported from the five sites in the last 5 years. Despite slight variations during the years and notable differences between the study sites, P. vivax continues to be an important contributor to the overall malaria burden in Ethiopia between 2020 and 2022. The highest proportion of P. vivax infections was reported from Batu (~ 55%) with the lowest from Gondar Zuria (~ 32%) (Fig 1 and S1 Table).
The five-year proportion of P. vivax in the study sites is indicated together with the number of reported cases (by species) from the study sites (using DHIS2 data from 2018–2022). The bar plots depict the prevalence of P. vivax and P. falciparum (left Y-axes) for each year (X-axes) in the study sites. The trend lines (right Y-axes) show the proportion of P. vivax by year. The map indicates the study sites with altitude.
The base map of the Ethiopian administrative boundary shapefile (2021) was obtained from the GADM database (https://gadm.org/). The DEM data were sourced from the USGS Earth Explorer (https://earthexplorer.usgs.gov/) via the SRTM sensor. Both datasets are open-licensed, with GADM free for academic use and USGS in the public domain, ensuring compatibility with the CC BY 4.0 license. The map was created using ArcGIS software.
Characteristics of the study population
A majority of adult (61.1%, 266/435) males (56.3%, 245/435) who were confirmed to have P. vivax mono- (95.4%, 415/435) or mixed-species (P. falciparum and P. vivax) infections (4.6%, 20/435) by qPCR who were included in this study. Of the clinical cases, 98.7% (369/374) were confirmed to have microscopy-positive infections whilst the rest (n = 5) were only positive by qPCR. Of the asymptomatic community infections recruited in Dilla town, 63.9% (39/61) were negative by microscopy and were positive only by qPCR. qPCR detected median parasitemia was different between clinical (17,139; IQR: 2,895–109,895, n = 374) and asymptomatic (8,027; IQR: 1,638–37,204, n = 61, p = 0.020) infections (Table 1). The study findings show that the PvDBP copy numbers vary across the study sites (p < 0.001, S2 Table).
DARC genotyping and PvDBP copy numbers vary between study sites
Most of the participants were heterozygous (77.7%, 338/435) or homozygous (19.1%, 83/435) Duffy positive. Only 3.2% (14/435) of the participants were Duffy negative. Duffy genotypes varied between the study sites. All participants in one of the study sites (Batu) were heterozygous Duffy positive whilst the least heterozygosity was detected in Dilla town both in its symptomatic (61.4%, 51/83) and asymptomatic (59.0%, 36/61) communities. Most of the Duffy negative participants were from Gondar Zuria (57.1%, 8/14) (Fig 2A and Table 1).
The pie charts at each site represent the proportion of Duffy genotypes (a), PvDBP copy numbers (b), and bar graphs illustrating the relationship between Duffy genotypes and CNV of PvDBP (c). The Y-axis in the figure panel (c) implies the proportion of individuals with Duffy blood group for each CNV.
For PvDBP CNV, 83.4% (363/435) infections had multiple copies; two to three (50.3%, 219/435) or more than three copies (33.1%, 144/435), while only 16.6% (72/435) infections had a single copy. Differences were observed between sites; a higher number of multiple copies were detected in Arba Minch Zuria with two to three 46.6% (41/88) and more than three copies 47.7% (42/88) (Fig 2B and Table 1). The least PvDBP multiple copies were detected in Dilla town: both among the community members (59.0%, 36/61) and the clinical cases (64.0%, 16/25). Overall, PvDBP copy numbers significantly varied between clinical cases and asymptomatic community samples (p < 0.001): multiple copies were detected in 87.4% (327/374) clinical infections versus 59.0% (36/61) asymptomatic community members.
The frequency of multiple PvDBP copies was higher among Duffy positives (84.3%, 355/421) than Duffy negatives (57.1%, 8/14, p = 0.034) (S2 Table). Both heterozygous (77.7%, 338/435, p = 0.005) and homozygous (19.1%, 83/435, p = 0.006) patients were more likely to have been infected by parasites with multiple PvDBP copies than Duffy negatives (Fig 2C and S2 Table).
The base map of the Ethiopian administrative boundary shapefile (2021) was obtained from the GADM database (https://gadm.org/), which is freely available for academic and research purposes.
