• Loading metrics

Leishmania major and Trypanosoma lewisi infection in invasive and native rodents in Senegal

  • Cécile Cassan ,

    Roles Investigation, Methodology, Validation, Writing – original draft

    Affiliation MIVEGEC, IRD, CNRS, Univ. Montpellier, Montpellier, France

  • Christophe A. Diagne,

    Roles Formal analysis, Methodology, Validation, Writing – original draft

    Affiliations CBGP, IRD, CIRAD, INRA, Montpellier SupAgro, Univ Montpellier, Montpellier, France, CBGP, IRD, CIRAD, INRA, Montpellier SupAgro, Univ Montpellier, Campus ISRA/IRD de Bel Air, Dakar, Sénégal, Département de Biologie Animale, Faculté des Sciences et Techniques, Université Cheikh Anta Diop (UCAD), Dakar, Sénégal

  • Caroline Tatard,

    Roles Investigation

    Affiliation CBGP, INRA, CIRAD, IRD, Montpellier SupAgro, Univ Montpellier, Montpellier, France

  • Philippe Gauthier,

    Roles Investigation

    Affiliation CBGP, IRD, CIRAD, INRA, Montpellier SupAgro, Univ Montpellier, Montpellier, France

  • Ambroise Dalecky,

    Roles Investigation

    Affiliation IRD, LPED (UMR AMU/IRD), Marseille, France

  • Khalilou Bâ,

    Roles Investigation

    Affiliation CBGP, IRD, CIRAD, INRA, Montpellier SupAgro, Univ Montpellier, Campus ISRA/IRD de Bel Air, Dakar, Sénégal

  • Mamadou Kane,

    Roles Investigation

    Affiliation CBGP, IRD, CIRAD, INRA, Montpellier SupAgro, Univ Montpellier, Campus ISRA/IRD de Bel Air, Dakar, Sénégal

  • Youssoupha Niang,

    Roles Investigation

    Affiliation CBGP, IRD, CIRAD, INRA, Montpellier SupAgro, Univ Montpellier, Campus ISRA/IRD de Bel Air, Dakar, Sénégal

  • Mamoudou Diallo,

    Roles Investigation

    Affiliation CBGP, IRD, CIRAD, INRA, Montpellier SupAgro, Univ Montpellier, Campus ISRA/IRD de Bel Air, Dakar, Sénégal

  • Aliou Sow,

    Roles Investigation

    Affiliation CBGP, IRD, CIRAD, INRA, Montpellier SupAgro, Univ Montpellier, Campus ISRA/IRD de Bel Air, Dakar, Sénégal

  • Carine Brouat,

    Roles Conceptualization, Funding acquisition, Investigation, Methodology, Project administration, Supervision, Validation, Writing – review & editing

    Affiliation CBGP, IRD, CIRAD, INRA, Montpellier SupAgro, Univ Montpellier, Montpellier, France

  • Anne-Laure Bañuls

    Roles Conceptualization, Formal analysis, Funding acquisition, Methodology, Project administration, Supervision, Validation, Writing – review & editing

    Affiliation MIVEGEC, IRD, CNRS, Univ. Montpellier, Montpellier, France

Leishmania major and Trypanosoma lewisi infection in invasive and native rodents in Senegal

  • Cécile Cassan, 
  • Christophe A. Diagne, 
  • Caroline Tatard, 
  • Philippe Gauthier, 
  • Ambroise Dalecky, 
  • Khalilou Bâ, 
  • Mamadou Kane, 
  • Youssoupha Niang, 
  • Mamoudou Diallo, 
  • Aliou Sow


Bioinvasion is a major public health issue because it can lead to the introduction of pathogens in new areas and favours the emergence of zoonotic diseases. Rodents are prominent invasive species, and act as reservoirs in many zoonotic infectious diseases. The aim of this study was to determine the link between the distribution and spread of two parasite taxa (Leishmania spp. and Trypanosoma lewisi) and the progressive invasion of Senegal by two commensal rodent species (the house mouse Mus musculus domesticus and the black rat Rattus rattus). M. m. domesticus and R. rattus have invaded the northern part and the central/southern part of the country, respectively. Native and invasive rodents were caught in villages and cities along the invasion gradients of both invaders, from coastal localities towards the interior of the land. Molecular diagnosis of the two trypanosomatid infections was performed using spleen specimens. In the north, neither M. m. domesticus nor the native species were carriers of these parasites. Conversely, in the south, 17.5% of R. rattus were infected by L. major and 27.8% by T. lewisi, while very few commensal native rodents were carriers. Prevalence pattern along invasion gradients, together with the knowledge on the geographical distribution of the parasites, suggested that the presence of the two parasites in R. rattus in Senegal is of different origins. Indeed, the invader R. rattus could have been locally infected by the native parasite L. major. Conversely, it could have introduced the exotic parasite T. lewisi in Senegal, the latter appearing to be poorly transmitted to native rodents. Altogether, these data show that R. rattus is a carrier of both parasites and could be responsible for the emergence of new foci of cutaneous leishmaniasis, or for the transmission of atypical human trypanosomiasis in Senegal.

Author summary

Biological invasions (the introduction and adaptation of living organisms to a new environment) are increasingly frequent due to worldwide intensification of human-associated exchanges. They can lead to the introduction of pathogens in new areas and favour the emergence of diseases. Rodents are prominent invasive species, and act as reservoirs in many infectious diseases. The aim of our study was to determine the link between the distribution and spread of two parasites, Leishmania spp. and Trypanosoma lewisi, and the progressive invasion of Senegal by two commensal rodent species, the house mouse Mus musculus domesticus, and the black rat Rattus rattus. We identified R. rattus as a potential reservoir for Leishmania major and T. lewisi in the southern part of Senegal. The presence of these two pathogens in R. rattus may be of different origins. The invader R. rattus could have been locally contaminated with L. major. Conversely, T. lewisi infection could have been introduced in Senegal by R. rattus, and seems to be poorly transmitted to native rodents. Altogether, these data show that R. rattus is a carrier of both parasites, and could be responsible for the emergence of new foci of cutaneous leishmaniasis or for the transmission of atypical human trypanosomiasis in Senegal.


Biological invasions are increasingly frequent due to the worldwide intensification of human-associated exchanges. They can have significant consequences on the biodiversity in many ecosystems [1]. For instance, they might lead to novel parasite-host combinations and have dramatic effects on the dynamics of diseases that affect wildlife, livestock and/or humans [2]. Disease emergence events associated with exotic pathogens imported by animal invaders have already been reported [3]. The dynamics of endemic diseases could also be affected by invasive species that act as novel hosts, or could negatively affect native host species [4].

Among invasive species, rodents indisputably represent the vertebrate group that has most often accompanied the global dispersion of humankind. Rodents act as reservoir for numerous zoonotic agents, such as helminths, bacteria or protozoa. The introduction of exotic rodents has been associated with the appearance of several new foci of infectious diseases in wildlife (e.g., [5,6,7]) and human populations (e.g., [8]). However, up to now, only few studies have evaluated the effects of both native and invasive host communities on infectious disease risk in invaded ecosystems. For instance, Billeter et al. [9] showed that the risk of transmission of bartonellosis to human was higher from native rodents than from invasive black rat in Uganda. In this study, we focused on the spatial distribution of two zoonotic protozoan parasites, Leishmania spp. and Trypanosoma lewisi, in native and invasive populations of commensal rodents in Senegal.

Leishmaniasis is a neglected disease that affects 0.9 to 1.6 million people worldwide and responsible for 20,000 to 40,000 deaths each year [10]. Moreover, 350 million people are considered at risk of contracting this infection [11]. Among the 30 Leishmania species described in mammals, 20 are pathogenic for humans [12,13]. These species are generally host-specific and restricted to particular geographical areas. The parasite is transmitted through the bite of infected female phlebotomine sand flies. In humans, they can cause different clinical forms: asymptomatic infection, visceral leishmaniasis (VL), mucocutaneous leishmaniasis (MCL) or cutaneous leishmaniasis (CL) [13]. VL is mainly caused by Leishmania infantum and Leishmania donovani. Human VL has never been detected in Senegal [10,14,15], but high human serological prevalence of Leishmania has been found in a focus near Thiès [16], where canine leishmaniasis caused by L. infantum has been recorded since last century and recently epidemiologically described [14,16,17,18]. Atypical MCL due to Leishmania major (the main causative agent of MCL is Leishmania braziliensis, whose distribution is restricted to South America [13]) can occur in Senegal but is very rare (Dr. Babacar Faye, personal communication, and [19]). Rural CL is caused by Leishmania major, and is endemic in West and North Africa, and in the Middle East. In Senegal the infection has been observed since the beginning of the 20th century [20,21,22,23,24] and is still reported [25, Dr Babacar Faye, personal communication]. Nevertheless, public health records about the prevalence and distribution of the disease remain limited.The epidemics appear as foci that then disappear, possibly in function of seasons, and dynamics of vectors and reservoir populations [21,24]. Gerbils are traditionally considered to be the L. major reservoirs in the Old World [11]. However, the parasite was identified in various rodent species (e.g., Mastomys erythroleucus, Arvicanthis niloticus and Gerbilliscus gambianus) near Thiès in Senegal [21,26,27]. These asymptomatic rodents (showing no cutaneous lesion) were found infected, by culture or molecular methods. In Senegal, L. major is transmitted by the vector Phlebotomus duboscqi [28]. P. duboscqi is found mainly in rodent burrows, termite mounds and tree holes, but also indoor; it feeds on many vertebrates, birds, reptiles and mammals (rabbits, rodents) and is also very anthropophilic. [18,28,29].

