Skip to main content
NCBI home page
One Health logoLink to One Health
. 2026 Sep 20;23:101575. doi: 10.1016/j.onehlt.2026.101575

Widespread transmission in diverse ecotypes challenges visceral leishmaniasis control in East Africa

Eva Iniguez a,⁎,1, Mercy Tuluso b,1, Steve Kiplagat c, Araya Gebresilassie d, Esayas Aklilu e, Olivia Battistoni f, Johnstone Ingonga b, John Mark Makwatta c, Mohamed Alamin g, Osman Dakien g,h, Alphine Chebet b, Esther Kaunda b, Patrick Huffcutt a, Serena Doh a, Pedro Cecilio i, Galgallo Bonaya c, Claudio Meneses a, Tiago D Serafim a, Myrthe Pareyn j, Mohamed Osman k, Eltahir AG Khalil g, Omran F Osman g, Brima M Younis g, Sithar Dorjee l, Guofa Zhou m, Jesus G Valenzuela a, Dan K Masiga c, Ahmed M Musa n, Asrat Hailu o, Abhay Satoskar f, Shaden Kamhawi a,⁎, Damaris Matoke-Muhia b,⁎⁎
PMCID: PMC13634277  PMID: 42831020

Abstract

East Africa is emerging as the global hot spot of visceral leishmaniasis (VL), yet efforts to eliminate it are hindered by substantial knowledge gaps in its ecoepidemiology. Here, we report on the high prevalence of Leishmania infection in Phlebotomus orientalis in Marsabit county, Kenya (3.9%), and Gedaref state, Sudan (3.6%), where this species comprised 99.8% (n = 1185) and 100% (n = 1350) of captured Phlebotomus females, respectively. In Aba Roba, Ethiopia, Phlebotomus martini accounted for 94.4% of 184 collected Phlebotomus females and had a lower infection rate of 1.5%. Phlebotomus orientalis and Phlebotomus martini exhibited different habitat and feeding preferences. While Phlebotomus orientalis was abundant in diverse peridomestic and sylvatic microhabitats, Phlebotomus martini was predominantly collected from termite hills. Moreover, Phlebotomus orientalis primarily fed on humans and less on domestic and sylvatic animals. In contrast, Phlebotomus martini exhibited zoophagic behavior, mostly feeding on cows and Ovis. Widespread transmission of Leishmania in our study sites is supported by high rK39 seroprevalence in both Kenya (17.9%) and Sudan (6.2%). An observed greater prevalence of antibodies to rK39 in individuals living near than away from VL cases in both Kenya (19.8% versus 7.4%, P = 0.0015) and Sudan (8.4% versus 2.1%, P = 0.0105) demonstrated that proximity to a VL case carries an increased risk of infection. Our findings highlight the need for a risk-based targeted site-specific one health elimination strategy that accounts for the intensity, diversity, and complexity of VL transmission in today's East Africa.

Keywords: East Africa, Visceral leishmaniasis, Sand fly ecology, Leishmania-infected sand flies, rK39 seroprevalence

Graphical abstract

A one health framework is needed to control visceral leishmaniasis transmission across diverse endemic regions in East Africa. Schematic representation of Leishmania donovani transmission cycles in our study sites highlights complex interactions among vectors, humans, animals, vegetation, and the environment. The environment influences vector bionomics in peridomestic and sylvatic settings for Phlebotomus orientalis (wide prevalence across microhabitats including Acacia and Balanites vegetation, vertisol cracks, animal burrows, and indoors), and Ph. martini (restricted to termite hills). Arrows depict transmission cycles involving human and animal populations moving from peridomestic to sylvatic settings (eg. temporary shelters) due to seasonal occupations as pastoralists and farmers. Risk of human infection is increased by proximity to VL cases (peridomestic transmission) or by seasonal migration (possible zoonotic transmission). Study sites: Marsabit county, Kenya (orange); Aba Roba, the Konso Zone, Ethiopia (blue); Gedaref state, Sudan (red). Created in BioRender. Iniguez, E. (2026) https://BioRender.com/oxq4oau.

Unlabelled Image

Open in a new tab

Highlights

  • •

    Ph. orientalis and Ph. martini sand flies show distinct habitat and feeding behavior.

  • •

    Widespread VL transmission confirmed by finding infected sand flies in Kenya, Sudan, and Ethiopia study sites.

  • •

    rK39 serology shows high asymptomatic rates in Kenya and Sudan study sites.

  • •

    Living near VL cases increases infection risk.

  • •

    A one health approach identified distinct microhabitats that act as transmission hotbeds, guiding targeted VL control strategies.

1. Introduction

East Africa (EA) is currently the epicenter of global transmission of visceral leishmaniasis (VL), a fatal neglected disease ranked second after malaria in mortality rates [1], [2]. VL in EA is epidemiologically and clinically diverse [3], [4], [5], and transmitted by several sand fly vector species including Phlebotomus orientalis, Ph. martini, and Ph. celiae that have divergent ecologies and bionomics [6], [7], [8], [9]. VL cases have recently surged in EA, emerging in new foci that disproportionally affected Kenya, Sudan, and Ethiopia, accounting for more than 65% of cases worldwide, mostly children [1], [10].

VL in EA had initially been targeted for control rather than elimination due to significant knowledge gaps [11]. In June 2024, the World Health Organization launched a framework to eliminate VL as a public health concern from EA by 2030 [12]. However, major challenges to VL elimination include differing treatment regimens, low compliance, poor access to care facilities [6], [12], [13], [14], [15], and significant knowledge gaps in ecoepidemiology, vector biology, and nature of reservoirs. Though vector species have been found in strong association with Acacia and Balanites woodlands, vertisol (black cotton soil), and termite hills [16], [17], [18], [19], linking these microhabitats to VL transmission in the region [[18], [19], [20], [21], [22], [23], [24]] needs to be demonstrated. Additionally, we need to determine the prevalence of asymptomatic individuals and their role as infectious reservoirs for sand flies [25], [26], [27]. Further hindering control of VL in EA is the presence of semi-nomadic populations, such as pastoralists and farmers living in temporary shelters [13], and populations displaced by conflict [28], [29], that are at a higher risk of contracting VL [28], [29], [30]. The complicated epidemiological landscape of VL in EA reinforces the need for a one health approach to fully elucidate drivers of transmission. In this study, we provide a comprehensive account of VL transmission dynamics in today's EA.

2. Methods

2.1. Ethics approval

The study was approved by the Scientific Research Unit, KEMRI, and licensed by the National Commission for Science, Technology and Innovation (clinical protocol # KEMRI/SERU/CBRD/249/4634) and by the State Ministry of Health and Social Development at Gedaref state, Sudan ethical committee (clinical protocol # UG/EC/3/2023) in Sudan. Both clinical protocols were translated into local languages. Communal consent was obtained by meeting with community leaders. Written consent was obtained from adults or guardians of minors before blood collection. Verbal consent was acquired from homeowners before entomological collections.

