Until recently, little or no endemic transmission of Plasmodium vivax has been reported in sub-Saharan Africa outside of the Horn of Africa (1). P. vivax was presumed to be largely absent because the Duffy blood group antigen was rare in persons living in the region. However, accumulating evidence of endemic P. vivax has indicated that this parasite might be present in many areas of sub-Saharan Africa, albeit at low levels, and Duffy antigen–negative persons can be infected and contribute to transmission (2).

Turkana County is in northwestern Kenya and shares a border with Uganda, South Sudan, and Ethiopia. Turkana county’s harsh climate is characterized by an average rainfall of <215 mm/year and daytime temperatures of 40°C. Malaria transmission in this region was predicted to occur in isolated pockets with epidemic potential only after unusual rainfall. However, reactive case detection conducted across central Turkana County documented year-round symptomatic and asymptomatic P. falciparum infections and confirmed perennial endemic transmission of malaria (3).

We hypothesized that P. vivax might also be circulating in Turkana County because of stable malaria transmission and proximity to Ethiopia, where P. vivax infections are endemic. To test this hypothesis, we extracted genomic DNA from 3,305 dried blood spots collected from household members of patients with P. falciparum infections; household members were enrolled in the study at their homes in catchment areas surrounding 3 rural and 3 urban health facilities in central Turkana County (3). The study was approved by the Moi University Institutional Research and Ethics Committee and Duke University Institutional Review Board.

We tested each DNA sample for P. vivax by using an established nested qualitative PCR protocol (4). Gel electrophoresis bands were identified independently by 2 observers. We randomly selected 15 extracts for retesting by probe-based real-time PCR with the same primer sequences to detect the same target; all PCR products were confirmed. For our analysis, we used nested qualitative PCR results.

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Figure. Prevalence of Plasmodium vivax infection in communities along the Turkwel River in study of P. vivaxprevalence in semiarid region of northern Kenya,...

The percentage of household members infected with P. vivax was 2.1% (69/3,305); of those, 45% (31/69) were co-infected with P. falciparum (Table). We detected P. vivax infections across our study transect throughout most of the year (Figure; Appendix Figures 1, 2); the highest (5.8%, 28/485) prevalence was recorded near an urban facility in the town of Lodwar. Infections were present across all age groups, but we observed a slightly higher (1.6%, 8/490) percentage of P. vivax monoinfections in children <5 years of age (Table). Ten P. vivax–infected participants reported malaria-like symptoms when they were screened; 7 of those were co-infected with P. falciparum. Only 3 P. vivax–infected participants had a malaria-like illness within 1 month before enrollment; none reported taking antimalarial drugs. None of the P. vivax–infected participants reported traveling outside of their subcounty within 2 months before enrollment; 16% (11/69) reported having a net for their sleeping space, which was slightly less than uninfected participants (19.7%, 468/2,376) who had a net.

The burden of P. vivax infections in sub-Saharan Africa remains unclear; infections are rarely diagnosed in a clinical setting and might often be asymptomatic. The recommended rapid diagnostic test in most countries of sub-Saharan Africa is P. falciparum–specific. Consequently, P. vivax infections might be underestimated or undocumented.

Strategies designed to eliminate P. falciparum are undermined by P. vivax because dormant P. vivax hypnozoites that can cause relapse and sustain transmission are difficult to detect and treat (5). Furthermore, P. vivax infections generate gametocytes before symptom onset, making detection and treatment challenging before onward transmission occurs. P. vivax infections could present a growing challenge in Kenya, even as P. falciparum is brought under control, a process that has been observed in co-endemic malaria settings in Southeast Asia (6).

We did not test participants for Duffy antigen expression, which could have affected their susceptibility to P. vivax. Estimated Duffy antigen positivity in Kenya is 5%–10% (7). P. vivax infections in Duffy-negative subjects have been documented in Africa (2). Characterization of Duffy antigen expression will be needed to understand the threat of P. vivax infections in Kenya.

Anopheles stephensi mosquitoes have been identified in Kenya (E.O. Ochomo et al., unpub. data, https://doi.org/10.21203/rs.3.rs-2498485/v1), and the potential expansion of this highly competent vector, which survives in urban and manmade habitats, could dramatically change malaria transmission patterns. Continued spread of this invasive vector into sub-Saharan Africa would place ≈126 million persons at risk for malaria (8). Identification of An. stephensi mosquitoes in Djibouti was linked with a >100-fold rise in malaria cases, including the first autochthonous cases of P. vivax reported in 2016 (9).

In conclusion, if emerging An. stephensi mosquitoes become established across Kenya in the presence of confirmed P. vivax cases, malaria elimination in Kenya will be substantially more difficult to achieve. Enhanced surveillance for both An. stephensi mosquitoes and P. vivax will be needed to inform control measures, and increased clinical resource allocation will enable detection and effective treatment of patients with P. vivax malaria.

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