P. vivax parasitemia is associated with Duffy genotype and PvDBP gene copy number
Using a generalized additive model, parasitemia was found associated with the ages of participants, DARC genotyping status, and PvDBP copy number. Parasitemia increases by 0.9 in adults (Density ratio [DR] = 0.9, 95% CI: 0.78–0.95) and children between the age of 5 and 15 (DR = 0.9, 95% CI: 0.80–0.97) as compared to under-fives (Fig 3B and Table 2). Parasitemia was strongly linked with Duffy genotyping status: higher among the Duffy positives (median 17,218 parasites/µL; interquartile range [IQR] 2,895–104,489, n = 421) than Duffy negatives (170; IQR: 78–24,132, n = 14, p = 0.004). Specifically, individuals both with heterozygous (DR = 1.3, 95% CI: 1.07–1.25, n = 338) and homozygous (DR = 1.2, 95% CI: 1.17–1.55, n = 83) Duffy blood groups had increased parasitemia compared to those with the Duffy negative blood group (Table 2). Among the Duffy positive genotypes, median parasitemia was high in heterozygous Duffy blood groups (21,176; IQR: 3,275–111,375) compared to the homozygous Duffy blood group (8,078; IQR: 1,445–36,192, p = 0.009) (Fig 3C). The median parasitemia was also higher in infections with 2 to 3 PvDBP copies (20,468; IQR 3,649–110,632; n = 219, p = 0.001) and 4 or more PvDBP copies (17,139; IQR 2,831–95,946; n = 144, p = 0.004) compared to single copies (5,673; IQR 249–76,605; n = 72) (Fig 3D). Infections with PvDBP gene copy numbers of two to three carried 1.1 times higher parasitemia (DR = 1.1, 95% CI: 1.07–1.24) compared to those with a single copy. Likewise, samples with PvDBP copy numbers of more than three had 1.2 times higher parasitemia (DR = 1.2, 95% CI: 1.07–1.25) compared to those with a single copy (16.6%, 72/435) (Table 2).
The distribution of parasitemia is shown for symptomatic and asymptomatic individuals (A), age groups (B), Duffy genotypes (C), and PvDBP copy numbers (D) and comparison between groups using the Wilcoxon rank sum test (A) and Kruskal test (B to D). P-values were obtained by Dunnett tests, with P. vivax parasitemia (in Log10 Pv18S copies per microliter) as the outcome and age groups, Duffy status, or PvDBP copy numbers as predictors.
Discussion
In five settings in Ethiopia that are sympatric for the two species, P. falciparum and P. vivax, the latter contributed to an overall 39% of malaria infections reported in the last five years. We determined the spatial heterogeneity of DARC genotyping and PvDBP CNV and their association with parasitemia in these sites. Most of the P. vivax infected individuals in this study were Duffy positive whilst only fourteen Duffy negative individuals were found infected with P. vivax. Most of the parasites had multiple copies of the PvDBP gene (83% isolates) with substantial differences between sites, ranging from 94.3% in Arba Minch to only 60.5% in Dilla. Parasites with multiple PvDBP copies were detected more commonly (87.4%) in clinical infections than asymptomatic infections (59.0%) as well as in Dufy positive heterozygous (84.0%) and homozygous patients (85.5%) than Duffy negative patients (61.5%). Our investigation confirmed that parasitemia is associated with age and DARC genotyping status of patients and the number of PvDBP gene copies of the parasites in Ethiopia.
Malaria has been increasing in the last five years in Ethiopia, especially starting from 2020. This may be attributed to service interruptions attributed to the COVID-19 pandemic [45], the emergence of drug and diagnostic-resistant parasites [39,46–48], mosquito resistance to insecticides [1,45], the expansion of the invasive vector Anopheles stephensi [39,49–53], and climate change [1,54]. The contribution of P. vivax has remained substantial in Ethiopia [55]. The overall burden of malaria linked with P. vivax and its trend observed in the present study is in line with the national figure [ 7,12,55].
This study confirms the low but appreciable presence of the P. vivax parasite reservoir in Duffy negative populations. Duffy positive genotypes predominate among populations susceptible to P. vivax infections [56]. P. vivax endemicity is associated with the Duffy gene expression in the population in which this parasite is more endemic in areas where a high population of Duffy-positives exists [57,58]. Our result was corroborated by a prior report that found 86.9% of heterozygous P. vivax patients, followed by 11.7% homozygous for Duffy positive and 1.4% negative. This supports the hypothesis that homozygous individuals are less susceptible to P. vivax infection [59]. A recent systematic review also revealed a varied relationship between the Duffy genotype/phenotype and P. vivax infection ranging from no evidence of malaria cases in the Duffy negative genotypes to a higher prevalence (99%) of P. vivax infections in these groups [60]. Another systematic review also confirmed a low prevalence of P. vivax infections among Duffy-negative individuals in East Africa [61]. However, there is limited global evidence to date to substantiate this phenomenon [36]. Our study was limited to addressing the relationship between allelic types and the risk of malaria.
