Steppe lemmings and Chinese hamsters as new potential animal models for the study of the Leishmania subgenus Mundinia (Kinetoplastida: Trypanosomatidae)

Tomas Becvar; Barbora Vojtkova; Lenka Pacakova; Barbora Vomackova Kykalova; Lucie Ticha; Petr Volf; Jovana Sadlova

doi:10.1371/journal.pntd.0011897

Abstract

Leishmania, the dixenous trypanosomatid parasites, are the causative agents of leishmaniasis currently divided into four subgenera: Leishmania, Viannia, Sauroleishmania, and the recently described Mundinia, consisting of six species distributed sporadically all over the world infecting humans and/or animals. These parasites infect various mammalian species and also cause serious human diseases, but their reservoirs are unknown. Thus, adequate laboratory models are needed to enable proper research of Mundinia parasites. In this complex study, we compared experimental infections of five Mundinia species (L. enriettii, L. macropodum, L. chancei, L. orientalis, and four strains of L. martiniquensis) in three rodent species: BALB/c mouse, Chinese hamster (Cricetulus griseus) and steppe lemming (Lagurus lagurus). Culture-derived parasites were inoculated intradermally into the ear pinnae and progress of infection was monitored for 20 weeks, when the tissues and organs of animals were screened for the presence and quantity of Leishmania. Xenodiagnoses with Phlebotomus duboscqi were performed at weeks 5, 10, 15 and 20 post-infection to test the infectiousness of the animals throughout the experiment. BALB/c mice showed no signs of infection and were not infectious to sand flies, while Chinese hamsters and steppe lemmings proved susceptible to all five species of Mundinia tested, showing a wide spectrum of disease signs ranging from asymptomatic to visceral. Mundinia induced significantly higher infection rates in steppe lemmings compared to Chinese hamsters, and consequently steppe lemmings were more infectious to sand flies: In all groups tested, they were infectious from the 5th to the 20th week post infection. In conclusion, we identified two rodent species, Chinese hamster (Cricetulus griseus) and steppe lemming (Lagurus lagurus), as candidates for laboratory models for Mundinia allowing detailed studies of these enigmatic parasites. Furthermore, the long-term survival of all Mundinia species in steppe lemmings and their infectiousness to vectors support the hypothesis that some rodents have the potential to serve as reservoir hosts for Mundinia.

Author summary

Leishmania parasites of the subgenus Mundinia are an emerging health and veterinary problem that should not be ignored. Being the most recent of all Leishmania described, many aspects of Mundinia biology are enigmatic. We have very scarce data on their life cycles and biology, thus proper laboratory research must be done to enable their better understanding. One of the most crucial parts of the life cycle of Leishmania is the development in the mammalian host. In the past, we worked on establishment of other laboratory models for the subgenus, but neither Arvicanthis, Mastomys, nor guinea pigs proved to be a good choice. Other authors performed experiments with BALB/c mice using various inoculation techniques, but they also failed. Here we describe the establishment of two new potential laboratory model species, Chinese hamsters and steppe lemmings, which proved to be susceptible to Mundinia and such findings will enable other scientists to continue in research of these parasites.

Citation: Becvar T, Vojtkova B, Pacakova L, Vomackova Kykalova B, Ticha L, Volf P, et al. (2024) Steppe lemmings and Chinese hamsters as new potential animal models for the study of the Leishmania subgenus Mundinia (Kinetoplastida: Trypanosomatidae). PLoS Negl Trop Dis 18(5): e0011897. https://doi.org/10.1371/journal.pntd.0011897

Editor: Nadira D. Karunaweera, University of Colombo Faculty of Medicine, SRI LANKA

Received: January 5, 2024; Accepted: April 16, 2024; Published: May 13, 2024

Copyright: © 2024 Becvar et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: The authors confirm that all data underlying the findings are fully available without restriction. All relevant data are within the paper and its Supporting Information files.