The genus Trypanosoma includes pathogens, such as Trypanosoma gambiense, T. cruzi and T. brucei brucei, responsible for sleeping sickness, Chagas’ disease and African animal trypanosomiasis, or nagana in livestock respectively, as well as other species considered to be non-pathogenic for humans [30]. However, atypical human trypanosomiases that are caused by species normally restricted to animals, for instance T. lewisi, have been recently described [31]. Humans are rare accidental hosts for T. lewisi, and only a few cases have been reported worldwide [30]. The infection is generally associated with mild symptoms, such as fever for a few days, but can require medical intervention in young children, and sometimes can lead to death [30]. T. lewisi has been identified for the first time in France in the 19th century, then in Poland in 1880 and 1901 [32]. It is now found in nearly all continents, including Africa where it was detected in rodents in Niger [33] and Nigeria [34,35], and in one infant in Gambia [36]. Nothing more is known on its presence in West Africa, and no human infection has been reported in Senegal up to now [30], but this atypical trypanosomiasis could be under-diagnosed, possibly due to asymptomatic carriage or to non specific symptoms. T. lewisi is described as quite host-specific, and infects rodents, mainly rats [37]. In rodents, its pathogenic potential is low. This parasite proliferates in blood and is rarely found in organs. T. lewisi is orally transmitted to rodents by ingestion of hematophagous arthropods or their faeces [38].

In Senegal, villages and towns are invaded by two major invasive rodent species (Global Invasive Species Database [39]): the house mouse Mus musculus domesticus and the black rat Rattus rattus. Historical records and molecular analyses have shown that these rodents were first brought to sea ports in Senegal by Europeans during the colonial period [40,41,42]. Over the last century, both taxa have spread to inland villages and towns due to the improvement of transport infrastructures, and the native commensal rodent species have been progressively evicted from the invaded habitats [41].

The goal of this study was to evaluate Leishmania and T. lewisi infection rates in commensal native and invasive rodent communities of Senegal in localities along invasion gradients, in order (i) to explore the link between rodent invasion and the spread of these two trypanosomatids, and (ii) to assess the health risk for human populations.

Materials and methods

Ethical statement

Trapping campaigns within localities and private land were conducted with the authorization of the appropriate institutional and household authorities. They were carried out under the framework agreement established between the Institut de Recherche pour le Développement and the Republic of Senegal, as well as with the Senegalese Head Office of Waters and Forests. None of the rodent species investigated here has protected status (see list of the International Union for Conservation of Nature). Handling procedures were performed under our lab agreement for experiments on wild animals (no. 34-169-1), and followed the official guidelines from the American Society of Mammalogists [43]. Euthanasia was performed as recommended by the Federation of European Laboratory Animal Science Associations (FELASA) for small rodents [44,45]. Sample transfers have been approved by the regional Head of Veterinary Service (Hérault, France).

Distribution of the rodents and sampling

The house mouse is now present in most of northern and central Senegal, whereas the black rat is distributed throughout the southern part of the country (Fig 1). Rodents were live-trapped in localities (villages and towns) along each of these two invasion gradients. We used data from historical records and longitudinal sampling surveys of rodent communities in Senegal carried out since the 1980s [41,46,47], in order to classify sampling localities into three categories related to invasion status: (i) in long-established invasion localities (LI), the house mouse or the black rat settled in large and permanent populations were present since more than a century, and have excluded native rodents; (ii) in recently invaded localities corresponding to invasion front (IF), exotic rodents arrived only recently (10–30 years ago) and currently coexist with native rodents; (iii) in non-invaded localities (NI), the house mouse and the black rat have never been detected, and only native rodents are known to occur. Three to six localities were systematically sampled per invasion category along each invasion gradient (Fig 1). In addition, rodents were trapped also in Mereto, a village in the Terres Neuves region (star in Fig 1). The lack of data on rodent communities in this village (it was not sampled for rodents before this study) prevented its classification into a specific invasion category: the village was created before 1972 [48], so exotic rodents may have arrived there for more than 40 years, or later.

Fig 1. Rodent sampling localities along the house mouse (Mus musculus domesticus) (circles) and black rat (Rattus rattus) (squares) invasion gradients in Senegal (West Africa).

Colour code: black for localities of long-established invasion (LI: rodent communities dominated by invasive species); grey for recently invaded localities or the invasion front (IF: sympatric areas with invasive rodents and native Mastomys erythroleucus, or Mastomys natalensis in KED); white for non-invaded localities (NI: only native rodents, Ma. erythroleucus in the north, Ma. natalensis in the south). Presence (blue dots for Leishmania major; violet dots for Trypanosoma lewisi + L. major) and absence (white dots) of parasites in the rodent species are also indicated. AEL = Aere Lao, BAN = Badi Nieriko, BOU = Boutougoufara, BRA = Bransan, DAG = Dagathie, DEN = Dendoudi, DIK = Diakène Wolof, DIT = Diattacounda, DIW = Diomandiou Walo, DOD = Dodel, DOL = Doumga Lao, GAL = Galoya, KED = Kedougou, LAM = Lambango, LOU = Lougue, MAK = Mako, MAR = Marsassoum, MBA = Mbakhana, NDB = Ndombo, SAM = Sare Maoundé, SEG = Segou, SOU = Soutouta, THL = Thilene, THW = Thiewlé, TOB = Tobor. MRT (= Mereto; hatched star) is invaded by both M. m. domesticus and R. rattus. The map was created using the QGIS software v. 2.18.7 (

Sampling was performed between March and April 2013 in the north, and between November 2013 and February 2014 in the south (including Mereto). Details on rodent trapping and identification, autopsy procedures and age determination are provided elsewhere [41,46,49]. Young animals were eliminated from the study. Each rodent was euthanized by cervical dislocation and dissected.

Sample preparation

For this study, the spleen and one ear from adult animals were collected. Tissues were stored at 4°C in 95% ethanol prior to use. DNA extraction was performed with the DNeasy Blood and Tissue Kit according to the manufacturer’s protocol (Qiagen, Courtaboeuf, France).

To verify the DNA quality and validate negative results, we performed a real time PCR targeting the rodent β-actin gene in all samples collected in southern Senegal with the primers bAqF (5’-TCCGTAAAGACCTCTATGCCAA-3’) and bAqR (5’-CAGAGTACTTGCGCTCAG-3’) [50] on a 7300 Real-Time PCR instrument (Applied Biosystems, Foster City, USA). Each 8μl reaction mix included 1.6μl of 5X Evagreen (Euromedex, Souffelweyersheim, France), 0.375μM of each primer, and 2μl of 100 times diluted DNA (about 5-10ng). Cycling conditions were: 15min initial denaturation followed by 45 amplification cycles (95°C for 10s, 60°C for 15s, 72°C for 30s) and a melting curve (95°C for 15s, 60°C for 30s, 5 acquisitions per sec. up to 95°C at 0.11°C/sec). Fluorescence detection was performed at the end of the amplification step. The expected size of the amplicon was 274b. Actin could not be amplified in 20 samples that were thus re-extracted to improve quality.

Molecular diagnosis of Leishmania

Leishmania detection was performed in spleen and a few ear samples. For Leishmania diagnosis, a highly sensitive nested PCR method to amplify the minicircle kinetoplastic DNA (kDNA) variable region was chosen, as described by Noyes et al. [51]. For the first PCR step, the primers CSB1XR (5’-ATT TTT CSG WTT YGC AGA ACG-3’) and CSB2XF (5’-SRT RCA GAA AYC CCG TTC A-3’) were used and the following conditions: 2min initial denaturation at 94°C, followed by 45 amplification cycles (94°C for 30s, 54°C for 1min, 72°C for 1min) and a final extension step of 72°C for 10min. Each 30μl reaction mix included 0.333μM of each primer, 42μM of each dNTP, 3μl of 10X buffer, 1U of Taq DNA polymerase (Roche Diagnostics, Meylan, France), and 3μl of template DNA. For the second PCR step, 3μl of the first PCR product were used with the same programme (but for the annealing temperature that was increased to 56°C) and the same reaction mix except the primers that were replaced by the following: LIR (5’-TCG CAG AAC GCC CCT-3’) and 13Z (5’-ACT GGG GGT TGG TGT AAA ATA G-3’). For this second step, the expected size of the amplicon was 500-800b depending on the Leishmania species. Leishmania complex identification was based on comparison of the size of the second PCR product on a 1.5% agarose gel in 0.5X TAE buffer with the reference strain profiles [51]. To confirm the diagnosis, amplicons from positive samples were cloned in the pGEM-T vector (Promega, Charbonnières-les-Bains, France) and sequenced with the LIR and 13Z primers. BLAST was used to compare the obtained sequences to those included in the NCBI and TriTrypDB databases. BLAST results were taken into account when they came from reference strains or from sequences published in peer-reviewed articles, and were well characterized in terms of species, origin and hosts, in order to avoid misidentifications. Our sequence data were also compared with sequences obtained with the same method from two L. major reference strains (MHOM/SU/73/5ASKH and MHOM/IL/1980/Friedlin) and from a L. major strain isolated from a Senegalese patient (LC-DKR). This last strain was isolated in 2008 at the Dermatology unit of the hospital A. Le Dantec in Dakar by Pr Babacar Faye (laboratory of Parasitology and Mycology, University Cheikh Anta Diop, Dakar) from a patient with typical L. major lesions; L. major species was confirmed with the nested PCR described above). The experimental clones were named using the field sample number to which a suffix was added. Sequences were submitted to GenBank (S1 Table, S1 Text).