2.2. Study sites

Fig. 1A depicts our endemic study sites in Marsabit county, Kenya [31], [32], the Konso zone, Ethiopia, and Gedaref state, Sudan, selected based on their stable VL endemicity (Fig. 1B).

Fig. 1.

Fig. 1

Open in a new tab

Study sites and incidence of visceral leishmaniasis.

(A) Map of East Africa showing the location of study sites. 1, Marsabit; 2, Konso; 3, Gedaref state. (B) Visceral leishmaniasis (VL) cases per 10,000 population over the past seven years in Marsabit county, Kenya; Konso, Ethiopia; and Gedaref state, Sudan. Incidence data are based on Ministry of Health reports from each country. Maps at QGIS software QGIS shapefile, https://open.africa/dataset/?tags=Shapefiles.

2.2.1. Kenya

Marsabit county has a population of 459,785 people [33], an arid to semi-arid tropical savanna climate with long (April to May) and short rains (November to December), and a temperature range of 15 °C to 26 °C [34]. Vegetation is dominated by Acacia trees. The community moves with their domestic animals from semi-permanent (manyatta) to temporary (fora) houses for up to six months in search of pasture and water.

2.2.2. Sudan

Gedaref state is urban with well-established villages. The rainy season spans June to October, and temperature averages 28 °C. The natural vegetation is a savanna woodland with Acacia and Balanites trees, and the principal soil type is black cotton vertisol. Livelihoods include agriculture needing seasonal labor migrants, and small animal husbandry. Dinder National Park, with a high VL prevalence (27.6%) reported among its wildlife soldiers [35], falls between the Sahel of Gedaref state and the Ethiopian Highlands and supports a diverse wildlife. We investigated the villages of Kunaynah, Sabouni, and Umslala-Houng (closest to Dinder National Park).

2.2.3. Ethiopia

The Konso zone is a semi-arid rugged terrain with hills, lowlands, and riverbeds. The long rains last from February to May, followed by short rains in September and October, and the temperature averages 24 °C. Dispersed homesteads and animal enclosures are scattered in lowlands or valleys, while villages are on hills or highlands. Many families own farms in the valleys and spend nights in temporary shelters (foras) herding cattle, goats, and sheep. Vegetation comprises a mixture of shrub and forests, including Acacia and Balanites trees, and the predominant soil type is sandy. Castellated active or eroded shrinking termite hills [8] are abundant. We investigated the villages of Galga, Goinada, Maira, and Lahalaha.

Unless otherwise specified, we will refer to our study sites as Kenya, Sudan, and Ethiopia.

2.3. Sand fly collection and species identification

Entomological collections were carried out from September 2023 to September 2024. Ecotypes included peridomestic (associated with humans) and sylvatic (away from humans) microhabitats (Supplementary Fig. 1). The total number of CDC light traps used per collection varied in each study site, but the same number of traps were placed per ecotype (Supplementary Table 1).

Phlebotomus sand flies were wet mounted for morphological identification. For identification, spermatheca and pharyngeal teeth were examined in females and external genitalia distinguished males [36], [37].

2.4. Rotation of Phlebotomus male genitalia

During the first 16-24 h post-emergence, the external genitalia rotate to orient the claspers for mating. We recorded unrotated, and partially or fully rotated male genitalia to identify newly emerged juveniles as a proxy to breeding sites [16], [19], [21].

2.5. Extraction of DNA from field-collected Phlebotomus female sand flies

Individual or pools of 5 midguts were collected in Kenya; pools of 10 sand flies (whole bodies or midguts) were collected in Sudan; and midgut samples were individually collected in Ethiopia. Blood-fed sand flies were preserved individually as midguts/whole bodies on Whatman 903 Protein Saver cards (Cytica), or in up to 50 μl of DNA/RNA Shield buffer (Zymo). DNA was extracted from samples using the Quick-DNA/RNA Miniprep Plus kit (Zymo) tissue protocol, or Tissue DNA kit (Omega). All samples were eluted to a final volume of 100 μl and stored at −20/−80 °C until further processing.

2.6. Blood meal analysis of field-collected samples by multiplex PCR

2.6.1. Standardization of a blood meal multiplex custom PCR panel

Female Lutzomyia longipalpis sand flies (4–6-day old) reared at the Laboratory of Malaria and Vector Research, NIAID, NIH, were allowed to feed for 2 h on whole blood from various species through an artificial membrane [38], and were processed 24 h post-feeding as positive controls. Blood was obtained from vendors under LMVR4 animal protocol or IRB human protocol 000331-I (human). Hyrax and camel blood were donated by Dan Masiga, (icipe).

A universal reverse primer [39] was used together with forward primers designed for this study [40], [41], [42] (Supplementary Table 2). Briefly, DNA (40 ng, 10 μl) was mixed with all primers and the DreamTaq™ Hot Start Green PCR Master Mix (Thermo) to a final volume of 30 μl. PCR conditions were 94 °C initial denaturation for 5 min, 35 cycles of 30 s at 94 °C, 30 s at 55 °C, and 60 s at 72 °C, and a final extension of 7 min at 72 °C. PCR products were separated on a 2% agarose gel. Bands were visualized on a c600 Azure Biosystems imager (Supplementary Fig. 3).

2.6.2. Bloodmeal analysis of field-collected samples

DNA from field samples was amplified, separated and visualized under the conditions described above. Samples that did not amplify were rerun using generic vertebrate cytochrome b (cytb) gene primers (0.2 μM) [43]. After confirming amplification by gel electrophoresis, products were purified using the GeneJet PCR purification kit (Thermo) and sequenced (Eurofins). Sequences were aligned with NCBI reference genomes to determine blood meal identity.

2.7. Leishmania detection and quantification by qPCR

A standard curve was created for each different sample condition by spiking an uninfected sand fly midgut, a whole body, or a pool of whole bodies, with 105 L. donovani (MHOM/SD/62/1S) parasites followed by DNA extraction and a 10-fold serial dilution into DNA of corresponding uninfected sample types (Supplementary Fig. 4).

We used qPCR to amplify the Leishmania kinetoplast minicircle using JW11 and JW12 primers [38] and 40 ng of DNA template. For unfed and gravid samples, we used SYBR green (Perfecta SYBR Green Fast Mix or FIREPol Master Mix Ready to Load) following the manufacture's conditions. For blood fed samples, a probe-based qPCR was used as described [38], [44]. Threshold values of 600 and 45 were applied for SYBR-Green and probe-based assays, respectively.