Funding: This work was supported by the project “Grant Schemes at CU” at the Univerzita Karlova v Praze (reg. no. 403 CZ.02.2.69/0.0/0.0/19_073/0016935 to TB, BV, BVK, LP and LT) and by the ERD Funds, project CePaV- 404 iP at the Ministerstvo Školství, Mládeže a Tělovýchovy (grant No. CZ.02.1.01/0.0/0.0/16_019/0000759 to JS and PV). This work was supported by European Commission, Horizon 2020 Infrastructure Infravec2 project (https://infravec2.eu/) under grant agreement No 731060. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

1. Introduction

Leishmaniases are a group of diseases caused by digenetic parasites of the genus Leishmania that naturally occur in tropical and subtropical regions around the world. These parasites are currently divided into four subgenera: Leishmania, Viannia, Sauroleishmania, and Mundinia [1]. With more than one million human cases a year, leishmaniases are among the world’s most important tropical diseases caused by protozoa. There are three main clinical manifestations of the disease: cutaneous, mucocutaneous, and visceral, with symptoms depending on many endogenous and exogenous factors, such as parasite species and virulence, infectious dose, the host immune response, and other biological, socio-economic and environmental factors [2–4].

Mundinia is the most recently described Leishmania subgenus consisting of six species, all of which have unknown reservoir hosts [5,6]. Biting midges (Diptera: Ceratopogonidae) are the most probable vectors of Mundinia, changing the established dogma that Leishmania are transmitted solely by phlebotomine sand flies (Diptera: Psychodidae) [7–10]. Three species of Mundinia do not infect humans; L. enriettii was found in domestic guinea pigs in Brazil, and even though nearly 80 years have passed since its discovery, it remains one of the most enigmatic species of Leishmania [11]. Leishmania macropodum, which causes cutaneous and mucocutaneous symptoms in kangaroos, is the only representative species of the Leishmania genus in Australia [12,13] and L. procaviensis has recently been described from hyrax (Procavia capensis) isolates in Namibia [6,14].

Other three species of Mundinia infect humans; the symptoms of the disease range from a single cutaneous lesion to general visceral infection [15–17]. Leishmania martiniquensis is the most widespread species of all known Leishmania, present in Central and North America, Europe and Asia [17–20] while L. orientalis is endemic to Southeast Asia [16,21–23]. In Thailand, a high prevalence of L. martiniquensis and several cases of L. orientalis have been described in immunosuppressed people [24], but cases of immunocompetent people showing signs of cutaneous or visceral leishmaniasis have also been reported [25]. The last member of the subgenus is recently described L. chancei [6] which causes cutaneous leishmaniasis in humans, mainly children and young adults in Ghana [15]. Based on the low number of human cases, the zoonotic circulation of this parasite is predicted [26].

Several mammalian species are currently incriminated for their potential involvement in Mundinia circulation. Black rats (Rattus rattus), cows, and horses are suggested candidates, since L. martiniquensis DNA was detected and viable parasites were cultured from these hosts [18–20,27]. However, it should be noted that detection of parasites or their DNA does not imply the reservoir potential of the host, since parasites can be present in the organism incidentally, be detectable, but not transmittable to other hosts. These organisms are so called “sinks” of infection. The opposite are so called “sources”, where parasites are detectable and further transmittable to other hosts [28].

In this study, we decided to test susceptibility to Mundinia and infectiousness to sand flies by xenodiagnoses in both a classic model laboratory animal, the BALB/c mouse, and two less established model rodent species, steppe lemming (Lagurus lagurus) and Chinese hamster (Cricetulus griseus). Laboratory mice (Mus musculus) have become one of the most widely used biological models in many areas of biomedical research, including the study of cutaneous and visceral leishmaniasis, mainly due to the wide range of inbred strains and transgenic animals. In contrast, Lagurus lagurus and Cricetulus griseus are genetically polymorphic wild rodents. They were chosen because they are commercially available and easy to maintain and several old publications [29–32] as well as more recent ones [33,34], have demonstrated their high susceptibility to various Leishmania parasites.

2. Materials and methods

2.1. Ethics statement

The animals were maintained and handled in the Charles University animal facility in Prague following institutional guidelines and Czech legislation (Act No. 246/1992 and 359/2012 coll. on Protection of Animals against Cruelty in present statutes at large), which complies with all relevant European Union and international guidelines for experimental animals. All experiments were approved by the Committee on the Ethics of Laboratory Experiments of Charles University in Prague and were performed with permission no. MSMT-7831/2020-3 of the Ministry of Education, Youth, and Sports. The investigators are certified for experimentation with animals by the Ministry of Agriculture of the Czech Republic.