Molecular diagnosis of Trypanosoma lewisi

T. lewisi detection was performed in spleen. Diagnosis in mouse samples was performed using a FRET-based real time PCR method to detect 18S rRNA with the primers TRYP A1 (5'-AGGAATGAAGGAGGGTAGTTCG-3’) and TRYP A2 (5'-CACACTTTGGTTCTTGATTGAGG-3') and the probes TRYP A3 (5’-LC640AGAATTTCACCTCTGACGCCCCAGTPhos-3’) and TRYP A4 (5’-GCTGTAGTTCGTCTTGGTGCGGTCTFlc-3’) [33], on a LightCycler LC 480 instrument (Roche Diagnostics, Meylan, France). Each 12μl reaction mix included 6μl of 2X Maxima Probes master mix (Thermo Fisher Scientific, Waltham, Massachusetts, USA), 0.5μM of each primer, 0.25μM of each probe, and 5μl of template DNA. After enzyme activation at 50°C for 1min and an initial denaturation step of 95°C for 10min, 50 amplification cycles were carried out at 95°C for 10s, 56°C for 10s, 72°C for 15s, followed by a melting curve (95°C for 15s, 56°C for 30s, 5 acquisitions per sec. up to 95°C at 0.11°C/sec). Fluorescence acquisition was performed with a Red 640 filter at the end of the annealing step. The expected size of the amplicon was 131bp.

The FRET-based real time PCR was chosen for its robustness and sensitivity (<9.5fg, S1 Fig). However, this PCR is not specific for T. lewisi, but detects also Leishmania (S1 Fig). This latter parasite was not detected in samples from the first campaign realized in the north of Senegal, but was detected in some of those from the second campaign in the central/south of Senegal (see Results). Therefore, we developed a more specific approach based on real time PCR amplification of T. lewisi mini-exon instead of the FRET-based real time PCR for this second campaign. The mini-exon amplification was realized with the primers ME-F (5'-GCTGACACCGGTTGGTTCTG -3’) and ME-R (5'-GACAGCAGCGTGAGCAATAA -3'). The 8μl reaction mix included 4μl 2X SYBR Green (Roche Diagnostics, Meylan, France), 0.6μM of each primer, and 2μl of template DNA and the following cycling conditions: initial denaturation step of 95°C for 5min, followed by 50 amplification cycles (95°C for 15s, 59°C for 20s, 72°C for 15s) and a melting curve (95°C for 5s, 59°C for 1min, 5 acquisitions per sec. up to 95°C at 0.11°C/sec) on a LightCycler LC 480 instrument (Roche Diagnostics, Meylan, France). Fluorescence acquisition was performed at the end of the amplification step. This PCR was specific to T. lewisi with a sensitivity of 1.9fg. The expected size of the amplicon was 140bp. Results were confirmed by amplicon sequencing (S2 Text) and comparison with sequence data from the NCBI database; sequences obtained showed homologies with the mini-exon sequence of the strain Molteno B3 (Genbank AJ250740.1).

Statistical analysis

Statistical analyses were restricted to R. rattus as positive samples for both Leishmania and T. lewisi were mainly identified in this host species (see Results). Generalized linear mixed models (GLMMs) assuming a binomial distribution were used to test whether prevalence levels (infected/non-infected) differed among invasion categories. The sampling locality was considered as a random effect. P-values were obtained by stepwise model simplification in likelihood-ratio tests and were considered significant when <0.05. The models were validated by checking the normality, independence and variance homogeneity of the residuals. All model analyses were performed with the R software v 3.2.2 [52] using the package lme4 v1.1–14 [53].


During the two sampling campaigns, the following commensal native rodents were trapped: Ma. erythroleucus in northern Senegal, and Ma. erythroleucus, Mastomys natalensis, A. niloticus and Praomys daltoni in southern or central Senegal (see Dalecky et al. 2015 [41] for a detailed description of commensal rodent communities in Senegal). Molecular diagnosis was performed on tissue specimens from 161 M. m. domesticus and 167 Ma. erythroleucus samples in the north (Table 1), and from 241 invasive rodent (212 R. rattus, 29 M. m. domesticus) and 219 native rodent (96 Ma. natalensis, 67 Ma. erythroleucus, 12 A. niloticus and 15 P. daltoni) samples in the south (Table 1) (Fig 1). Mereto was the only locality where native rodents coexisted with both mice and rats.

Table 1. Summary of rodents trapped in localities of Mus musculus domesticus or Rattus rattus long-established invasion (LI), invasion front (IF) and no invasion (NI).

Leishmania diagnosis

None of the trapped rodents presented any clinical sign of leishmaniasis. Molecular diagnosis was first performed on spleen samples. All rodents sampled in the north of Senegal (Ma. erythroleucus and M. m. domesticus) were negative for Leishmania (Fig 1).

Conversely, in the south, 37 R. rattus, one M. m. domesticus (from Mereto) and two native rodent samples (Ma. erythroleucus in Soutouta; A. niloticus in Mereto) were positive for Leishmania (Table 2 and Fig 1). The positive samples represented 12.5% and 11.2% of all specimens in the LI and IF localities, respectively, excluding Mereto. No Leishmania infection was detected in samples from NI localities (Fig 1). In R. rattus samples (the main host), Leishmania prevalence was similar between LI and IF localities (F1.199 = 0.43, p = 0.511).

Table 2. Leishmania major and Trypanosoma lewisi infection in rodents trapped in southern Senegal.

Number of positive/total number of rodents in localities of Rattus rattus long-established invasion (LI), invasion front (FI) and no invasion (NI), and in Mereto.

Comparison of the amplicon sizes obtained for the R. rattus samples and the Leishmania reference strains of the Old World led to the identification of L. major in all positive R. rattus samples (Fig 2). However, the diagnosis could not be confirmed by direct sequencing of such amplicons. This can be explained by the high heterogeneity of the minicircle kDNA sequences. Each band on agarose gel could include different amplicons of equal size for a single Leishmania species. Diagnosis with more specific, but less sensitive molecular methods based on detection of Leishmania ITS1, HSP70 or mini-exon, as well as other methods based on kDNA amplification, also failed (S2 Fig). Therefore, the amplicons of all the positive nested PCR samples were cloned. The cloning was successful for seven samples (43 clones were successfully analysed), as well as for two L. major reference strains (Friedlin and 5ASKH) and one L. major clinical isolate (LC-DKR) (26 clones analysed for these three L. major strains) (S1 Table).

Fig 2. Results of the nested PCR assay used for the identification of Leishmania parasites in rodent samples and reference strains.

Lane 1, negative control; lane 2, L. infantum (MHOM/BL/67/ITMAP263); lane 3, L. tropica (MHOM/SU/74/K27); lane 4, L. major (MHOM/SU/1973/5ASKH); lane 5, sample 3119 (negative); lanes 6 to 8, samples 3394, 3441 and 3493 (positive).

Twenty of 26 sequences from reference strains displayed homologies with sequences in databases. Interestingly, six sequences of excellent quality of the reference strain clones (for instance, the Friedlin-x and LC-DKR-p clones) could not be extensively matched with database sequences (<80 nucleotides), implying that kDNA sequences of L. major are not yet exhaustively present in databases. In addition, six sequences from the reference strains displayed homology with kDNA sequences from other strains of the same species (for instance, the Friedlin-v clone with the LC-DKR-v clone, with 84% similarity on 572 bp) or even of other species but with low similarity (the Friedlin-d clone with Leishmania braziliensis, with 91% similarity on 53 bp, see S1 Table), in accordance with the idea that each Leishmania strain has different minicircle kDNA classes that could be either species-specific or shared with other species [54,55].

In the same way, 20 of 43 field sample clones displayed no similarity with the database sequences nor with the reference strain sequences. Nevertheless, two field sample clones (clones 3394–11 and 3394-11c) showed sequence similarity (more than 70% identity with high probability and high score) with L. major in databases (e.g. LMJLV39_SCAF000371, 73% identity in TriTrypDB on 630 bp) and 21 field sample clones, obtained from five different samples (3441, 3851, 3167, 3192, 3767) showed strong sequence similarity to the sequence of LC-DKR-u (Senegalese L. major control strain) (S1 Table). Interestingly, no clone from field sample showed similarity with sequences of clones of the other two reference L. major strains used in this study, 5ASKH (isolated in Russia) and Friedlin (isolated in Israel). In conclusion, five of the seven positive field samples tested by cloning could be identified as similar to the Senegalese L. major strain. Therefore, based on the amplicon size identity and sequence comparison, it was assumed that all the other positive samples belonged to the L. major species. Altogether, these results designated R. rattus as carrier of L. major parasites.

Leishmania diagnosis was also performed using DNA isolated from the ear of 64 rodents trapped in the south, chosen among nested PCR positive and negative samples using spleen DNA. Overall, specimens of different species, native or invasive, and the three locality categories (not invaded, recently invaded, and of long-established invasion) were represented. All tested ears were negative.