Discovery of the molecular mechanisms of successful infection in Duffy negative and Duffy positive individuals made it clear that Duffy negative individuals are not completely resistant to P. vivax infections, but suggest a role in resistance to clinical disease [62]. The high proportion of Duffy positives may serve as a potential source of P. vivax infection for Duffy negatives [24]. The high frequency of this phenotype in our population coupled with P. vivax endemicity demands tailored interventions targeting P. vivax. Yet, it is vital to examine the transmission pathways and the genetic diversity of P. vivax among Duffy positive and Duffy negative populations.
The high proportion of P. vivax infections with multiple PvDBP copies in this study supported by previous studies, confirmed that multi-copy PvDBP infections are common in Ethiopia [31,36,63]. Gene duplication can generate new gene functions or alter gene expression pathways [64]. We have observed differences between the study sites which might be linked with the parasites’ genetic diversity. The highest PvDBP CNV was reported from Arba Minch, where a recent study demonstrated the highest haplotype and nucleotide diversity of P. vivax [65]. A higher proportion of multiple PvDBP copy parasites was observed in symptomatic patients compared to the asymptomatic participants, supported by a previous report [36]. As it is well known the parasite uses this ligand-receptor interaction to establish infection in RBCs and limited or enhanced availability or binding of the two pairs might play an important role in determining the parasitemia level. In our previous study including a community cohort where we followed asymptomatic infected participants over a year, some infections progressed to clinical high density infections whilst others remained around the limit of detection (low density infections) throughout the study [66,67]. The inclusion of asymptomatic community members from all study sites could have provided important insights. Our study was limited to include asymptomatic individuals only from one of the study sites. Future investigation with appropriate study design that includes the asymptomatic community might support generating a definitive conclusion.
In our study, we observed higher PvDBP copies in patients with Duffy positive genotypes compared with Duffy negatives although there were very few Duffy negative observations. Similar PvDBP expansion were detected at lower frequencies in Cambodia, Brazil, and India where a small proportion of Duffy negative individuals live [31]. Debatably, PvDBP copy numbers may not be selected in response to the Duffy-negativity barrier [42]. No specific PvDBP sequence polymorphisms that are associated with Duffy negative erythrocyte invasion by the parasites were reported [68]. On the contrary, higher CNVs of PvDBP1 were observed in Duffy negative than positive [32]. Further studies may elucidate the significance of PvDBP CNV in Duffy positive and Duffy negative erythrocyte invasion mechanisms.
Duffy positive P. vivax-infected participants had high parasitemia compared with Duffy negatives in this study, in line with other reports from Ethiopia [20,22,33] and elsewhere [21,69]; whilst another report from western Ethiopia reported conflicting results [70]. The high parasitemia observed in the under-five children which might be linked with premature immunity at an early age suggests an important role they might play in sustaining the transmission reservoir [71]. A significant number of P. vivax infections with high parasitemia levels could be a challenge for malaria elimination programs targeting P. vivax. On the other hand, the observed low parasite load in Duffy negative individuals might suggest low invasion competence of P. vivax [27,33], despite small Duffy negative samples. Parasite invasion of reticulocytes is determined to be reduced in the absence of the Duffy antigen expression [72]. Our data noted that all of the infections detected in Duffy negative individuals were not missed by microscopy albeit low density. Molecular methods might provide better estimates of the silent P. vivax parasites reservoir that could remain undetected by routine diagnostic tests (microscopy and RDT). P. vivax could survive in Duffy negative individuals without eliciting any symptoms, but the importance of these infections to the infectious reservoir needs further investigation [73]. Based on the low parasite densities detected in this study, Duffy negative vivax patients may be less infectious to mosquitoes but their relative contribution remains to be quantified [22,33].
Within the Duffy positive population, we observed higher parasitemia in heterozygous than homozygous individuals contrasting with other studies [20,22,33], which reported a comparable range of parasitemia between the two groups. This deviation could be due to study site-level variations of parasitemia and Duffy genotype in our study. For instance, in Batu, where all of the patients were heterozygous, the maximum median parasitemia was detected. Contrary to the above studies, a previous study in Ethiopia reported that gametocytemia levels in heterozygous individuals were found to be relatively higher than in homozygous groups [74]. Further investigations are required if these are biologically relevant. Taking into account previous studies that did not find an association between parasitemia and PvDBP copy number [31,36], several lines of arguments might explain the observation in this study. Considering that most infections had multiple copies of PvDBP, the interaction between DARC and PvDBP can be enhanced and that might explain the higher parasitemia we observed in the heterozygous population. Until very recently, the interaction between DARC and PvDBP was solely hypothesized to be behind an invasion of reticulocytes. No evidence exists to date that polymorphisms in the PvRBP2b and CD71 play any role in the successful invasion of reticulocytes and subsequently parasitemia.