2.2. Parasites and rodents

Leishmania enriettii (MCAV/BR/45/LV90), L. macropodum (MMAC/AU/2004/AM-2004), L. chancei (MHOM/GH/2012/GH5), L. orientalis (MHOM/TH/2014/LSCM4) and four strains of L. martiniquensis (MHOM/MQ/1992/MAR1; MHOM/TH/2011/Cu1R1; MHOM/TH/2019/Cu2 and MEQU/CZ/2019/Aig1) were used. Parasites were maintained at 28°C in M199 medium supplemented with 20% fetal calf serum (Gibco, Prague, Czech Republic), 1% BME vitamins (Sigma-Aldrich, Prague, Czech Republic), 2% sterile urine and 250 μg/ml amikacin (Amikin, Bristol-Myers Squibb, Prague, Czech Republic). All strains were kept in a cryobank with 2–3 subpassages in vitro prior to experimental infections of rodents. Before experimental infection, the parasites were washed by centrifugation (1400×g/5 min) and resuspended in saline solution.

BALB/c mice (Velaz, Prague, Czech Republic), steppe lemmings and Chinese hamsters (Karel Kapral s.r.o., Prague, Czech Republic) were maintained in groups of 5 specimens in T4 boxes (58 × 37 × 20 cm) (Velaz, Prague, Czech Republic), equipped with bedding (SubliCZ, Sojovice, Czech Republic), breeding material (Woodwool, Miroslav Vlk s.r.o., Czech Republic) and hay (Krmne smesi Kvidera, Spalene Porici, Czech Republic), provided with a feed mixture ST-1 (Krmne smesi Kvidera, Spalene Porici, Czech Republic) and water ad libitum, with a 12 h light/12 h dark photoperiod, temperature of 22–25°C and relative humidity of 50–60%.

2.3. Experimental infections and xenodiagnoses of rodents

Eighty specimens of three rodent species (Mus musculus, Cricetulus griseus and Lagurus lagurus) anesthetised with ketamin/xylazin (62 mg/kg and 25 mg/kg) were injected with 10⁷ stationary-stage promastigotes (from 5–7 days old cultures, depending on the species) in 5 μl of sterile saline solution intradermally into the left ear pinnae. The ear pinna was chosen because it is the most appropriate site for monitoring the development of lesions and it is also the site where sand flies naturally bite, therefore suitable for xenodiagnoses [33,35,36]. The course of infection was recorded weekly.

Xenodiagnoses were performed at weeks 5, 10, 15 and 20 p.i. using Phlebotomus duboscqi. Five to six-day-old sand flies were placed in plastic vials covered with fine nylon mesh and allowed to feed on the ear pinnae of anesthetised animals. The proven vectors of Mundinia are unknown and a laboratory colony of the permissive species Culicoides sonorensis (Pirbright institute, UK) is not available for routine experiments in the Czech Republic, but early infections in vectors are nonspecific [37,38], so Mundinia can multiply before defecation of the sand fly, which in P. duboscqi is on day 4–5 PBM [39]. The engorged individuals were maintained for two days at 25°C and then stored in tissue lysis buffer (Roche, Prague, Czech Republic) at -20°C in pools of 5 females for subsequent DNA isolation and nested PCR. Specimen for xenodiagnoses were chosen randomly at the beginning of the experiment and the same animals were used for all xenodiagnoses trials.

At the end of the experiments (20 weeks p.i.) the rodents were euthanised, dissected, and tissues from ears, paws, tail, ear-draining lymph nodes, spleen, and liver were stored at − 20°C for subsequent analyses of parasite load (DNA isolation, nested PCR and qPCR).

2.4. Morphometry of parasites

The inocula were fixed with methanol, stained with Giemsa, and examined under the light microscope with an oil-immersion objective. One hundred and thirty randomly selected promastigotes from each infection dose were photographed using the QuickPHOTO MICRO programme for further analysis in the ImageJ programme. The body and flagellar lengths of the parasites were measured and metacyclic forms were distinguished, based on the criteria of Walters (1993) and Cihakova and Volf (1997) [40,41]: body length < 14 μm and flagellar length 2 times body length.

2.5. DNA isolation and nested PCR of sand fly and rodent samples

Total DNA isolated using the High Pure PCR Template Preparation Kit (Roche Diagnostics, Indianapolis, IN) according to the manufacturer’s instructions was used as a template for nested PCR amplification (detection limit 10⁻² parasites per sample) with outer primers amplifying 332 bp region of 18S sequence (forward primer 18SN1F 5’- GGA TAA CAA AGG AGC AGC CTC TA3’ and reverse primer 18SN1R 5’–CTC CAC ACT TTG GTT CTT GAT TGA-3’) and inner primers amplifying the 226 bp long 18S sequence (forward primer 18SN2F 5′-AGA TTA TGG AGC TGT GCG ACA A-3′ and reverse primer 18SN2R 5′-TAG TTC GTC TTG GTG CGG TC-3′) previously used by Sadlova et al. 2022 [42]. The samples were then analysed using 0,8% agarose gel. The reaction mixtures and cycling conditions were as follows:

1. step of PCR: 3 μl of genomic DNA, 0.5 μl forward primer 18SN1F (10 μM), 0.5 μl reverse primer 18SN1R (10 μM), 10 μl of 2x EmeraldAmp GT PCR Master Mix (Takara Bio), 6 μl of ddH2O. Step 1, 94°C for 3 min 30 s; step 2, 94°C for 30 s; step 3, 60°C for 30 s; step 4, 72°C for 25 s; step 5, 72°C for 7 min; followed by cooling at 12°C. Steps 2–4 were repeated 35 times.

2. step of PCR: 1 μl of 1. step PCR reaction, 0.5 μl forward primer 18SN2F (10 μM), 0.5 μl reverse primer 18SN2R (10 μM), 10 μl 2x EmeraldAmp GT PCR Master Mix (Takara Bio), 8 μl ddH2O. Step 1, 94°C for 30 s; step 2, 94°C for 30 s; step 3, 60°C for 30 s; step 4, 72°C for 20 s; step 5, 72°C for 7 min; followed by cooling at 12°C. Steps 2–4 were repeated 35 times.

2.6. Quantitative PCR

Parasite quantification by quantitative PCR (qPCR) was performed in a Bio-Rad iCycler & iQ Real-Time PCR Systems using the SYBR Green detection method (SsoAdvanced Universal SYBR Green Supermix, Bio-Rad, Hercules, CA). Primers targeting the 226 bp long 18S sequence (forward primer 18SN2F 5′-AGA TTA TGG AGC TGT GCG ACA A-3′ and reverse primer 18SN2R 5′-TAG TTC GTC TTG GTG CGG TC-3′) were used. One microlitre of DNA was used per individual reaction. PCR amplifications were performed in duplicates using the following conditions: 98°C for 2:30 min followed by 40 repetitive cycles: 98°C for 10 s and 60°C for 20 s. PCR water was used as a negative control. Detection limit of used assay is 10³ parasites per sample. A series of 10-fold dilutions of L. martiniquensis promastigote DNA, ranging from 5 × 10⁶ to 5 × 10¹ parasites per PCR reaction, was used to prepare a standard curve. Quantitative results were expressed by interpolation with a standard curve. To monitor non-specific products or primer dimers, a melting analysis was performed from 70 to 95°C at the end of each run, with a slope of 0.5°C/c, and 6 s at each temperature. The parasite loads in tested tissues were classified into three categories: low, < 1000 parasites; moderate, 1000–10 000 parasites; heavy, > 10 000 parasites.

2.7. Statistical analysis

Differences in infection rates were analysed using Fishers Exact Tests in SPSS version 27.

3. Results

3.1. Development of Mundinia in BALB/c mice

Two independent trials, each with groups of 5 animals inoculated with culture-derived Mundinia species, were processed; in total, 10 BALB/c mice infected with each Mundinia species (L. enriettii, L. macropodum, L. orientalis, L. chancei, and four strains of L. martiniquensis) were studied. The representation of the infective metacyclic stages in the inocula ranged between 0–32% (S1 Table).

Mice did not show weight loss or external signs of infection throughout the experiment (S2 Table). Xenodiagnoses were performed at weeks 5, 10, 15 and 20 p.i. on 4/10 mice from each group and revealed no positive results by PCR. Also, all examined tissue samples (i.e., ears, paws, tail, ear-draining lymph nodes, spleen and liver) were negative for the presence of Leishmania DNA by PCR (S3 Table). In summary, BALB/c mice were resistant to all Mundinia species tested.