It is worth mentioning that for some samples (about 7%, generally not R. rattus samples), PCR products that were clearly smaller than the expected amplicon size for any Leishmania species (about 200-300bp, S3 Fig) were also observed. Sequence analysis of these amplicons showed similarities with GenBank rodent sequences, suggesting the unspecific amplification of the rodent host genome (S3 Fig).

Trypanosoma lewisi diagnosis

All rodents trapped in the north of Senegal (Ma. erythroleucus and M. m. domesticus) were negative for T. lewisi (FRET-based real time PCR method).

In the south including Mereto, 59 R. rattus, six M. m. domesticus, one A. niloticus, and one Ma. erythroleucus (spleen samples were positive for T. lewisi (mini-exon real time PCR method) (Table 2). Except R. rattus, all the rodents were from Mereto. No infection was detected in NI localities. In R. rattus samples (the main host), T. lewisi prevalence was similar in LI and IF localities (F1.199 = 1.11, p = 0.292).

None of the 13 R. rattus spleen samples from Mereto was positive for T. lewisi.

Some R. rattus samples were positive for both L. major and T. lewisi: 9/98 (9.2%) in LI localities, and 3/101 (3%) in IF localities (12/199, 6.0%, for both locality categories) (Table 2). In LI localities, L. major prevalence was higher among R. rattus individuals that were also infected by T. lewisi than among those that were not (26.5% vs 9.4%, GLMM F1.199 = 4.74, p = 0.029).


Leishmania major

The nested PCR allowed detecting 40 positive spleen samples with amplicons of molecular weight similar to L. major species. All tested ears were negative, suggesting that ear specimens from rats with or without skin lesions should not be used for L. major detection. Direct sequencing of the positive nested PCR to confirm diagnosis was previously used by other authors [56] or by our team [18]. However, the high heterogeneity and number of minicircle kDNA sequences existing at strain level can prevent the direct sequencing. Indeed, the polymerase may amplify only one minicircle target or several at the same time, depending on the Leishmania strain and the experimental conditions. When this happens, as in our study, cloning is necessary prior to sequencing and species identification. Sequencing of field sample clones allowed finding homologies with cloned sequences of a L. major strain isolated from a Senegalese patient (LC-DKR). The high-throughput sequencing of minicircle PCR products developed on vectors and Leishmania strains from the New World could overcome these issues [55].

Finally, 17.5% of all trapped R. rattus individuals and only two native rodents were positive for L. major in the south of Senegal (including Mereto), whereas all rodents trapped in the north were negative.

Gerbils are the main L. major reservoir: Rhombomys opimus (great gerbil) in Central Asia, Psammomys obesus (fat sand rat) in Middle East, and P. obesus and Meriones sp. in North Africa [57,58]. L. major prevalence in these species varies from 0 up to 70%, according to the diagnostic method and the season [59]. More recently, M. m. domesticus (42.9% in [60] and 7.1% in [56]) and R. rattus (12.5% [56]) also have been identified as carriers in the Middle East. So far, in West Africa, L. major was detected in native Mastomys spp., G. gambianus and A. niloticus [21,26,27,61]. In Senegal, the prevalence determined by tissue culture was about 7.3% in Ma. erythroleucus, 5.8% in G. gambianus [26], and 2.7% in A. niloticus [62]. However, no parasite by microscopic observation nor typical cutaneous lesion was identified in any rodent species in this country [63]. Among the previous studies conducted in Senegal in the 1980s, very few R. rattus individuals were found to be carriers, because this rodent species was not sampled at that time in the studied regions (villages near Dakar and Thiès, in the centre of Senegal) [21,22,27,64,65,66,67,68]. Thus, rats were not suspected to play a role in L. major transmission in Senegal.

R. rattus has been probably introduced in Senegal from Europe, where L. major has never been detected [11]. CL is known to be endemic in Senegal since 1933 [64]. Our data showing R. rattus individuals positive for L. major in Senegal suggest that invading black rats have acquired the pathogen after their introduction. The finding that native Mastomys spp. and A. niloticus individuals, which were previously described as L. major reservoirs in Senegal, were not infected in our study could be explained by many factors, such as the ecology of native rodents compared with that of R. rattus, a different susceptibility between native and invasive rodents, Leishmania ecology (transmission generally occurs within a very limited area, micro-foci [71]), L. major life cycle characteristics (wild zoonotic cycle) and/or the ecology of P. duboscqi (its main vector) that is more abundant in fields and farming areas than inside villages and towns [69]. A sampling in the surrounding of the villages invaded by R. rattus is needed to confirm the presence of a native wild rodent reservoir. Particularly, the screening of L. major in native gerbils, which are common in non-commensal habitats of southern Senegal (e.g., G. gambianus, G. guineae and Taterillus gracilis [46]), could bring information.

On the other hand, no rodent was positive for L. major in the northern region invaded only by M. m. domesticus. This could be explained by the absence of L. major in this region, or by a difference in susceptibility between mice and rats. No comparative experimental study was available, but several epidemiological works showed more cases of L. major infection in R. rattus than in M. m. domesticus [56,70]. In Mereto, the village invaded by both R. rattus and M. m. domesticus, 16.7% (n = 1/6) of A. niloticus, 3.4% (n = 1/29) of M. m. domesticus and 23.1% (n = 3/13) of R. rattus individuals were positive for L. major (Table 2), supporting again the hypothesis of a higher susceptibility of R. rattus to L. major, compared with M. m. domesticus.

Trypanosoma lewisi

We found the highest prevalence of T. lewisi among R. rattus in the south of Senegal including Mereto (29.6%). Two native rodents (n = 2) and six mice were also positive, but only in Mereto where rats and mice coexist. No infection was found in the north.

T. lewisi prevalence in R. rattus is consistent with several previous works often based on microscopic observation rather than on molecular diagnostic tools. T. lewisi has been identified in the Rattini group of rodents on all continents: Europe [32,46,65], Africa [33,34,35,36,72,73], Middle East [74], Asia [75,76,77,78,79,80,81], North [82], Central [83] and South [84,85,86,87,88] America, Hawaii [89], New Zealand [90] and Australia [91], where it caused the extinction of two native host rodent species in Christmas Island [6]. In Niger, T. lewisi prevalence is higher in R. rattus (71%) than in native rodents (6%) [33]. In the other cited studies, the reported prevalence ranges from 4.6 to 82.3%, and varies according to the season, sex and age of rodents [86,90]. Prevalence may also vary according to the tissue chosen for diagnostic. Indeed, spleen have been successfully used in previous studies because it is highly blood-supplied tissue [33,35]. Nevertheless, it may lead to underestimate prevalence levels of the parasite that poorly enters organs, compared to blood samples [37]. T. lewisi distribution and lack of positive samples among native rodents in our study strongly suggest that Senegal was T. lewisi-free before the arrival of R. rattus from Europe. The GLMM analysis did not find any significant difference in R. rattus prevalence between LI and IF localities, possibly due to the large variability at IF localities (Table 2). Indeed, T. lewisi was not detected in two IF localities (SOU, BOU: Fig 1), whereas its prevalence in the other two (KED, BAN, Fig 1, Table 2) was comparable to that in LI localities. These differences may be related to the variable R. rattus introduction times in IF areas. Indeed, Soutouta (SOU) and Boutougoufara (BOU) are marginal rural localities that have undoubtedly been colonized very late by R. rattus, because their connection to commercial networks dates from 2007 [92,93]. Conversely, Kedougou (KED) and Badi Nieriko (BAN) were colonized by R. rattus in the 1990s [41]. The prevalence heterogeneity at the IF is consistent with the hypothesis of T. lewisi introduction by R. rattus, and reflects classical enemy loss [49] in localities that have been very recently invaded. T. lewisi infection has been associated with a large mortality rate in non-Rattini species [35], which could lead to under-estimation of parasite transmission in these hosts. If this parasite affects severely native rodents, its “spill-over” [49] from the host that introduced it would confer to rats a strong advantage at the IF.

Like for L. major, the absence of T. lewisi infection among rodents in the north may suggest a lower sensitivity of mice. Mice and rats are both used as laboratory models for T. lewisi infection, but mouse experimental infection sometimes fails, while rat infection is more reproducible [76,94]. At first glance, the data obtained in Mereto challenge this idea: in Mereto 21% of M. m. domesticus and two native rodents (A. niloticus and Ma. erythroleucus) were infected, while no R. rattus (0/13) was positive for T. lewisi. Trypanosoma musculi, another trypanosomatid species very close to T. lewisi, could be responsible for the infection [95]. Indeed, T. lewisi and T. musculi, morphologically indistinguishable, were characterized on the basis of host specificity, rats being preferential hosts for T. lewisi and mice for T. musculi [95]. Moreover, these two species appear also difficult to distinguish on a molecular point of view [95,96]. We first tested the molecular methods used in our study for T. lewisi diagnosis on the reference strains of T. lewisi and T. musculi (Partinico II strain kindly provided by Pr Philippe Vincendeau and Mrs Pierrette Courtois). The gel electrophoresis banding patterns and the sequencing did not allow distinguishing these two parasites (S4A and S4B Fig). Secondly, the only published PCR protocol able to differentiate T. lewisi from T. musculi [96] was also tested. The results obtained in our assays were non reproducible, non-specific and not sensitive enough (S4C and S4D Fig). Furthermore, the only two sequences available for T. musculi in Genbank database make difficult the design of a new specific and more sensitive molecular diagnostic method. Therefore, at this stage, we cannot deduce which of T. musculi or T. lewisi is the parasite responsible for infection in Mereto positive mice.