2.8. Human case-control study design and dried blood spot collection

As Ph. orientalis is the vector of L. donovani in both Marsabit, Kenya, and Gedaref, Sudan, we assessed seroprevalence of Leishmania rK39 antibodies in these study foci.

We conducted a case–control study of household members of VL cases, and five neighboring households selected purposively based on proximity to VL cases. For Kenya, non-neighboring controls lived within 10 km from areas of index VL cases. In Sudan, control households were in Sabouni, an adjacent village at 80 km from Kunaynah with a similar ecology that supports sand flies. VL cases were diagnosed and treated at reference hospitals according to national guidelines [12], [45]. Participants were household members meeting inclusion criteria (age ≥ 24 months in Kenya and ≥ 6 months in Sudan).

A total of 752 participants (male to female ratio of 1:1.54) with a median age of 14 years (range, 2–85 years) were enrolled from Laisamis and Karare villages, Marsabit county, Kenya. In Kunaynah village, Gedaref state, Sudan, we enrolled a total of 435 participants (male to female ratio of 1:1.3) with a median age of 15 years (range, 5 months – 85 years). Of those, 108 and 145 were control household members in Kenya and Sudan, respectively.

Dried blood spots (DBSs) were collected from household members of 48 VL cases in Marsabit county, Kenya, and five VL cases in Kunaynah village, Gedaref state, Sudan (Supplementary Table 3) and their corresponding control households. Blood was collected by finger prick using a lancet, spotted on a Whatman 903 protein saver card (Cytiva), and preserved at room temperature.

2.9. Leishmania rK39 ELISA

ELISA plates (Thermo Fisher Scientific) were coated overnight at 4 °C with 25 ng/well of rK39 antigen (donated by Steve Reed, Infectious Disease Research Institute, now the Access to Advanced Health Institute). DBSs were eluted with 300 μl of PBS 0.05% Tween 20 (Sigma) and incubated for 4 h shaking at room temperature or overnight at 4 °C. Plates were blocked with TBST/20% donor horse serum (DHS) for 2 h and 25 μl of the eluted DBS sample was diluted in TBST/5% DHS (1:4 dilution) to a total volume of 100 μl/well in duplicates. Subsequent ELISA steps were completed as described [44]. Non-endemic negative controls obtained from individuals living in Nairobi, Kenya, were included in each plate for normalization.

2.10. Statistical analysis

A chi-square goodness-of-fit test was performed for the entomology data, followed by post-hoc pairwise comparisons using standardized residual (Z-score) analysis with Bonferroni correction to identify differences among ecotypes within each country. A Bonferroni-adjusted critical value of |Z| 2.69 (corresponding to P value ≤0.007) was considered significant. ELISA data was normalized using the plate cutoff calculated by the mean optical density (O.D.) of non-endemic negative controls +5 SD, followed by log transformation. After normalization, samples were statistically compared using the Mann-Whitney test. Odds ratio was used to assess the risk for having rK39 antibodies (categorized as positive or negative) between endemic index case household members and neighborhood households versus non-neighboring households (Kenya), or non-neighborhood controls (Sudan), and its significance was tested using Fisher's exact test. A univariable binary logistic regression was performed to assess the effect of distance in meters from index case households to neighboring versus non-neighboring households on rK39 seropositivity (risk by log-odds of positive outcome after normalization). The ‘margins’ command of Stata was used to compute adjusted log-odds of seropositivity at selected distances from index case households. A P value of ≤0.05 was considered significant.

3. Results

3.1. Distinct microhabitat and breeding preferences of vector species

Ph. orientalis was the predominant VL vector species in Kenya, and Sudan, comprising 99.8% (n = 1185) and 100% (n = 1350) of Phlebotomus females, respectively (Fig. 2A). In contrast, Ph. martini/Ph. celiae females (morphologically indistinguishable) were the most prevalent of five Phlebotomus species in Ethiopia, accounting for 94.4% of the collection (n = 184; Fig. 2A).

Fig. 2.

Fig. 2

Open in a new tab

Distribution of different sand fly species across distinct microhabitats in endemic foci in Kenya, Sudan and Ethiopia study sites.

(A) Relative abundance of collected Phlebotomus females by species. (B) Relative abundance of Phlebotomus females by ecotype.

There was a significant difference in habitat distribution across all study sites (x2 = 1352.9, d.f. = 12, P < 0.0001). Ph. orientalis females in Kenya, were mainly collected from outdoor habitats associated with peridomestic and sylvatic vegetation (45.0%, P < 0.0001), mostly Acacia trees, followed by animal enclosures (25.9%, P < 0.0001; Fig. 2B). In Sudan, Ph. orientalis was primarily captured in vegetation (31.8%, P < 0.0001) and inside houses (27.2%, P < 0.0001). In Ethiopia, Ph. martini/Ph. celiae females were mainly captured from termite hills (74.3%, P < 0.0001) that were prevalent in both peridomestic and sylvatic ecotypes, followed by temporary shelters (8.2%; Fig. 2B).

Ph. orientalis represented 100% of all collected males in Kenya and Sudan (Supplementary Fig. 2 A), and were mostly collected from habitats where females predominated (Supplementary Fig. 2B). Preliminary evidence of breeding sites, identified by males having unrotated or partially rotated genitalia, implicated outdoor ecotypes for both Kenya and Sudan (Supplementary Fig. 2C), potentially indicating that Ph. orientalis may not be breeding indoors in either country.

In Ethiopia, Ph. martini was the main male species, and was collected mostly from termite hills, including 72.9% with unrotated or partially rotated genitalia (Supplementary Fig. 2 A-C). This implicates termite hills as a significant breeding site for Ph. martini. The relative abundance of Ph. martini (93.2%) over Ph. celiae (5.8%) males indicates predominance of the former species in our female collections. As such, we will use Ph. martini instead of Ph. martini/Ph. celiae for females from here on.

3.2. Vector species exhibit different feeding preferences

The multiplex PCR panel, tailored to amplify blood from humans and seven animals present in our study sites (Supplementary Fig. 3, Supplementary Table 2), identified 63.8% (n = 270/423) of blood meals. After cytb gene sequencing of unknowns, this increased to 96.9%. Diversity of blood meal sources was considerably higher in Kenya, compared to either Sudan, or Ethiopia (Fig. 3A-C). Sand flies with multiple blood meals were higher than expected for Kenya, (x2 = 44.1, d.f. = 15, P < 0.0001) and Sudan (x2 = 17.8, d.f. = 6, P < 0.0067), and several females contained mixed human and domestic or sylvatic animal blood meals (Fig. 3A-C). Humans were identified as the dominant host for Ph. orientalis in both Kenya, and Sudan, representing 35.8% (P < 0.0001) and 74.8% (P < 0.0001) of single blood meals, and 74.6% and 100% of mixed blood meals, respectively (Fig. 3A,B). Of the collected 12 Ph. martini blood fed specimens in Ethiopia, most fed on animals, suggesting a more zoophagic behavior (Fig. 3C). Blood fed females were collected from the same ecotypes as unfed/gravid sand flies (Fig. 3D).