Despite the majority of the study participants being Duffy positive, infections have been detected in Duffy negative individuals as well; these were low density in general. The high prevalence of multi-copy PvDBP observed among Ethiopian P. vivax isolates, combined with the high rate of Duffy positivity, that were directly related to the observed parasitemia levels corroborates the fact that the parasite evolved to maximize transmission success. These could challenge the malaria elimination efforts of the country. Considering that gametocyte production mirrors asexual parasitemia in P. vivax, high density infections are likely to produce more gametocytes and contribute substantially to the maintenance of transmission in endemic settings. In addition, P. vivax could survive in Duffy negative individuals without eliciting any symptoms, but the importance of these infections to the infectious reservoir needs further investigation. In general, the relative public health importance of P. vivax infections in Duffy positive and Duffy negative individuals, the impact of PvDBP CNVs, and their relative contribution to the maintenance of the infectious reservoir needs to be investigated in detail in natural infections.
Supporting information
S1 Fig. A scatter plot of parasitemia versus age with an overlay of a non-linear LOWESS line (red line).
The plot represents the relationship between the variables visually, with the non-linear relationship being represented by the line.
https://doi.org/10.1371/journal.pntd.0012837.s001
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S2 Fig. Diagnostic plots for assessing linearity assumptions and fitting models (lm model).
These plots can provide insights into the relationship between the parasitemia and the predictor variables (Age, PvDBP copy number, Duffy blood group). Residuals vs. Fitted: This plot shows the differences between observed and predicted values (on the y-axis and the fitted values on the x-axis). This helps identify patterns and non-linearities in the relationship between the response and predictors. A good fit is indicated by residuals that are randomly distributed around zero. Normal Q-Q (Q-Q Residuals) Plot: This graph compares the quantiles of standardized residuals to those of a normal distribution. It helps determine whether the residuals have a normal distribution. If the points roughly follow a straight line, the assumption of normality is probably met. Scale-Location Plot: This plot assists in checking for constant variance (homoscedasticity) of residuals. Ideally, the points should be randomly scattered around a horizontal line. Residuals vs. Leverage: This plot helps to identify influential observations that have a large impact on the model fit. Observations with high leverage and large residuals may have a strong influence on the estimated coefficients.
https://doi.org/10.1371/journal.pntd.0012837.s002
(TIF)
S3 Fig. Diagnostic plots for assessing fitting models (GAM model).
Observed Vs Predicted: This plot shows the observed parasitemia against the fitted values from the GAM model. The y-axis represents the observed response, and the x-axis represents the fitted values. It helps assess the overall fit of the model. Ideally, the points should fall along a straight line, indicating that the model captures the relationship between the predictors and the response variable. Any systematic deviations from the line could suggest issues with the model’s ability to capture the underlying patterns in the data.
https://doi.org/10.1371/journal.pntd.0012837.s003
(TIF)
S1 Table. The trend of confirmed P. vivax and P. falciparum cases in each study site between 2018 and 2022.
https://doi.org/10.1371/journal.pntd.0012837.s004
(XLSX)
S2 Table. PvDBP duplication referring to age category, sex, infection type, Duffy blood group, and site Ethiopia (N = 435).
https://doi.org/10.1371/journal.pntd.0012837.s005
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S3 Table. Result of the ordinal logistic regression model for significantly associated variables with PvDBP copy number, Ethiopia (N = 435).
https://doi.org/10.1371/journal.pntd.0012837.s006
(XLSX)
Acknowledgments
We would like to acknowledge the study participants, the field research team, the community facilitators, and regional and district health officers for their support during sample collection and transportation.
References
- 1.
WHO. World Malaria Report 2023. Geneva: World Health Organization; 2023.
- 2. Howes RE, Battle KE, Mendis KN, Smith DL, Cibulskis RE, Baird JK, et al. Global epidemiology of Plasmodium vivax. Am J Trop Med Hyg. 2016;95(6 Suppl):15–34. pmid:27402513
- 3. Rosenberg R. Plasmodium vivax in Africa: hidden in plain sight?. Trends Parasitol. 2007;23(5):193–6. pmid:17360237
- 4. Twohig K, Pfeffer D, Baird K, Price R, Zimmerman P, Hay S, et al. Growing evidence of Plasmodium vivax across malaria-endemic Africa. PLoS Negl Trop Dis. 2019;13(1):e0007140.