The prevalent pattern of T. lewisi-positive R. rattus co-infected with L. major in LI suggests positive interactions between the two parasites. Previous studies showed that T. lewisi could weaken the immune system of its host and thus favour the acquisition of other infections, such as Toxoplasma gondii [97,98] or Cryptococcus neoformans [99].

Putative consequences on human health

Data about L. major distribution in Senegal are still limited, with the exception of the Dakar region [23] and nearby localities [21,22], where it is endemic. To date, no data is available about human cases caused by L. major in the north nor in the south of this country. Classically, the transmission of L. major takes place essentially in rural environments, peridomestic and farming areas, where the burrows are, with an epidemiological cycle involving native rodents as main reservoirs [18,21,100]. We can imagine that a second transmission cycle involving R. rattus might occur predominantly inside villages and indoors. The increasing presence of R. rattus, which has a behaviour strongly linked to human activities and can move and settle in new territories, engenders a risk of emergence of new spots of human CL. It would be interesting to determine the number of cases and spatial distribution of human CL in Senegal, and to assess the available means and knowledge for its diagnosis.

Only nine cases of T. lewisi infection in humans have been reported in the world, in Malaysia [101], India [76,102,103,104,105], Gambia [36] and Thailand [106]. Patients were often immunologically weak infants, living in poor hygiene conditions, and in close contact with contaminated rats in and around houses [36,76,101]. Symptoms were generally mild, except in one child [105]. The diagnosis was mostly based on the morphological identification of T. lewisi by microscopic examination of blood drops [76,101,102,103,104], and was rarely confirmed with molecular tools [36,105,106]. However, human infections could be underestimated, because the identified patients lived far from health centres and T. lewisi infection is usually associated with non-specific and transient symptoms (fever, lethargy, anorexia) [30]. Very few epidemiological data on humans have been published and no routine and specific serological tests are available yet [107]. Nevertheless, 12 of the 187 farmers tested were serologically positive in China, without any apparent symptom [108], reflecting the existence of asymptomatic carriers. The risk for immunologically weak people, such as patients with AIDS, remains to be evaluated. Given the high prevalence of T. lewisi in rodents in the south of Senegal, it could be interesting to assess the seroprevalence in humans in this region. This could be performed by the international network for atypical human trypanosomiases that has been set up in Africa [107].

In summary, we identified the black rat R. rattus as a potential reservoir for L. major and T. lewisi in the southern part of Senegal. These two infections appeared to obey to two different models. The invader R. rattus could have been be locally infected by the endemic parasite L. major, and is potentially more susceptible than native commensal rodents. Conversely, T. lewisi infection could have been introduced in Senegal by R. rattus, but seems to be poorly transmitted to native rodents by R. rattus, although this point remains to be investigated. The high prevalence of both parasites in R. rattus, which is anthropogenic and relentlessly gaining new territories, could increase the risk of transmission/emergence of new foci of human CL in urban areas and of sporadic cases of human trypanosomiasis.

Supporting information

S1 Table. BLAST analysis of the sequences obtained by PCR amplification for Leishmania detection.

For Leishmania diagnosis, the nested PCR method to amplify the minicircle kinetoplastic DNA (kDNA) variable region described by Noyes et al. [51] was used, and then amplicons were separated by electrophoresis before cloning. * BLAST results were retained only if they corresponded to the sequences of reference strains or to sequences of isolates described in peer-reviewed articles and well characterized in terms of species, origin and hosts, in order to avoid misidentifications. ** When the results obtained with one database were highly significant and with high score whereas low similarity was found with the second one, only the highly significant results are provided.


S1 Text. Sequences obtained by PCR amplification for Leishmania detection.


S2 Text. Sequences obtained by the mini-exon PCR amplification for Trypanosoma lewisi detection.

All the sequences presented were obtained from R. rattus samples.


S1 Fig. Study of the FRET-based real time PCR on 18S rRNA used for T. lewisi detection.

(A) Cp values of amplification of a T. lewisi DNA scale and of other trypanosomatids. (B) Efficiency curve. This method was chosen for its robustness and sensitivity (<9.5fg/μl). However, this PCR was not specific for T. lewisi, but detected also other trypanosomes as well as Leishmania.


S2 Fig. Negative results obtained with other molecular methods tested to confirm Leishmania diagnosis.

Field samples were chosen among samples that were positive with the nested PCR used for the Leishmania diagnosis and tested with other molecular methods. The sensitivity of these protocols was not sufficient to confirm the diagnosis made with the nested PCR on kDNA minicircles.W stands for water (negative control). Primers were the following: ITS1: LITSR, ITS1R [109]; nested ITS1 step 1: LITSR, LITSV, step 2: LITSR, L5.8S [110]; HSP70: forward [111], HSP70ant [112]; mini-exon: Fme, Rme [113]; ITS2: LGITSF2, LGITSR2 [114]; rDNA: rDNA-10F, rDNA-14R [115]; kDNA: L.MC-1S, L.MC-1R [116]; kDNA (real-time PCR HRM): MLF, MLR [117].


S3 Fig. Unspecific amplification of rodent DNA with the nested PCR on kDNA minicircles.

(A) Exampe of electrophoresis gel showing unspecific amplification obtained from a few field samples (7%) with the nested PCR on kDNA minicircles (samples marked with *, lower bands). (B) Example of sequence obtained from the sample CB3834 and presenting 85% homology on 406b with mouse DNA sequences found in public databases (for example GenBank AL772311.19). Overall, seven amplification products from seven field samples were directly sequenced and showed homology with rodent DNA.


S4 Fig. Tests for T. musculi as another possible interpretation for the T. lewisi positive results in Mus musculus domesticus samples from Mereto.

(A) Electrophoresis gel of the T. lewisi mini-exon PCR performed on T. musculi and other reference strains. Our T. lewisi diagnosis was not strictly specific and could amplify T. musculi. (B) Sequencing of the mini-exon PCR product from T. musculi. There were too many variability and too few data in databases to differentiate T. lewisi and T. musculi on the basis of mini-exon sequencing. (C) Electrophoresis gel of amplification of different reference strains with T. musculi kDNA maxi-circles PCR (upper part). The four T. musculi samples were extracted independently from the same blood sample of an infected mouse. Primers used were TM1F, TM1R [96]. In the lower part, amplification of the same samples with the 18S rRNA PCR showed that the T. musculi kDNA maxi-circles PCR seemed to be not very sensitive. (D) Electrophoresis gel of T. musculi kDNA maxi-circles performed on field samples (upper part: rat samples; lower part: mouse samples). This PCR gave frequent (33/66, 50%) non specific amplification, probably from rodents DNA. We obtained no band of size corresponding to specific amplification.



We are indebted to all Senegalese people who allowed rodent trapping in their homes. We thank S. Piry for his help to create Fig 1. We thank Prof. Babacar Faye (Service de Parasitologie–Mycologie, Faculté de Médecine et Pharmacie, Université Cheikh Anta Diop, Dakar, Senegal) for providing the Leishmania major Senegalese strain (LC-DKR) and Prof. Philippe Vincendeau and Mrs Pierrette Courtois (UMR 177 IRD/CIRAD/Université de Bordeaux, INTERTRYP, Bordeaux, France) for providing Trypanosoma musculi strain (Partinico II). We also thank Elisabetta Andermarcher for assistance in preparing and editing the manuscript.