Fig. 3.

Fig. 3

Open in a new tab

A mosaic of blood meal sources observed in leishmaniasis vectors across endemic foci in Kenya, Sudan and Ethiopia study sites.

(A-C) Host blood meal identity of blood fed females in Kenya (A), Sudan (B), and Ethiopia (C) with a single or multiple sources of blood detected. (D) Relative abundance of collected blood fed Phlebotomus females by ecotype.

3.3. Leishmania-infected vector species were recovered from diverse microhabitats

Using qPCR (Supplemental Fig. 4) on individual (ni) or pooled (np) sand flies, we detected Leishmania-infected specimens in both peridomestic and sylvatic niches across all study sites (Fig. 4A), and from distinct ecotypes (Fig. 4B-C). The minimal Leishmania infection rate for Ph. orientalis was similar in Kenya, (26/648, 3.9%) and Sudan, (10/281, 3.6%), and lower for Ph. martini (3/183, 1.5%) in Ethiopia (Fig. 4A). None of the other species were infected.

Fig. 4.

Fig. 4

Open in a new tab

Distribution of Leishmania-infected sand flies from distinct and diverse microhabitats reveals hotspots of active transmission near and away human habitation.

(A) Number of Leishmania-infected sand flies collected from peridomestic or sylvatic habitats. (B) Habitat productivity of infected unfed/gravid female Ph. orientalis sand flies in Kenya and Sudan, and Ph. martini/Ph. celiae in Ethiopia. (C) Host blood meal identity of infected blood fed Ph. orientalis and Ph. martini/Ph. celiae females. Specimens were processed individually or as pools of up to 5–10 sand flies (B) or individually (C).

The majority of Leishmania-infected Ph. orientalis were collected from vegetation in sylvatic sites for Kenya, compared to human-associated microhabitats in Sudan (Fig. 4B-C, Supplementary Tables 1 and 4), though in both study sites, they fed predominantly on humans (Fig. 4C, Supplementary Tables 4 and 5), confirming the vector's anthropophilic behavior. Notably, three and one infected sand flies from Kenya fed on gazelles and a rodent, respectively (Fig. 4C, Supplementary Tables 4 and 5). In Ethiopia, Leishmania-infected Ph. martini (1.8%, n = 3/171) were all captured in termite hills (Fig. 4B). Highlighting the focality of VL transmission in all study sites, we found clusters of Leishmania-infected Ph. orientalis around VL patients and recovered two infected Ph. martini females nine months apart from the same castellated termite hill (Supplementary Tables 1 and 4).

3.4. Leishmania rK39 seroprevalence reveals an increased risk of infection for individuals living near VL cases

We conducted a case-control study of household members of VL cases (Supplementary Table 3) and five neighboring households for comparison against non-neighboring controls.

For Kenya and Sudan, rK39 titers were significantly higher at 19.8% (118/596) and 8.4% (24/285) for participants living with or near VL cases, compared to 7.4% (8/108) and 2.1% (3/145) for those living farther away, respectively (Fig. 5 A-B). The odds ratio calculated by Fisher's test for those living with or near a VL case was 3.1 (95%CI = 1.5–6.4, P = 0.0015) and 4.4 (95%CI = 1.4–12.6, P = 0.0105) for Kenya and Sudan, respectively. Additionally, logistic regression demonstrated that the likelihood of developing rK39 antibodies decreases significantly with distance from a VL case for both Kenya (Fig. 5C; Likelihood ratio = 19.9, P < 0.0001; z = −3.1, P = 0.0017) and Sudan (Fig. 5D; Likelihood ratio = 7.2, P = 0.0072; z = −2.3, P = 0.0221). The same outcome was observed when the analysis excluded VL index case household members (Supplemental Fig. 5 A-B). Of note, though serum was collected within 1–4 months from diagnosis and treatment, only 22/48 (Kenya) and none (Sudan) of the diagnosed and treated VL patients had detectable antibodies by rK39 ELISA (Supplementary Table 3).

Fig. 5.

Fig. 5

Open in a new tab

Prevalence of Leishmania rK39 antibodies in our study sites indicate high risk of infection in Marsabit county, Kenya, and Gedaref state, Sudan.

(A,B) Leishmania rK39 antibody prevalence measured from dry blood spots (DBS) collected from individuals living with or near and away from recently reported VL cases in Laisamis and Karare, Marsabit county (A) and Kunaynah, Gedaref state (B). Data was normalized using the plate cutoff calculated by the mean optical density (O.D.) of non-endemic negative controls +5 SD. Non-endemic negative controls were obtained from individuals living in Nairobi, Kenya. Samples with O.D. ≥0.00 after normalization were considered positive. (C,D) Univariable binary logistic regression analysis was performed to assess the effect of distance (meters) from index case households to neighboring and control households, on rK39 positivity in Marsabit, Kenya (C), and Gedaref, Sudan (D). The ‘margins’ command of Stata was used to compute adjusted log-odds of seropositivity at selected distances from the index households. Analysis includes index household members. A P value of ≤0.05 was considered significant by Mann-Whitney test (A,B), or univariable binary logistic regression (C,D); *P < 0.05, **P < 0.01.

4. Discussion

We provide evidence of widespread and intense VL transmission across our diverse study sites, recovering infected specimens from both sylvatic and peridomestic habitats that reinforce previous findings [8], [18], [20], [22], [46], [47].

We recovered Leishmania-infected Ph. orientalis from a variety of microhabitats within our study sites in Kenya, and Sudan, frequently from the same location months apart. This indicates that Ph. orientalis has adapted to diverse ecological microhabitats driving widespread yet focal hotbeds of VL transmission. Earlier collections from Helat-Belo, Gedaref state, Sudan, conducted during the dry season in March to June 2016–2018, found unfed and blood fed Leishmania-infected Ph. orientalis mainly outside houses and in sylvatic ecotypes [18]. In our study, performed during the rainy season (June to September), Leishmania-infected Ph. orientalis females were collected mainly indoors and had fed soley on humans, suggesting a potential shift in vector behavior, with sand flies likely resting inside the house after feeding on humans. Surprisingly, we collected juvenile males indoors in Sudan, possibly due to the widespread nature of vertisols (black cotton soil) in the village, including inside houses, that serve as a breeding site for this vector [19], and creating an ideal environment for focal and active VL transmission indoors. Further confirmation of this shift in vector behavior and transmission pattern is needed to inform control measures that are currently focused on outdoor applications.