- 5. Howes RE, Patil AP, Piel FB, Nyangiri OA, Kabaria CW, Gething PW, et al. The global distribution of the Duffy blood group. Nat Commun. 2011;2(1):266. pmid:21468018
- 6. Gunalan K, Niangaly A, Thera MA, Doumbo OK, Miller LH. Plasmodium vivax infections of Duffy-negative erythrocytes: historically undetected or a recent adaptation?. Trends Parasitol. 2018;34(5):420–9. pmid:29530446
- 7. Ketema T, Bacha K, Getahun K, Portillo HA, Bassat Q. Plasmodium vivax epidemiology in Ethiopia 2000-2020: a systematic review and meta-analysis. PLoS NeglTrop Dis. 2021;15(9):e0009781. pmid:34525091
- 8. Sintasath DM, Ghebremeskel T, Lynch M, Kleinau E, Bretas G, Shililu J, et al. Malaria prevalence and associated risk factors in Eritrea. Am J Trop Med Hyg. 2005;72(6):682–7 pmid:15964950
- 9. Seyfarth M, Khaireh BA, Abdi AA, Bouh SM, Faulde MK. Five years following first detection of Anopheles stephensi (Diptera: Culicidae) in Djibouti, Horn of Africa: populations established-malaria emerging. Parasitol Res. 2019;118(3):725–32. pmid:30671729
- 10. Bousema T, Youssef RM, Cook J, Cox J, Alegana VA, Amran J, et al. Serologic markers for detecting malaria in areas of low endemicity, Somalia, 2008. Emerg Infect Dis. 2010;16(3):392–9. pmid:20202412
- 11. Elgoraish AG, Elzaki SEG, Ahmed RT, Ahmed AI, Fadlalmula HA, Abdalgader Mohamed S, et al. Epidemiology and distribution of Plasmodium vivax malaria in Sudan. Trans R Soc Trop Med Hyg. 2019;113(9):517–24. pmid:31162590
- 12. Taffese HS, Hemming-Schroeder E, Koepfli C, Tesfaye G, Lee M-c, Kazura J, et al. Malaria epidemiology and interventions in Ethiopia from 2001 to 2016. Infect Dis Poverty. 2018;7(1):1–9.
- 13. Armstrong J. Susceptibility to vivax malaria in Ethiopia. Trans R Soc Trop Med Hyg. 1978;72(4):342–4.
- 14. Mathews H, Armstrong J. Duffy blood types and vivax malaria in Ethiopia. Am J Trop Med Hyg. 1981;30(2):299–303.
- 15. Chaudhuri A, Polyakova J, Zbrzezna V, Williams K, Gulati S, Pogo AO. Cloning of glycoprotein D cDNA, which encodes the major subunit of the Duffy blood group system and the receptor for the Plasmodium vivax malaria parasite. Proc Natl Acad Sci USA. 1993;90(22):10793–7. pmid:8248172
- 16. Ovchynnikova E, Aglialoro F, Bentlage AE, Vidarsson G, Salinas ND, von Lindern M, et al. DARC extracellular domain remodeling in maturating reticulocytes explains Plasmodium vivax tropism. Blood. 2017;130(12):1441–4.
- 17. Gruszczyk J, Kanjee U, Chan L-J, Menant S, Malleret B, Lim NT, et al. Transferrin receptor 1 is a reticulocyte-specific receptor for Plasmodium vivax. Science. 2018;359(6371):48–55.
- 18. Malleret B, Li A, Zhang R, Tan KS, Suwanarusk R, Claser C, et al. Plasmodium vivax: restricted tropism and rapid remodeling of CD71-positive reticulocytes. Blood. 2015;125(8):1314–24. pmid:25414440
- 19. Miller L, Mason S, Clyde D, McGinniss M. The resistance factor to Plasmodium vivax in blacks. The Duffy-blood-group genotype, FyFy. N Engl J Med. 1976;295(6):302–4.
- 20. Abebe A, Bouyssou I, Mabilotte S, Dugassa S, Assefa A, Juliano JJ, et al. Potential hidden Plasmodium vivax malaria reservoirs from low parasitemia Duffy-negative Ethiopians: molecular evidence. PLoS NeglTrop Dis. 2023;17(7):e0011326. pmid:37399221
- 21. Ahmed S, Pestana K, Ford A, Elfaki M, Gamil E, Elamin AF, et al. Prevalence and distribution of Plasmodium vivax Duffy Binding Protein gene duplications in Sudan. PLoS One. 2023;18(7):e0287668. pmid:37471337
- 22. Lo E, Russo G, Pestana K, Kepple D, Abagero BR, Dongho GBD, et al. Contrasting epidemiology and genetic variation of Plasmodium vivax infecting Duffy-negative individuals across Africa. Int J Infect Dis. 2021;108:63–71. pmid:33991680
- 23. Zimmerman PA. Plasmodium vivax infection in Duffy-negative people in Africa. Am J Trop Med Hyg. 2017;97(3):636–8. pmid:28990906
- 24. Ménard D, Barnadas C, Bouchier C, Henry-Halldin C, Gray LR, Ratsimbasoa A, et al. Plasmodium vivax clinical malaria is commonly observed in Duffy-negative Malagasy people. Proc Natl Acad Sci USA. 2010;107(13):5967–71. pmid:20231434
- 25. King CL, Adams JH, Xianli J, Grimberg BT, McHenry AM, Greenberg LJ, et al. Fya/Fyb antigen polymorphism in human erythrocyte Duffy antigen affects susceptibility to Plasmodium vivax malaria. Proc Natl Acad Sci U S A. 2011;108(50):20113–8.