  1. 1. Wardle DA, Bardgett RD, Callaway RM, Van der Putten WH. Terrestrial ecosystem responses to species gains and losses. Science. 2011;332(6035):1273–7. pmid:21659595
  2. 2. Young HS, Parker IM, Gilbert GS, Sofia Guerra A, Nunn CL. Introduced Species, Disease Ecology, and Biodiversity-Disease Relationships. Trends Ecol. Evol. 2017;32(1):41–54. pmid:28029377
  3. 3. Dunn AM, Hatcher MJ. Parasites and biological invasions: parallels, interactions, and control. Trends Parasitol. 2015;31(5):189–99. pmid:25613560
  4. 4. Poulin R, Paterson RA, Townsend CR., Tompkins DM, Kelly DW. Biological invasions and the dynamics of endemic diseases in freshwater ecosystems: Invasions and freshwater diseases. Freshw. Biol. 2011;56(4):676–88.
  5. 5. de Bruyn PJN, Bastos ADS, Eadie C, Tosh CA, Bester MN. Mass Mortality of Adult Male Subantarctic Fur Seals: Are Alien Mice the Culprits? PLoS One. 2008;3(11):e3757. pmid:19018284
  6. 6. Wyatt KB, Campos PF, Gilbert MT, Kolokotronis SO, Hynes WH, DeSalle R, et al. Historical mammal extinction on Christmas Island (Indian Ocean) correlates with introduced infectious disease. PLoS One. 2008;3(11):e3602. pmid:18985148
  7. 7. Harris DB. Review of negative effects of introduced rodents on small mammals on islands. Biol. Invasions. 2009;11(7):1611–30.
  8. 8. Hayman DTS, McDonald KD, Kosoy MY. Evolutionary history of rat-borne Bartonella: the importance of commensal rats in the dissemination of bacterial infections globally. Ecol. Evol. 2013;3(10):3195–203. pmid:24223261
  9. 9. Billeter SA, Borchert JN, Atiku LA, Gage KL, Kosoy MY. Bartonella Species in Invasive Rats and Indigenous Rodents from Uganda. Vector-Borne Zoonotic Dis. 2014;14(3):182–8. pmid:24575846
  10. 10. Alvar J, Vélez ID, Bern C, Herrero M, Desjeux P, Cano J, et al. Leishmaniasis worldwide and global estimates of its incidence. PLoS One. 2012;7:e35671. pmid:22693548
  11. 11. WHO Expert Committee on the Control of the Leishmaniases & World Health Organization. Control of the leishmaniases: report of a meeting of the WHO Expert Committee on the Control of Leishmaniases. World Health Organization. 2010.
  12. 12. Bañuls AL, Hide M, Prugnolle F. Leishmania and the leishmaniases: a parasite genetic update and advances in taxonomy, epidemiology and pathogenicity in humans. Adv. Parasitol. 2007;64:1–109. pmid:17499100
  13. 13. Akhoundi M, Kuhls K, Cannet A, Votypka J, Marty P, Delaunay P, et al. A Historical Overview of the Classification, Evolution, and Dispersion of Leishmania Parasites and Sandflies. PLoS Negl. Trop. Dis. 2016;10(3):e0004349. pmid:26937644
  14. 14. Ranque P, Bussiéras J, Chevalier JL, Quilici M, Mattei X. Present importance of dog leishmaniasis in Senegal. Value of immunologic diagnosis. Possible incidence in human pathology. Bull. Académie Natl. Médecine. 1970;154(21):510–2.
  15. 15. Fraga J, Montalvo AM, De Doncker S, Dujardin JC, Van der Auwera G. Phylogeny of Leishmania species based on the heat-shock protein 70 gene. Infect. Genet. Evol. 2009;10(2):238–45. pmid:19913110
  16. 16. Faye B, Bucheton B, Bañuls AL, Senghor MW, Niang AA, Diedhiou S, et al. Seroprevalence of Leishmania infantum in a rural area of Senegal: analysis of risk factors involved in transmission to humans. Trans. R. Soc. Trop. Med. Hyg. 2011;105(6):333–40. pmid:21543098
  17. 17. Faye B, Bañuls AL, Bucheton B, Dione MM, Bassanganam O, Hide M, et al. Canine visceral leishmaniasis caused by Leishmania infantum in Senegal: risk of emergence in humans? Microbes Infect. Inst. Pasteur. 2010;12(14–15):1219–25.
  18. 18. Senghor MW, Niang AA, Depaquit J, Ferté H, Faye MN, Elguero E, et al. Transmission of Leishmania infantum in the Canine Leishmaniasis Focus of Mont-Rolland, Senegal: Ecological, Parasitological and Molecular Evidence for a Possible Role of Sergentomyia Sand Flies. PLoS Negl. Trop. Dis. 2016;10(11):e0004940. pmid:27806051
  19. 19. Bañuls AL, Bastien P, Pomares C, Arevalo J, Fisa R, Hide M. Clinical pleiomorphism in human leishmaniases, with special mention of asymptomatic infection. Clin. Microbiol. Infect. 2011;17:1451–1461. pmid:21933304
  20. 20. Kimutai A, Kamau Ngure P, Kiprotich Tonui W, Muita Gicheru M, Bonareri Nyamwamu L. Leishmaniasis in northern and western Africa: a review. African Journal of Infectious Diseases. 2009;3(1):14–25.
  21. 21. Dedet JP, Hubert B, Desjeux P, Derouin F. Ecology of a cutaneous leishmaniasis focus in the Thies region (Senegal, West Africa). 5. Spontaneous infection and disease reservoir role of various wild rodent species]. Bull. Société Pathol. Exot. Ses Fil. 1981;74(1):71–7.
  22. 22. Blanchot M, Lusina D, Beunier E. Interepidemic surveillance of a cutaneous leishmaniasis focus in Senegal. Médecine Trop. Rev. Corps Santé Colon. 1984;44(1):35–40.
  23. 23. Ndongo S, Dieng MT, Dia D, Sy TN, Leye A, Diop MT, et al. Cutaneous leishmaniasis in hospital area: epidemiological and clinical aspects, about 16 cases. Dakar Méd. 2004;49(3):207–10. pmid:15776620
  24. 24. Boakye DA, Wilson M, Kweku M. A review of leishmaniasis in west Africa. Ghana Med. J.2005;39(3):94–7. pmid:17299551
  25. 25. WHO/Department of Control of Neglected Tropical Diseases. Global leishmaniasis update, 2006–2015: a turning point in leishmaniasis surveillance. Weekly epidemiological record. 2017.
  26. 26. Dedet JP, Derouin F, Hubert B, Schnur LF, Chance ML. Isolation of Leishmania major from Mastomys erythroleucus and Tatera gambiana in Senegal (West Africa). Ann. Trop. Med. Parasitol. 1979;73(5):433–7. pmid:534447
  27. 27. Kamara Bomba L, Saf’ianova VM, Goncharov DB, Emel’ianova LP. The serological examination of the population for leishmaniasis and the detection of Leishmania in rodents in the Republic of Guinea. Med. Parazitol. (Mosk.). 1992;2:42–46.
  28. 28. Dedet JP, Desjeux P, Derouin F. Ecology of a focus of cutaneous leishmaniasis in the area of Thies, Senegal, West Africa. 4. Spontaneous infestation and biology of Phlebotomus duboscqi Neveu-Lemaire 1906. Bull. Société Pathol. Exot. Ses Fil. 1980;73(3):266–6.
  29. 29. Ba Y, Trouillet J, Thonnon J, Fontenille D. Phlébotomes du Sénégal (Diptera-Psychodidae): peuplement et dynamique des populations de la région de Mont-Rolland. Parasite. 1998;5:143–150. pmid:9754310
  30. 30. Truc P, Büscher P, Cuny G, Gonzatti MI, Jannin J, Joshi P, et al. Atypical human infections by animal trypanosomes. PLoS Negl. Trop. Dis. 2013;7(9): e2256. pmid:24069464
  31. 31. Lun ZR, Reid SA, Lai DH, Li FJ. Atypical human trypanosomiasis: a neglected disease or just an unlucky accident? Trends Parasitol. 2009;25(3):107–8. pmid:19162551
  32. 32. Karbowiak G, Wita I. The cases of infection of brown rats Rattus norvegicus with Trypanosoma lewisi (Kent, 1880) Laveran i Mesnil, 1901, in the area of Warsaw agglomeration. Wiad. Parazytol. 2001;47(3):377–82. pmid:16894750
  33. 33. Dobigny G, Poirier P, Hima K, Cabaret O, Gauthier P, Tatard C, et al. Molecular survey of rodent-borne Trypanosoma in Niger with special emphasis on T. lewisi imported by invasive black rats. Acta Trop. 2011;117(3): 183–8. pmid:21126503
  34. 34. Ugbomoiko U. Factors affecting the prevalence of protozoan parasites of small mammals in southern Nigeria. Parasitica. 1997;53:5–13.
  35. 35. Tatard C, Garba M, Gauthier P, Hima K, Artige E, Dossou DKHJ, et al. Rodent-borne Trypanosoma from cities and villages of Niger and Nigeria: A special role for the invasive genus Rattus? Acta Trop. 2017;171:151–8. pmid:28373037
  36. 36. Howie S, Guy M, Fleming L, Bailey W, Noyes H, Faye JA, et al. A Gambian infant with fever and an unexpected blood film. PLoS Med. 2006;3(9):e355. pmid:17002503
  37. 37. Hoare CA. The Stercoraria. Subgenus Herpetosoma Doflein 1901. In: The trypanosomes of mammals. Oxford and Edinburgh, Blackwell Scientific publications; 1972. pp. 214–239.
  38. 38. Maraghi S, Wallbanks KR, Molyneux DH. Oral transmission of trypanosomes of the subgenus Herpetosoma from small mammals. Parasitol. Res. 1995;81:693–695. pmid:8570586
  39. 39. Global Invasive Species Database (