In Ethiopia, Ph. martini were predominantly collected from termite hills, both in sylvatic and peridomestic settings, pointing to a restricted habitat preference compared to Ph. orientalis. Additionally, all three Leishmania-infected sand flies were recovered from termite hills reinforcing findings of a previous study [8] and highlighting the importance of this microhabitat in sustaining transmission by Ph. martini. In-depth studies focused on termite hills are needed to fully elucidate the involvement of this microhabitat in VL transmission.

Despite reported exposure to Leishmania in domestic and wild animal species in EA [48], [49], [50], [51], [52], live parasites have not been isolated from any thus far. Irrespective of whether domestic or wild animals are reservoirs, VL endemicity in scarcely populated areas supports the existence of a zoonotic transmission cycle in EA [18], [23], [53]. Establishing whether animal reservoirs are involved in sustaining VL transmission in EA remains a critical knowledge gap for elimination strategies.

Finding infected sand flies near VL cases from all our study sites affirms the importance of human reservoirs in maintaining transmission, and the need for continued surveillance and rapid treatment of cases. This is further reinforced by finding a significantly higher proportion of positive rK39 antibodies and an increased risk associated with proximity to VL cases in both Marsabit, Kenya, and Gedaref, Sudan. Importantly, the high prevalence of rK39 antibodies in asymptomatic participants provides further evidence of prevalent and efficient transmission and stresses the urgency of determining their involvement as potential reservoirs [15], [54].

A limitation of our study is the use rK39 to assess seroprevalence and the small sample size of VL index cases in Sudan. The sensitivity of rK39 in EA is known to be suboptimal [55], [56], [57]. In our study, this is reinforced by the loss of rK39 positivity in recently diagnosed VL cases emphasizing the need for diagnostics with better specificity and sensitivity in EA. Future studies with a larger sample size are also needed to assess the increased risk of living in proximity to VL cases. Our study adopted a one health framework, investigating the association between humans, animals, vegetation, and the environment to uncover distinct hotbeds driving VL transmission in EA. Moving forward, we need in-depth longitudinal studies focused on these identified hotbeds of infection to better understand the requisites supporting transmission in specific microhabitats.

CRediT authorship contribution statement

Eva Iniguez: Writing – review & editing, Writing – original draft, Visualization, Validation, Supervision, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. Mercy Tuluso: Writing – review & editing, Methodology, Investigation, Data curation. Steve Kiplagat: Writing – review & editing, Methodology, Investigation, Data curation. Araya Gebresilassie: Writing – review & editing, Methodology, Investigation, Data curation. Esayas Aklilu: Writing – review & editing, Methodology, Investigation, Data curation. Olivia Battistoni: Writing – review & editing, Methodology, Investigation. Johnstone Ingonga: Writing – review & editing, Methodology, Investigation, Data curation. John Mark Makwatta: Writing – review & editing, Methodology, Investigation. Mohamed Alamin: Writing – review & editing, Methodology, Investigation. Osman Dakien: Writing – review & editing, Methodology, Investigation, Data curation. Alphine Chebet: Writing – review & editing, Investigation. Esther Kaunda: Writing – review & editing, Methodology, Investigation, Data curation. Patrick Huffcutt: Writing – review & editing, Validation, Methodology, Investigation, Formal analysis, Data curation. Serena Doh: Writing – review & editing, Methodology, Investigation. Pedro Cecilio: Writing – review & editing, Methodology, Investigation. Galgallo Bonaya: Writing – review & editing, Methodology, Investigation, Conceptualization. Claudio Meneses: Writing – review & editing, Methodology, Investigation. Tiago D. Serafim: Writing – review & editing, Methodology, Investigation. Myrthe Pareyn: Writing – review & editing, Investigation. Mohamed Osman: Writing – review & editing, Investigation. Eltahir A.G. Khalil: Writing – review & editing, Methodology, Investigation, Data curation. Omran F. Osman: Writing – review & editing, Methodology, Investigation, Data curation. Brima M. Younis: Writing – review & editing, Supervision, Methodology, Investigation, Data curation. Sithar Dorjee: Writing – review & editing, Software, Methodology, Formal analysis, Data curation. Guofa Zhou: Writing – review & editing, Software, Investigation, Formal analysis, Data curation. Jesus G. Valenzuela: Writing – review & editing, Supervision, Resources, Investigation, Funding acquisition. Dan K. Masiga: Writing – review & editing, Visualization, Supervision, Project administration, Investigation, Funding acquisition, Conceptualization. Ahmed M. Musa: Writing – review & editing, Visualization, Supervision, Project administration, Investigation, Funding acquisition, Formal analysis, Conceptualization. Asrat Hailu: Writing – review & editing, Visualization, Supervision, Project administration, Investigation, Funding acquisition, Formal analysis, Data curation. Abhay Satoskar: Writing – review & editing, Visualization, Supervision, Project administration, Investigation, Funding acquisition, Conceptualization. Shaden Kamhawi: Writing – review & editing, Writing – original draft, Visualization, Supervision, Resources, Project administration, Investigation, Funding acquisition, Formal analysis, Conceptualization. Damaris Matoke-Muhia: Writing – review & editing, Visualization, Supervision, Project administration, Investigation, Funding acquisition, Conceptualization.

Ethics declaration

Written informed consent to take part in the study and to publish the article has been obtained from all participants or their legal representatives. The privacy rights of participants have been observed.

This study included organ or tissue donors. This study includes human biological material and consent was obtained by donors, or their next of kin or legal representatives, for use in this study and for publication of the article. The samples used in this research were not sourced from executed prisoners or prisoners of conscience.

This study was performed in compliance with relevant laws, regulatory frameworks and guidelines where the research took place. This study was approved by the The study was approved by the Scientific Research Unit, KEMRI, and licensed by the National Commission for Science, Technology and Innovation, and by the State Ministry of Health and Social Development at Gedaref state, Sudan ethical committee in Sudan. Human blood was obtained from IRB human protocol 000331-I (human). (Approval No. For Kenya, clinical protocol # KEMRI/SERU/CBRD/249/4634 and for Sudan clinical protocol # UG/EC/3/2023. Human blood was obtained from IRB human protocol 000331-I (human).)

This study was conducted in accordance with the ARRIVE (Animal Research: Reporting of In Vivo Experiments) guidelines. This study was approved by the National Institute of Allergy and Infectious Diseases (NIAID/NIH) (Approval No. Mice blood obtained from LMVR4).

Declaration of generative AI and AI-assisted technologies in the writing process

During the preparation of this work, the authors used GPT-4o (OpenAI, NIAID GenAI Toolkit) in order to improve readability and language of the manuscript. After using this tool/service, the authors reviewed and edited the content as needed and take full responsibility for the content of the publication.