- 26. Bouyssou I, El Hoss S, Doderer-Lang C, Schoenhals M, Rasoloharimanana LT, Vigan-Womas I, et al. Unveiling P. vivax invasion pathways in Duffy-negative individuals. Cell Host Microbe. 2023;31(12):2080–92.e5. pmid:38056460
- 27. Dechavanne C, Dechavanne S, Metral S, Roeper B, Krishnan S, Fong R, et al. Duffy Antigen Expression in Erythroid Bone Marrow Precursor Cells of Genotypically Duffy Negative Individuals. bioRxiv. 2018;508481.
- 28. Golassa L, Amenga-Etego L, Lo E, Amambua-Ngwa A. The biology of unconventional invasion of Duffy-negative reticulocytes by Plasmodium vivax and its implication in malaria epidemiology and public health. Malar J. 2020;19(1):1–10.
- 29. Kepple D, Hubbard A, Ali MM, Abargero BR, Lopez K, Pestana K, et al. Plasmodium vivax from Duffy-negative and Duffy-positive individuals share similar gene pools in east Africa. J Infect Dis. 2021;224(8):1422–31. pmid:33534886
- 30. Popovici J, Roesch C, Rougeron V. The enigmatic mechanisms by which Plasmodium vivax infects Duffy-negative individuals. PLoS Pathog. 2020;16(2):e1008258. pmid:32078643
- 31. Hostetler JB, Lo E, Kanjee U, Amaratunga C, Suon S, Sreng S, et al. Independent origin and global distribution of distinct Plasmodium vivax Duffy binding protein gene duplications. PLoS NeglTrop Dis. 2016;10(10):e0005091. pmid:27798646
- 32. Gunalan K, Lo E, Hostetler JB, Yewhalaw D, Mu J, Neafsey DE, et al. Role of Plasmodium vivax Duffy-binding protein 1 in invasion of Duffy-null Africans. Proc Natl Acad Sci USA. 2016;113(22):6271–6. pmid:27190089
- 33. Abate A, Bouyssou I, Mabilotte S, Doderer-Lang C, Dembele L, Menard D, et al. Vivax malaria in Duffy-negative patients shows invariably low asexual parasitaemia: implication towards malaria control in Ethiopia. Malar J. 2022;21(1):230. pmid:35915453
- 34. Pimpat Y, Leelawat K, Lek-Uthai U. Red blood cell Duffy antigen receptor for chemokines and susceptibility to Plasmodium vivax infection in Thais. Southeast Asian J Trop Med Public Health. 2016;47(5):885–9 pmid:29620340
- 35. Kasehagen LJ, Mueller I, Kiniboro B, Bockarie MJ, Reeder JC, Kazura JW, et al. Reduced Plasmodium vivax erythrocyte infection in PNG Duffy-negative heterozygotes. PLoS One. 2007;2(3):e336. pmid:17389925
- 36. Lo E, Hostetler JB, Yewhalaw D, Pearson RD, Hamid MM, Gunalan K, et al. Frequent expansion of Plasmodium vivax Duffy Binding Protein in Ethiopia and its epidemiological significance. PLoS NeglTrop Dis. 2019;13(9):e0007222.
- 37.
Puji BSA, Din S, John Kevin B. Challenges in the control and elimination of Plasmodium vivax Malaria. In: Sylvie M, Vas D, editors. Towards Malaria Elimination. Rijeka: IntechOpen; 2018. p. Ch. 4.
- 38. Woldearegai TG, Kremsner PG, Kun JF, Mordmüller B. Plasmodium vivax malaria in Duffy-negative individuals from Ethiopia. Trans R Soc Trop Med Hyg. 2013;107(5):328–31. pmid:23584375
- 39. Emiru T, Getachew D, Murphy M, Sedda L, Ejigu LA, Bulto MG, et al. Evidence for a role of Anopheles stephensi in the spread of drug and diagnosis-resistant malaria in Africa. Nat Med. 2023;29(12):3203–11.