  40. 40. Konečný A, Estoup A, Duplantier JM, Bryja J, Bâ K, Galan M, et al. Invasion genetics of the introduced black rat (Rattus rattus) in Senegal, West Africa. Mol. Ecol. 2013;22(2):286–300. pmid:23206272
  41. 41. Dalecky A, Bâ K, Piry S, Lippens C, Diagne CA, Kane M, et al. Range expansion of the invasive house mouse Mus musculus domesticus in Senegal, West Africa: a synthesis of trapping data over three decades, 1983–2014: Range expansion of the house mouse in Senegal. Mammal Rev. 2015;45(3):176–90.
  42. 42. Lippens C, Estoup A, Hima MK, Loiseau A, Tatard C, Dalecky A, et al. Genetic structure and invasion history of the house mouse (Mus musculus domesticus) in Senegal, West Africa: a legacy of colonial and contemporary times. Heredity. 2017;119(2):64–75. pmid:28353686
  43. 43. Sikes RS, Gannon WL. Guidelines of the American Society of Mammalogists for the use of wild mammals in research. J. Mammal. 2011;92(1):235–53.
  44. 44. AVMA Panel on Euthanasia. American Veterinary Medical Association. 2000 Report of the AVMA Panel on Euthanasia. J. Am. Vet. Med. Assoc. 2001;218(5):669–96. pmid:11280396
  45. 45. Joint Working Group on Veterinary Care. Guidelines for the veterinary care of laboratory animals: report of the FELASA/ECLAM/ESLAV Joint Working Group on Veterinary Care. Lab. Anim. 2008;42:1–11. pmid:18348761
  46. 46. Granjon L, Duplantier JM. Les rongeurs de l’Afrique sahélo-soudanienne. IRD Editions; 2009.
  47. 47.
  48. 48. Garenne M, Lombard J. La migration dirigée des Sereer vers les Terres Neuves (Sénégal). in La migration: changements sociaux et développement: projets de développement et politiques migratoires 317–332. ORSTOM;1988.
  49. 49. Diagne C, Ribas A, Charbonnel N, Dalecky A, Tatard C, Gauthier P, et al. Parasites and invasions: changes in gastrointestinal helminth assemblages in invasive and native rodents in Senegal. Int. J. Parasitol. 2016;46(13–14):857–69. pmid:27670366
  50. 50. Tayeh A, Tatard C, Kako-Ouraga S, Duplantier JM, Dobigny G. Rodent host cell/Lassa virus interactions: evolution and expression of α-Dystroglycan, LARGE-1 and LARGE-2 genes, with special emphasis on the Mastomys genus. Infect. Genet. Evol. 2010;10(8):1262–1270. pmid:20674789
  51. 51. Noyes HA, Reyburn H, Bailey JW, Smith D. A nested-PCR-based schizodeme method for identifying Leishmania kinetoplast minicircle classes directly from clinical samples and its application to the study of the epidemiology of Leishmania tropica in Pakistan. J. Clin. Microbiol. 1998;36(10):2877–2881. pmid:9738037
  52. 52. R Core Team, 2016. A language and environment for statistical computing. R Foundation for Statistical Computing, 2016. Available from
  53. 53. Bates D, Maechler M, Bolker B, Walker S. Linear mixed-effects models using Eigen and S4. 2017.
  54. 54. Brewster S, Barker DC. Analysis of minicircle classes in Leishmania (Viannia) species. Trans. R. Soc. Trop. Med. Hyg. 2002;96(Suppl 1):S55–63.
  55. 55. Kocher A, Valière S, Bañuls AL, Murienne J. High-throughput sequencing of kDNA amplicons for the analysis of Leishmania minicircles and identification of Neotropical species. Parasitology. 2017;1–8. Epub 2017 nov 16.
  56. 56. Davami MH, Motazedian MH, Kalantari M, Asgari Q, Mohammadpour I, Sotoodeh-Jahromi A, Solhjoo K, Pourahmad M. Molecular Survey on Detection of Leishmania Infection in Rodent Reservoirs in Jahrom District, Southern Iran. J. Arthropod-Borne Dis. 2014;8(2):139–146. pmid:26114127
  57. 57. Saliba EK, Oumeish OY. Reservoir hosts of cutaneous leishmaniasis. Clin. Dermatol. 1999;17(3):275–7. pmid:10384866
  58. 58. Aoun K, Bouratbine A. Cutaneous leishmaniasis in North Africa: a review. Parasite. 2014;21:1–14.
  59. 59. Fichet-Calvet E, Jomâa I, Ben Ismail R, Ashford RW. Leishmania major infection in the fat sand rat Psammomys obesus in Tunisia: interaction of host and parasite populations. Ann. Trop. Med. Parasitol. 2003;97(6):593–603. pmid:14511558
  60. 60. Parhizkari M, Motazedian MH, Asqari Q, Mehrabani D. The PCR-based detection of Leishmania major in Mus musculus and other rodents caught in southern Iran: a guide to sample selection. Ann. Trop. Med. Parasitol. 2011;105:319–23. pmid:21871168
  61. 61. Ikeh EI, Ajayi JA, Nwana EJ. Mastomys natalensis and Tatera gambiana as probable reservoirs of human cutaneous leishmaniasis in Nigeria. Trans. R. Soc. Trop. Med. Hyg. 1995;89(1):25–6. pmid:7747299
  62. 62. Ranque P, Quilici M, Camerlynck P. Arvicanthis niloticus (rodent, Muridae), a reservoir of the virus causing leishmaniasis in Senegal. Bull. Société Pathol. Exot. Ses Fil. 1974;67(2):167–75.
  63. 63. Niang AA, Trouillet J, Faye O. Interepidemic supervision of the Leishmania focus of Keur Moussa (Thies, Senegal). Parasite. 1998;5(1):51–9. pmid:9754297
  64. 64. Riou M, Advier M. Leishmaniose cutanée contractée au Sénégal. Bull. Société Pathol. Exot. Ses Fil. 1933;26:254–6.
  65. 65. Dedet JP, Pradeau F, de Lauture H, Philippe G, Sankalé M. Ecology of the focus of cutaneous leishmaniasis in the region of Thies (Senegal, West Africa). 3. Evaluation of the endemicity in the human population. Bull. Société Pathol. Exot. Ses Fil. 1979;72(5–6):451–61.
  66. 66. Desjeux P, Bray RS, Dedet JP, Chance M. Differentiation of canine and cutaneous leishmaniasis strains in Senegal. Trans. R. Soc. Trop. Med. Hyg. 1982;76(1):132–3.
  67. 67. Dedet JP, Desjeux P, Derouin F. Ecology of a focus of cutaneous leishmaniasis in the Thiès region (Senegal, West Africa). 4. Spontaneous infestation and biology of Phlebotomus duboscqi Neveu-Lemaire 1906. Bull. Société Pathol. Exot. Ses Fil. 1982;75(5[2]):588–98.
  68. 68. Dedet JP, Saf'Janova VM, Desjeux P, Emelyanova LP, Schnur LF, Chance ML. Ecology of a reservoir of cutaneous leishmaniasis in the region of Thiès (Senegal, West Africa). 6. Characterization and types of isolated Leishmania strains. Bull. Société Pathol. Exot. Ses Fil. 1982;75(2):155–68.
  69. 69. Senghor MW, Faye MN, Diarra K, Elguero E, Gaye O, Bañuls AL, et al. Ecology of phlebotomine sand flies in the rural community of Mont Rolland (Thiès region, Senegal): area of transmission of canine leishmaniasis. PloS One. 2011;6:e14773. pmid:21445295
  70. 70. Samy AM, Doha SA, Kenawy MA. Ecology of cutaneous leishmaniasis in Sinai: linking parasites, vectors and hosts. Mem. Inst. Oswaldo Cruz. 2014;109(3):299–306. pmid:24821060
  71. 71. Rougeron V, Bañuls AL, Carme B, Simon S, Couppié P, Nacher M, Hide M, et al. Reproductive strategies and population structure in Leishmania: substantial amount of sex in Leishmania Viannia guyanensis. Mol. Ecol. 2011;20(15):3116–27 pmid:21722225
  72. 72. Abdel-Aal A, Abou-Eisha A. The role of rats as reservoir of some internal parasites with possible public health implications in the Suez Canal area. Assiut. Vet. Med. J. 1997;37:174–85.
  73. 73. A Allam KAM, El Bassioni SO, Nour El-Hoda AM. Some field and laboratory studies on Rattus rattus subspecies and Rattus norvegicus in two Egyptian governorates of different ecological characters. J. Egypt. Soc. Parasitol. 2002;32(3):805–12. pmid:12512813
  74. 74. Molan AL, Hussein MM. A general survey of blood and tissue parasites of some rodents in Arbil province, Iraq. APMIS. 1988;Suppl.3:47–9.
  75. 75. Laha R, Hemaprasanth H, Bhatta-Charva D. Observations on prevalence of Trypanosoma lewisi infection in wild rats and a trial on its adaptation in unnatural host. J. Appl. Anim. Biol. 1997;6:5–8.
  76. 76. Verma A, Manchanda S, Kumar N, Sharma A, Goel M, Banerjee PS, et al. Trypanosoma lewisi or T. lewisi-like infection in a 37-day-old Indian infant. Am. J. Trop. Med. Hyg. 2011;85(2):221–4. pmid:21813838
  77. 77. Linxian H, Wenchuan Y, Wenfeng P, Yuguang L. A survey on the parasitic protozoa of rodents from Xiamen Offujian province and Tacheng of Xinjiang Autonomous Region, China. Wuyi Sc J. 1994;12c:144–8.
  78. 78. Tang HJ, Lan YG, Wen YZ, Zhang XC, Desquesnes M, Yang TB, et al. Detection of Trypanosoma lewisi from wild rats in Southern China and its genetic diversity based on the ITS1 and ITS2 sequences. Infect. Genet. Evol. 2012;12(5):1046–51. pmid:22449774