Funding

Funding support for this study was obtained from the NIH under the Tropical Medicine Research Centers (TMRC) grant (U01AI168619). This research was supported by the Intramural Research Program of the National Institutes of Health (NIH). The contributions of the NIH authors are considered Works of the United States Government. The findings and conclusions presented in this paper are those of the authors and do not necessarily reflect the views of the NIH or the U.S. Department of Health and Human Services.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgements

We thank the community, administrative officers, and health care workers from all our study sites. We are also grateful to Marsabit County, Kenya, for supporting us with fieldwork implementation and logistics. We thank Professor EL-Hassan Center for Tropical Diseases staff, community leaders and the population in the Kunaynah village, Gedaref state, Sudan. We thank the Karat Zone Health Bureau, Ethiopia, for supporting this work and the Arbaminch University for hosting the project in Southern Ethiopia. We also acknowledge Rogers Asamba (KEMRI), and David Mbuvi (ICIPE) for their valuable support during fieldwork; Dr. Geyeto Garra (Ethiopia) for support given to this project; Mr. Haile Gebremariam (Ethiopia) for technical assistance.

Footnotes

Appendix A

Supplementary data to this article can be found online at https://doi.org/10.1016/j.onehlt.2026.101575.

Contributor Information

Eva Iniguez, Email: eva.iniguez@nih.gov.

Shaden Kamhawi, Email: skamhawi@niaid.nih.gov.

Damaris Matoke-Muhia, Email: DMatoke@kemri.go.ke.

Appendix A. Supplementary data

Supplementary material

mmc1.pdf (5.7MB, pdf)

Data availability

All data produced in the present work are contained in the manuscript.