- 40. Wampfler R, Mwingira F, Javati S, Robinson L, Betuela I, Siba P, et al. Strategies for detection of Plasmodium species gameocytes. PLoS One. 2013;8(9):e76316. pmid:24312682
- 41. Hermsen CC, Telgt DS, Linders EH, Locht L, Eling WM, Mensink EJ, et al. Detection of Plasmodium falciparum malaria parasites in vivo by real-time quantitative PCR. Mol Biochem Parasitol. 2001;118(2):247–51.
- 42. Roesch C, Popovici J, Bin S, Run V, Kim S, Ramboarina S, et al. Genetic diversity in two Plasmodium vivax protein ligands for reticulocyte invasion. PLoS NeglTrop Dis. 2018;12(10):e0006555. pmid:30346980
- 43.
Available from: https://github.com/solsisay/Plasmodium-vivax-Duffy-binding-protein-copy-number-variation-and-Duffy-genotype
- 44. Tadesse, Fitsum G. Spatial distribution of Plasmodium vivax Duffy binding protein copy number variation and Duffy genotype, and their association with parasitemia in Ethiopia [Dataset]. Dryad. https://doi.org/10.5061/dryad.7pvmcvf40.
- 45.
WHO. Global technical strategy and targets for malaria 2016–2030; 2024.
- 46. Assefa A, Fola AA, Tasew G. Emergence of Plasmodium falciparum strains with artemisinin partial resistance in East Africa and the Horn of Africa: is there a need to panic?. Malar J. 2024;23(1):34. pmid:38273360
- 47. Mihreteab S, Platon L, Berhane A, Stokes BH, Warsame M, Campagne P, et al. Increasing prevalence of artemisinin-resistant HRP2-negative malaria in Eritrea. N Engl J Med. 2023;389(13):1191–202. pmid:37754284
- 48. Fola AA, Feleke SM, Mohammed H, Brhane BG, Hennelly CM, Assefa A, et al. Plasmodium falciparum resistant to artemisinin and diagnostics have emerged in Ethiopia. Nat Microbiol. 2023;8(10):1911–9. pmid:37640962
- 49. Ahmed A, Pignatelli P, Elaagip A, Abdel Hamid MM, Alrahman OF, Weetman D. Invasive malaria vector Anopheles stephensi mosquitoes in Sudan, 2016-2018. Emerg Infect Dis. 2021;27(11):2952–4. pmid:34670658
- 50. Ahmed A, Khogali R, Elnour M-AB, Nakao R, Salim B. Emergence of the invasive malaria vector Anopheles stephensi in Khartoum State, Central Sudan. Parasit Vectors. 2021;14(1):511.
- 51. Hawaria D, Kibret S, Zhong D, Lee M-C, Lelisa K, Bekele B, et al. First report of Anopheles stephensi from southern Ethiopia. Malar J. 2023;22(1):373. pmid:38066610
- 52. Liu Q, Wang M, Du Y-T, Xie J-W, Yin Z-G, Cai J-H, et al. Possible potential spread of Anopheles stephensi, the Asian malaria vector. BMC Infect Dis. 2024;24(1):333. pmid:38509457
- 53. Yared S, Gebresilassie A, Aklilu E, Abdulahi E, Kirstein OD, Gonzalez-Olvera G, et al. Building the vector in: construction practices and the invasion and persistence of Anopheles stephensi in Jigjiga, Ethiopia. Lancet Planet Health. 2023;7(12):e999–e1005. pmid:38056970
- 54. Ryan SJ, Lippi CA, Zermoglio F. Shifting transmission risk for malaria in Africa with climate change: a framework for planning and intervention. Malar J. 2020;19(1):1–14.
- 55.
NMEP. Ethiopia Malaria Elimination Strategic Plan: 2021-2025. Addis Ababa: Ministry of Health, 2020.
- 56. Miller LH, Mason SJ, Dvorak JA, McGinniss MH, Rothman IK. Erythrocyte receptors for (Plasmodium knowlesi) malaria: Duffy blood group determinants. Science. 1975;189(4202):561–3. pmid:1145213
- 57. Dalgleish T, Williams J, Golden A, Perkins N, Barrett L, Barnard P. Dsitribution of blood groups in the east African Somali Population. J Exp Psychol Gen. 2007;136(1):23–42.
- 58. Abdelraheem MH, Albsheer MM, Mohamed HS, Amin M, Abdel Hamid MM. Transmission of Plasmodium vivax in Duffy-negative individuals in central Sudan. Trans R Soc Trop Med Hyg. 2016;110(4):258–60.