  79. 79. Jittapalapong S, Inpankaew T, Sarataphan N, Herbreteau V, Hugot JP, Morand S, et al. Molecular detection of divergent trypanosomes among rodents of Thailand. Infect. Genet. Evol. 2008;8(4):445–9. pmid:17904918
  80. 80. Milocco C, Kamyingkird K, Desquesnes M, Jittapalong S, Herbreteau V, Chaval Y, et al. Molecular demonstration of Trypanosoma evansi and Trypanosoma lewisi DNA in wild rodents from Cambodia, Lao PDR and Thailand. Transbound. Emerg. Dis. 2013;60(1):17–26.
  81. 81. Alias SN, Sahimin N, Edah MA, Mohd-Zain SN. Epidemiology of blood parasitic infections in the urban rat population in peninsular Malaysia. Trop. Biomed. 2014;31(2):230–40. pmid:25134892
  82. 82. Eyles DE. Incidence of Trypanosoma lewisi and Hepatozoon muris in the Norway rat. J. Parasitol. 1952;38(3):222–5. pmid:12981595
  83. 83. Calero C. Incidence of Trypanosoma lewisi, Sarcocystis muris, species of Spirochaeta and microfilarial larvae in rats in Panama city and suburbs. J. Parasitol. 1952;38(4[1]):369.
  84. 84. Herrera L, Urdaneta-Morales S. Synanthropic rodent reservoirs of Trypanosoma (Schizotrypanum) cruzi in the valley of Caracas, Venezuela. Rev. Inst. Med. Trop. São Paulo.1997;39(5):279–82. pmid:9661306
  85. 85. Maia da Silva F, Marcili A, Ortiz PA, Epiphanio S, Campaner M, Cartão-Dias JL, et al. Phylogenetic, morphological and behavioural analyses support host switching of Trypanosoma (Herpetosoma) lewisi from domestic rats to primates. Infect. Genet. Evol. 2010;10(4):522–9. pmid:20156599
  86. 86. Linardi PM, Botelho JR. Prevalence of Trypanosoma lewisi in Rattus norvegicus from Belo Horizonte, State of Minas Gerais, Brazil. Mem. Inst. Oswaldo Cruz. 2002;97(3):411–4. pmid:12048574
  87. 87. Pinto CM, Ocaña-Mayorga S, Lascano MS, Grijalva MJ. Infection by trypanosomes in marsupials and rodents associated with human dwellings in Ecuador. J. Parasitol. 2006;92(6):1251–55. pmid:17304802
  88. 88. Franjola R, Soto G, Montefusco A. Prevalence of protozoa infections in synanthropic rodents in Valdivia City, Chile. Bol. Chil. Parasitol. 1995;50(3–4):66–72. pmid:8762669
  89. 89. Kartman L. Observations on Trypanosoma lewisi and Grahamella sp. in the blood of rats from the Hamakua district, Island of Hawaii. J. Parasitol. 1954;40(5[1]):571–9.
  90. 90. Laird M. Blood parasites of mammals in New Zealand. Zool Pub Victoria U College. 1951;9:1–14.
  91. 91. Thompson CK, Godfrey SS, Thompson RCA. Trypanosomes of Australian mammals: A review. Int. J. Parasitol. Parasites Wildl. 2014;3(2):57–66. pmid:25161902
  92. 92. Lucaccioni H. Espaces, liens, et santé: dynamiques d’invasion d’un hôte de zoonoses dans un territoire en mutation. Le cas du rat noir au Sénégal oriental. M. Sc. Thesis, University of Paris Ouest. 2016.
  93. 93. Lucaccioni H, Granjon L, Dalecky A, Fossati O, Le Fur J, Duplantier JM, et al. From Human Geography to Biological Invasions: The Black Rat Distribution in the Changing Southeastern of Senegal. PloS One. 2016;11:e0163547. pmid:27661619
  94. 94. Desquesnes M, Ravel S, Cuny G. PCR identification of Trypanosoma lewisi, a common parasite of laboratory rats. Kinetoplastid Biol. Dis. 2002;1(1):1–6.
  95. 95. Hoare CA. The trypanosomes of mammals. (Blackwell, 1972).
  96. 96. Hong XK, Zhang X, Fusco OA, Lan YG, Lun ZR, Lai DH. PCR-based identification of Trypanosoma lewisi and Trypanosoma musculi using maxicircle kinetoplast DNA. Acta Trop. 2017;171:207–212. pmid:28427957
  97. 97. Guerrero OM, Chinchilla M, Abrahams E. Increasing of Toxoplasma gondii (Coccidia, Sarcocystidae) infections by Trypanosoma lewisi (Kinetoplastida, Trypanosomatidae) in white rats. Rev. Biol. Trop. 1997;45(2):877–82. pmid:9458990
  98. 98. Catarinella Arrea G, Chinchilla Carmona M, Guerrero Bermúdez OM, Abrahams E. Effect of Trypanosoma lewisi (Kinetoplastida: Trypanosomatidae) on the infection of white rats with Toxoplasma gondii (Eucoccidia: Sarcocystidae) oocysts. Rev. Biol. Trop. 1998;46(4):1121–23. pmid:10751936
  99. 99. Gross NT, Guerrero OM, Chinchilla M, Jarstrand-Hall C. Trypanosoma lewisi-induced immunosuppression: the effects on alveolar macrophage activities against Cryptococcus neoformans. Exp. Parasitol. 2006;113(4):262–6. pmid:16546172
  100. 100. Ashford R W, Bettini S. Ecology and epidemiology: Old World. In: Peters W, Killick-Kendrick R, editors. The Leishmaniases in Biology and Medicine. London: Academic Press; 1987(1). p. 365.
  101. 101. J Johnson PD. A case of infection by Trypanosoma lewisi in a child. Trans. R. Soc. Trop. Med. Hyg. 1933;26(5):467–8.
  102. 102. Shrivastava KK, Shrivastava GP. Two cases of Trypanosoma (Herpetosoma) species infection of man in India. Trans. R. Soc. Trop. Med. Hyg. 1974;68:143–4. pmid:4617938
  103. 103. Kaur R, Gupta VK, Dhariwal AC, Jain DC, Shiv L. A rare case of trypanosomiasis in a two month old infant in Mumbai, India. J. Commun. Dis. 2007;39:71–4. pmid:18338683
  104. 104. Shah I, Ali US, Andankar P, Joshi RR. Trypanosomiasis in an infant from India. J. Vector Borne Dis. 2011;48:122–3. pmid:21715739
  105. 105. Doke PP, Kar A. A fatal case of Trypanosoma lewisi in Maharashtra, India. Ann Trop Med Public Health. 2011;4:91–5.
  106. 106. Sarataphan N, Vongpakorn M, Nuansrichay B, Autarkool N, Keowkarnkah T, Rodtian P, et al. Diagnosis of a Trypanosoma lewisi-like (Herpetosoma) infection in a sick infant from Thailand. J. Med. Microbiol. 2007;56(8):1118–21.
  107. 107. Truc P, Nzoumbou-Boko R, Desquesnes M, Semballa S, Vincendeau P. Atypical human trypanosomoses. Médecine Santé Trop. 2014;24(3):249–52.
  108. 108. Liu JH, Liu AQ. Investigation of monkeys experimentally infection and human population naturally infected with Trypanosoma lewisi from Changchun region. J. Chin. Zoonosis. 1990;6:40–1.
  109. 109. Toz SO, Culha G, Zeyrek FY, Ertabaklar H, Alkan MZ, Vardarlı AT, et al. A real-time ITS1-PCR based method in the diagnosis and species identification of Leishmania parasite from human and dog clinical samples in Turkey. PLoS Negl Trop Dis. 2013;7(5):e2205. pmid:23675543
  110. 110. El Tai NO, Osman OF, E , Presber W, Schönian G. Genetic heterogeneity of ribosomal internal transcribed spacer in clinical samples of Leishmania donovani spotted on filter paper as revealed by single-strand conformation polymorphisms and sequencing. Trans R Soc Trop Med Hyg. 2000;94(5):575–9. pmid:11132393
  111. 111. Cássia-Pires R, Boité MC, D'Andrea PS, Herrera HM, Cupolillo E, Jansen AM, Roque AL. Distinct Leishmania species infecting wild caviomorph rodents (Rodentia: Hystricognathi) from Brazil. PLoS Negl Trop Dis. 2014;8(12):e3389. pmid:25503973
  112. 112. Fraga J, Montalvo AM, De Doncker S, Dujardin JC, Van der Auwera G. Phylogeny of Leishmania species based on the heat-shock protein 70 gene. Infect Genet Evol. 2010;10(2):238–45. pmid:19913110
  113. 113. Marfurt J, Nasereddin A, Niederwieser I, Jaffe CL, Beck HP, Felger I. Identification and differentiation of Leishmania species in clinical samples by PCR amplification of the miniexon sequence and subsequent restriction fragment length polymorphism analysis. J Clin Microbiol. 2003;41(7):3147–53. pmid:12843055
  114. 114. de Almeida ME, Steurer FJ, Koru O, Herwaldt BL, Pieniazek NJ, da Silva AJ. Identification of Leishmania spp. by molecular amplification and DNA sequencing analysis of a fragment of rRNA internal transcribed spacer 2. J Clin Microbiol. 2011;49(9):3143–9. pmid:21752983
  115. 115. Odiwuor SO, Saad AA, De Doncker S, Maes I, Laurent T, El Safi S, et al. Universal PCR assays for the differential detection of all Old World Leishmania species. Eur J Clin Microbiol Infect Dis. 2011;30(2):209–18. pmid:20936316
  116. 116. Kato H, Uezato H, Katakura K, Calvopiña M, Marco JD, Barroso PA, et al. Detection and identification of Leishmania species within naturally infected sand flies in the andean areas of ecuador by a polymerase chain reaction. Am J Trop Med Hyg. 2005;72(1):87–93. pmid:15728872
  117. 117. Ceccarelli M, Galluzzi L, Migliazzo A, Magnani M. Detection and characterization of Leishmania (Leishmania) and Leishmania (Viannia) by SYBR green-based real-time PCR and high resolution melt analysis targeting kinetoplast minicircle DNA. PLoS One. 2014;9(2):e88845. pmid:24551178