References

  • 1.World Health Organization Leishmaniasis. 2024. https://www.who.int/news-room/fact-sheets/detail/leishmaniasis (accessed 17 May 2024).
  • 2.eBioMedicine Leishmania: an urgent need for new treatments. EBioMedicine. 2023;87 doi: 10.1016/j.ebiom.2023.104440. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Baleela R., Llewellyn M.S., Fitzpatrick S., Kuhls K., Schonian G., Miles M.A., et al. Leishmania donovani populations in eastern Sudan: temporal structuring and a link between human and canine transmission. Parasit. Vectors. 2014;7:496. doi: 10.1186/s13071-014-0496-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Zackay A., Cotton J.A., Sanders M., Hailu A., Nasereddin A., Warburg A., et al. Genome wide comparison of Ethiopian Leishmania donovani strains reveals differences potentially related to parasite survival. PLoS Genet. 2018;14 doi: 10.1371/journal.pgen.1007133. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Zackay A., Nasereddin A., Takele Y., Tadesse D., Hailu W., Hurissa Z., et al. Polymorphism in the HASPB repeat region of east African Leishmania donovani strains. PLoS Negl. Trop. Dis. 2013;7 doi: 10.1371/journal.pntd.0002031. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.J. Alvar, M. den Boer, D.A. Dagne, Towards the elimination of visceral leishmaniasis as a public health problem in East Africa: reflections on an enhanced control strategy and a call for action, Lancet Glob. Health 9 (2021) e1763–e9. doi: 10.1016/S2214-109X(21)00392-2. [DOI] [PMC free article] [PubMed]
  • 7.Elnaiem D.E. Ecology and control of the sand fly vectors of Leishmania donovani in East Africa, with special emphasis on Phlebotomus orientalis. J. Vector Ecol. 2011;36(Suppl. 1):S23–S31. doi: 10.1111/j.1948-7134.2011.00109.x. [DOI] [PubMed] [Google Scholar]
  • 8.Gebre-Michael T., Lane R.P. The roles of Phlebotomus martini and P.Celiae (Diptera: Phlebotominae) as vectors of visceral leishmaniasis in the aba Roba focus, southern Ethiopia. Med. Vet. Entomol. 1996;10:53–62. doi: 10.1111/j.1365-2915.1996.tb00082.x. [DOI] [PubMed] [Google Scholar]
  • 9.Gebre-Michael T., Malone J.B., Balkew M., Ali A., Berhe N., Hailu A., et al. Mapping the potential distribution of Phlebotomus martini and P. orientalis (Diptera: Psychodidae), vectors of kala-azar in East Africa by use of geographic information systems. Acta Trop. 2004;90:73–86. doi: 10.1016/j.actatropica.2003.09.021. [DOI] [PubMed] [Google Scholar]
  • 10.Ruiz-Postigo J.A., Madjou S., Maia-Elkhoury A.N., Valadas S., Warusavithana S., Osman M., et al. Global leishmaniasis surveillance: 2021, assessing the impact of the COVID-19 pandemic. Wkly Epidemiol. Rec. 2022;97:575–590. [Google Scholar]
  • 11.World Health Organization . World Health Organization; Geneva: 2020. Ending the Neglect to Attain the Sustainable Development Goals: A Road Map for Neglected Tropical Diseases 2021–2030.https://iris.who.int/bitstream/handle/10665/338565/9789240010352-eng.pdf [Google Scholar]
  • 12.World Health Organization . World Health Organization; Geneva: 2024. Strategic Framework for the Elimination of Visceral Leishmaniasis as a Public Health Problem in Eastern Africa 2023–2030.https://iris.who.int/server/api/core/bitstreams/8b870651-422c-4c04-95ec-1e5e887d2449/content [Google Scholar]
  • 13.Alvar J., Beca-Martínez M.T., Argaw D., Jain S., Aagaard-Hansen J. Social determinants of visceral leishmaniasis elimination in eastern Africa. BMJ Glob. Health. 2023;8 doi: 10.1136/bmjgh-2023-012638. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Wasunna M., Musa A., Hailu A., Khalil E.A., Olobo J., Juma R., et al. The Leishmaniasis East Africa platform (LEAP): strengthening clinical trial capacity in resource-limited countries to deliver new treatments for visceral leishmaniasis. Trans. R. Soc. Trop. Med. Hyg. 2016;110:321–323. doi: 10.1093/trstmh/trw031. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Iniguez E., Masiga D., Bern C., Srikantiah S. Adapting lessons from the Indian subcontinent to accelerate elimination of visceral leishmaniasis as a public health problem in East Africa. PLoS Negl. Trop. Dis. 2026;20 doi: 10.1371/journal.pntd.0014088. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Gebresilassie A., Kirstein O.D., Yared S., Aklilu E., Moncaz A., Tekie H., et al. Species composition of phlebotomine sand flies and bionomics of Phlebotomus orientalis (Diptera: Psychodidae) in an endemic focus of visceral leishmaniasis in Tahtay Adiyabo district, Northern Ethiopia. Parasit. Vectors. 2015;8:248. doi: 10.1186/s13071-015-0849-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Hassaballa I.B., Torto B., Sole C.L., Tchouassi D.P. Exploring the influence of different habitats and their volatile chemistry in modulating sand fly population structure in a leishmaniasis endemic foci, Kenya. PLoS Negl. Trop. Dis. 2021;15 doi: 10.1371/journal.pntd.0009062. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Khogali A., Elnaiem D.A., Diaz-Reganon R., Jibreel T., Nour B.Y.M., Abdelrahman S.H., et al. Infection of Leishmania donovani in Phlebotomus orientalis sand flies at different microhabitats of a kala-azar endemic village in eastern Sudan. Trop. Med. Infect. Dis. 2024;9 doi: 10.3390/tropicalmed9020040. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Moncaz A., Kirstein O., Gebresellassie A., Lemma W., Yared S., Gebre-Michael T., et al. Characterization of breeding sites of Phlebotomus orientalis - the vector of visceral leishmaniasis in northwestern Ethiopia. Acta Trop. 2014;139:5–14. doi: 10.1016/j.actatropica.2014.06.013. [DOI] [PubMed] [Google Scholar]
  • 20.Elnaiem D.E., Osman O.F. Evidence for active transmission of visceral leishmaniasis within a village in eastern Sudan. Acta Trop. 1998;71:305–309. doi: 10.1016/s0001-706x(98)00067-9. [DOI] [PubMed] [Google Scholar]
  • 21.Hailu A., Balkew M., Berhe N., Meredith S.E., Gemetchu T. Is Phlebotomus (Larroussius) orientalis a vector of visceral leishmaniasis in south-West Ethiopia? Acta Trop. 1995;60:15–20. doi: 10.1016/0001-706x(95)00093-t. [DOI] [PubMed] [Google Scholar]
  • 22.Kirstein O.D., Skrip L., Abassi I., Iungman T., Horwitz B.Z., Gebresilassie A., et al. A fine scale eco-epidemiological study on endemic visceral leishmaniasis in north ethiopian villages. Acta Trop. 2018;183:64–77. doi: 10.1016/j.actatropica.2018.04.005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Zijlstra E.E., el-Hassan A.M. Leishmaniasis in Sudan. Visceral leishmaniasis. Trans. R. Soc. Trop. Med. Hyg. 2001;95(Suppl. 1):S27–S58. doi: 10.1016/s0035-9203(01)90218-4. [DOI] [PubMed] [Google Scholar]
  • 24.Votypka J., Pavlasova M., Volfova V., Volf P. Rotation of male genitalia in various species of phlebotomine sandfly. Med. Vet. Entomol. 2015;29:355–360. doi: 10.1111/mve.12132. [DOI] [PubMed] [Google Scholar]
  • 25.Mondal D., Bern C., Ghosh D., Rashid M., Molina R., Chowdhury R., et al. Quantifying the infectiousness of post-kala-azar dermal Leishmaniasis toward sand flies. Clin. Infect. Dis. 2019;69:251–258. doi: 10.1093/cid/ciy891. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Singh O.P., Tiwary P., Kushwaha A.K., Singh S.K., Singh D.K., Lawyer P., et al. Xenodiagnosis to evaluate the infectiousness of humans to sandflies in an area endemic for visceral leishmaniasis in Bihar, India: a transmission-dynamics study. Lancet Microbe. 2021;2:e23–e31. doi: 10.1016/S2666-5247(20)30166-X. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Singh O.P., Chaubey R., Kushwaha A.K., Fay M.P., Sacks D., Sundar S. Visceral Leishmaniasis-human immunodeficiency virus-Coinfected patients are highly infectious to sandflies in an endemic area in India. J. Infect. Dis. 2024;229:1909–1912. doi: 10.1093/infdis/jiae156. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Al-Salem W., Herricks J.R., Hotez P.J. A review of visceral leishmaniasis during the conflict in South Sudan and the consequences for east African countries. Parasit. Vectors. 2016;9:460. doi: 10.1186/s13071-016-1743-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Berry I., Berrang-Ford L. Leishmaniasis, conflict, and political terror: a spatio-temporal analysis. Soc. Sci. Med. 2016;167:140–149. doi: 10.1016/j.socscimed.2016.04.038. [DOI] [PubMed] [Google Scholar]