- 59. Lo E, Yewhalaw D, Zhong D, Zemene E, Degefa T, Tushune K, et al. Molecular epidemiology of Plasmodium vivax and Plasmodium falciparum malaria among Duffy-positive and Duffy-negative populations in Ethiopia. Malar J. 2015;14(1):1–10.
- 60. Picón-Jaimes YA, Lozada-Martinez ID, Orozco-Chinome JE, Molina-Franky J, Acevedo-Lopez D, Acevedo-Lopez N, et al. Relationship between Duffy Genotype/Phenotype and prevalence of Plasmodium vivax Infection: a systematic review. Trop Med Infect Dis. 2023;8(10):463. pmid:37888591
- 61. Wilairatana P, Masangkay FR, Kotepui KU, De Jesus Milanez G, Kotepui M. Prevalence and risk of Plasmodium vivax infection among Duffy-negative individuals: a systematic review and meta-analysis. Sci Rep. 2022;12(1):3998. pmid:35256675
- 62. Ryan J, Stoute J, Amon J, Dunton R, Mtalib R, Koros J, et al. Evidence for transmission of Plasmodium vivax among a duffy antigen negative population in Western Kenya. Am J Trop Med Hyg. 2006;75(4):575.
- 63. Auburn S, Getachew S, Pearson RD, Amato R, Miotto O, Trimarsanto H, et al. Genomic analysis of Plasmodium vivax in southern Ethiopia reveals selective pressures in multiple parasite mechanisms. J Infect Dis. 2019;220(11):1738–49. pmid:30668735
- 64. Lynch M, Conery JS. The evolutionary demography of duplicate genes. J Struct Funct Genomics. 2003;3(1-4):35–44 pmid:12836683
- 65. Abebe A, Dieng CC, Dugassa S, Abera D, Shenkutie TT, Assefa A, et al. Genetic differentiation of Plasmodium vivax duffy binding protein in Ethiopia and comparison with other geographical isolates. Malar J. 2024;23(1):55. pmid:38395885
- 66. Hailemeskel E, Tebeje SK, Ramjith J, Ashine T, Lanke K, Behaksra SW, et al. Dynamics of asymptomatic Plasmodium falciparum and Plasmodium vivax infections and their infectiousness to mosquitoes in a low transmission setting of Ethiopia: a longitudinal observational study. Int J Infect Dis. 2024;143:107010. pmid:38490637
- 67. Tadesse FG, Slater HC, Chali W, Teelen K, Lanke K, Belachew M, et al. The relative contribution of symptomatic and asymptomatic Plasmodium vivax and Plasmodium falciparum infections to the infectious reservoir in a low-endemic setting in Ethiopia. Clin Infect Dis. 2018;66(12):1883–91. pmid:29304258
- 68. Hoque MR, Elfaki MMA, Ahmed MA, Lee S-K, Muh F, Ali Albsheer MM, et al. Diversity pattern of Duffy binding protein sequence among Duffy-negatives and Duffy-positives in Sudan. Malar J. 2018;17(1):1–10.
- 69. Albsheer MMA, Hussien A, Kwiatkowski D, Hamid MMA, Ibrahim ME. The Duffy T-33C is an insightful marker of human history and admixture. Meta Gene. 2020;26:100782.
- 70. Bradley L, Yewhalaw D, Hemming-Schroeder E, Jeang B, Lee M-C, Zemene E, et al. Epidemiology of Plasmodium vivax in Duffy negatives and Duffy positives from community and health centre collections in Ethiopia. Malar J. 2024;23(1):76. pmid:38486245
- 71. Doolan DL, Dobaño C, Baird JK. Acquired immunity to malaria. Clin Microbiol Rev. 2009;22(1):13–36. pmid:19136431
- 72. Albsheer MM, Pestana K, Ahmed S, Elfaki M, Gamil E, Ahmed SM, et al. Distribution of Duffy phenotypes among Plasmodium vivax infections in Sudan. Genes. 2019;10(6):437.
- 73. Little E, Shenkutie TT, Negash MT, Abagero BR, Abebe A, Popovici J, et al. Prevalence and characteristics of Plasmodium vivax Gametocytes in Duffy-Positive and Duffy-Negative Populations across Ethiopia. Am J Trop Med Hyg. 2024;110(6):1091–9. pmid:38626749
- 74. Abate A, Hassen J, Dembele L, Menard D, Golassa L. Differential transmissibility to Anopheles arabiensis of Plasmodium vivax gametocytes in patients with diverse Duffy blood group genotypes. Malar J. 2023;22(1):136. pmid:37098534