  • 30.Boodman C., van Griensven J., Gupta N., Diro E., Ritmeijer K. Anticipating visceral leishmaniasis epidemics due to the conflict in northern Ethiopia. PLoS Negl. Trop. Dis. 2023;17 doi: 10.1371/journal.pntd.0011188. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.de Souza D.K., Picado A., Bessell P.R., Liban A., Wachira D., Mwiti D., et al. Strengthening visceral leishmaniasis diagnosis capacity to improve access to care in Kenya: the example of Marsabit County. Front. Trop. Dis. 2021;2 doi: 10.3389/fitd.2021.809757. [DOI] [Google Scholar]
  • 32.Mewara A., Gudisa R., Padhi B.K., Kumar P., Sah R., Rodriguez-Morales A.J. Visceral leishmaniasis outbreak in Kenya-a setback to the elimination efforts. New Microbes New Infect. 2022;49-50 doi: 10.1016/j.nmni.2022.101060. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Kenya National Bureau of Statistics . Kenya National Bureau of Statistics; Nairobi, Kenya: 2019. Kenya population and housing census volume II: distribution of population by administrative units. 2019. [Google Scholar]
  • 34.Kenya Meteorological Department . Kenya Meteorological Department; Nairobi, Kenya: 2025. State of the Climate Kenya 2024.https://meteo.go.ke [Google Scholar]
  • 35.Ibrahim B., Elmadani M., Hemeda S., Gadallh A.A.M., Elawad Ahmed A.E. Prevalence of visceral leishmaniasis among wildlife rangers in Dinder National Park, Sudan. J. Infect. Dev. Ctries. 2026;20:755–763. doi: 10.3855/jidc.20568. [DOI] [PubMed] [Google Scholar]
  • 36.Abonnenc E., Minter D. 1965. Tables D'identification Bilingues Des phlébotomes de la région éthiopienne = Bilingual Keys for the Identification of the Sandflies of the Ethiopian Region, Paris. [Google Scholar]
  • 37.Lewis D.J. The phlebotomine sandflies (Diptera: Psychodidae) of the oriental region. Bull. Br. Mus. Nat. Hist. Entomol. 1978;37:217–343. [Google Scholar]
  • 38.Dey R., Joshi A.B., Oliveira F., Pereira L., Guimaraes-Costa A.B., Serafim T.D., et al. Gut microbes egested during bites of infected sand flies augment severity of Leishmaniasis via inflammasome-derived IL-1beta. Cell Host Microbe. 2018;23:134–43 e6. doi: 10.1016/j.chom.2017.12.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Kent R.J., Norris D.E. Identification of mammalian blood meals in mosquitoes by a multiplexed polymerase chain reaction targeting cytochrome B. Am. J. Trop. Med. Hyg. 2005;73:336–342. [PMC free article] [PubMed] [Google Scholar]
  • 40.Irwin D.M., Kocher T.D., Wilson A.C. Evolution of the cytochrome b gene of mammals. J. Mol. Evol. 1991;32:128–144. doi: 10.1007/BF02515385. [DOI] [PubMed] [Google Scholar]
  • 41.Collini M., Albonico F., Hauffe H.C., Mortarino M. Identifying the last bloodmeal of questing sheep tick nymphs (Ixodes ricinus L.) using high resolution melting analysis. Vet. Parasitol. 2015;210:194–205. doi: 10.1016/j.vetpar.2015.04.007. [DOI] [PubMed] [Google Scholar]
  • 42.B. Parodi, O. Aresu, D. Bini, R. Lorenzini, F. Schena, P. Visconti, et al., Species identification and confirmation of human and animal cell lines: a PCR-based method, Biotechniques 32 (2002) 432–4, 6, 8–40. doi: 10.2144/02322rr05. [DOI] [PubMed]
  • 43.Abbasi I., Cunio R., Warburg A. Identification of blood meals imbibed by phlebotomine sand flies using cytochrome b PCR and reverse line blotting. Vector Borne Zoonotic Dis. 2009;9:79–86. doi: 10.1089/vbz.2008.0064. [DOI] [PubMed] [Google Scholar]
  • 44.Iniguez E., Priyamvada K., Dubey P., Bindroo J., Shahnawaz M., Alam A., et al. High seropositivity to rK39 and capture of Leishmania-infected sand flies in Bihar and Jharkhand emphasize the need for post-elimination surveillance of visceral Leishmaniasis in India. J. Infect. Dis. 2026;233:e802–e811. doi: 10.1093/infdis/jiaf543. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45.Kenya Ministry of Health . Kenya Ministry of Health; Nairobi, Kenya: 2017. Prevention, Diagnosis and Treatment of Visceral Leishmaniasis (Kala-Azar) in Kenya: National Guidelines.https://www.afrikadia.org/wp-content/uploads/2018/08/VL_Guidelines_Kenya_2017.pdf [Google Scholar]
  • 46.Hassan M.M., Elraba'a F.M., Ward R.D., Maingon R.D., Elnaiem D.A. Detection of high rates of in-village transmission of Leishmania donovani in eastern Sudan. Acta Trop. 2004;92:77–82. doi: 10.1016/j.actatropica.2004.06.001. [DOI] [PubMed] [Google Scholar]
  • 47.Perkins P.V., Githure J.I., Mebrahtu Y., Kiilu G., Anjili C., Ngumbi P.S., et al. Isolation of Leishmania donovani from Phlebotomus martini in Baringo district, Kenya. Trans. R. Soc. Trop. Med. Hyg. 1988;82:695–700. doi: 10.1016/0035-9203(88)90204-0. [DOI] [PubMed] [Google Scholar]
  • 48.Dereure J., El-Safi S.H., Bucheton B., Boni M., Kheir M.M., Davoust B., et al. Visceral leishmaniasis in eastern Sudan: parasite identification in humans and dogs; host-parasite relationships. Microbes Infect. 2003;5:1103–1108. doi: 10.1016/j.micinf.2003.07.003. [DOI] [PubMed] [Google Scholar]
  • 49.Elnaiem D.A., Hassan M.M., Maingon R., Nureldin G.H., Mekawi A.M., Miles M., et al. The Egyptian mongoose, Herpestes ichneumon, is a possible reservoir host of visceral leishmaniasis in eastern Sudan. Parasitology. 2001;122:531–536. doi: 10.1017/s0031182001007594. [DOI] [PubMed] [Google Scholar]
  • 50.Gebresilassie A., Yared S., Aklilu E., Kirstein O.D., Moncaz A., Tekie H., et al. Host choice of Phlebotomus orientalis (Diptera: Psychodidae) in animal baited experiments: a field study in Tahtay Adiyabo district, northern Ethiopia. Parasit. Vectors. 2015;8:190. doi: 10.1186/s13071-015-0807-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 51.Hassan M.M., Osman O.F., El-Raba'a F.M., Schallig H.D., Elnaiem D.E. Role of the domestic dog as a reservoir host of Leishmania donovani in eastern Sudan. Parasit. Vectors. 2009;2 doi: 10.1186/1756-3305-2-26. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Kassahun A., Sadlova J., Benda P., Kostalova T., Warburg A., Hailu A., et al. Natural infection of bats with Leishmania in Ethiopia. Acta Trop. 2015;150:166–170. doi: 10.1016/j.actatropica.2015.07.024. [DOI] [PubMed] [Google Scholar]
  • 53.Elnaiem D.A., Hassan H.K., Ward R.D. Phlebotomine sandflies in a focus of visceral leishmaniasis in a border area of eastern Sudan. Ann. Trop. Med. Parasitol. 1997;91:307–318. doi: 10.1080/00034989761157. [DOI] [PubMed] [Google Scholar]
  • 54.Singh O.P., Hasker E., Sacks D., Boelaert M., Sundar S. Asymptomatic Leishmania infection: a new challenge for Leishmania control. Clin. Infect. Dis. 2014;58:1424–1429. doi: 10.1093/cid/ciu102. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 55.Hagos D.G., Schallig H., Kiros Y.K., Abdulkadir M., Wolday D. Performance of rapid rk39 tests for the diagnosis of visceral leishmaniasis in Ethiopia: a systematic review and meta-analysis. BMC Infect. Dis. 2021;21:1166. doi: 10.1186/s12879-021-06826-w. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 56.Iniguez E., Kamhawi S. Discovery of new antigens for serodiagnosis of visceral leishmaniasis in humans and dogs adds valuable tools to our arsenal. mBio. 2024;15:e0107924. doi: 10.1128/mbio.01079-24. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 57.Kassa M., Abdellati S., Cnops L., Bremer Hinckel B.C., Yeshanew A., Hailemichael W., et al. Diagnostic accuracy of direct agglutination test, rK39 ELISA and six rapid diagnostic tests among visceral leishmaniasis patients with and without HIV coinfection in Ethiopia. PLoS Negl. Trop. Dis. 2020;14 doi: 10.1371/journal.pntd.0008963. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary material

mmc1.pdf (5.7MB, pdf)

Data Availability Statement

All data produced in the present work are contained in the manuscript.

Articles from One Health are provided here courtesy of Elsevier

RESOURCES