3.2. Development of Mundinia in Chinese hamsters (Cricetulus griseus)

In total, 10 Chinese hamsters were infected with each of the culture-derived Mundinia tested: L. enriettii, L. macropodum, L. orientalis, L. chancei, and four strains of L. martiniquensis. In total, 80 animals were used. The representation of infectious metacyclic stages in the inocula ranged between 0–57% (S1 Table)

Similar to BALB/c mice, Chinese hamsters did not show external signs of infection or weight loss (S2 Table) during the experiment. However, hamsters infected with L. enriettii, L. chancei, L. orientalis, and L. martiniquensis were infectious to sand flies throughout the experiment from week 5 p.i. to week 15 p.i. in L. enriettii and to week 20 p.i. in L. chancei, L. orientalis, and L. martiniquensis, when the experiment was terminated (Table 1). Always, 4/10 animals per group were used for xenodiagnoses trials. Nested PCR showed positivity of the inoculated ears in 6/8 tested groups at the end of the experiment, only samples from animals infected with L. macropodum and L. martiniquensis Aig1 were negative. Leishmania enriettii DNA was found in the inoculated ear of one animal and also in draining lymph nodes of the inoculated ear of another animal. The inoculated ears of animals infected with L. chancei were positive in 30% and parasites were not detected in any other tissue. In hamsters infected with L. orientalis, 40% of the ears were positive by nested PCR, while in one case the parasites were detected in the draining lymph nodes of the inoculated ear. Animals infected with L. martiniquensis MAR1 showed 20% of inoculated ears and draining lymph nodes positive for parasite DNA. In animals infected with L. martiniquensis Cu1R1, 30% of the inoculated ears were positive by nested PCR while no parasite DNA was detected in any other tissue. The highest proportion of infected animals was found in the group infected with L. martiniquensis Cu2 where 80% of the inoculated ears were positive and parasite DNA was also detected in 10% of the hind paws (Fig 1). Quantitative PCR showed that only a low numbers of parasites (<1000) were present in all the tissues tested (S3 Table).

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Fig 1. Anatomical distribution of parasites in Chinese hamsters determined by nested PCR.

The results are presented by the balloon graph where the size corresponds to the percentage of infected tissues from the total sum of tested organs of the same type. IE = inoculated ears; CE = contralateral ears; LN-IE = draining lymph nodes of inoculated ears; LN-CE = draining lymph nodes of the contralateral ears.

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Table 1. Infectiousness of tested Chinese hamsters to sand flies during xenodiagnoses.

https://doi.org/10.1371/journal.pntd.0011897.t001

3.3. Development of Mundinia in steppe lemmings (Lagurus lagurus)

As with the previous host species, groups of 5 steppe lemmings were inoculated with five culture derived species of Mundinia (L. enriettii, L. macropodum, L. orientalis, L. chancei, and four strains of L. martiniquensis) and the experiment was repeated twice; in total, 10 individuals of steppe lemmings were infected by each parasite species. The proportion of metacyclic stages in inocula ranged between 0–25% (S1 Table).

In steppe lemmings, most Mundinia species and strains produced significantly higher infection rates than in Chinese hamsters (Figs 2 and 3). The pathological signs of infection (Figs 4 and 5) including weight changes (S2 Table) in steppe lemmings varied from asymptomatic through cutaneous to visceral form. However, specimens in all groups were infectious to sand flies from week 5 p.i. until the end of the experiment by week 20 p.i. (Table 2).

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Fig 2. Infection rates in Chinese hamsters (Cricetulus griseus) and steppe lemmings (Lagurus lagurus) infected with Mundinia.

The numbers of animals evaluated are shown in the graph. P values indicate difference between host species; n.s., nonsignificant difference, *, P < 0.05, **, P < 0.01, ***, P < 0.0001.

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Fig 3. Anatomical distribution of parasites in steppe lemmings determined by nested PCR.

The results are presented by the balloon graph where the size corresponds to the percentage of infected tissues from the total sum of tested organs of the same type. IE = inoculated ears; CE = contralateral ears; LN-IE = draining lymph nodes of inoculated ears; LN-CE = draining lymph nodes of the contralateral ears.

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Fig 4. Pathological changes caused by Mundinia in the steppe lemming (Lagurus lagurus).

A) healthy ear before inoculation; B) lesion formation in animal infected with L. martiniquensis Cu2 12 weeks p.i.; C) dry lesion in animal infected with L. martiniquensis Cu2 15 weeks p.i.; D) partial pinna necrosis in animal infected with L. chancei 13 weeks p.i.; E) destruction of the ear and dissemination of parasites around the eye in animal infected with L. martiniquensis Cu2 19 weeks p.i.; F) hair loss in abdomen of animal infected with L. chancei 12 weeks p.i. (animal on the right).

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Fig 5. Graphical table showing pathological changes observed in steppe lemmings (Lagurus lagurus) infected with Mundinia.

The categories were: asymptomatic (white); swelling (yellow); dry lesion (green); partial destruction of the pinna (orange); destruction of the pinna (red); abdomen hair loss (blue); weight loss (grey); death (black). Axis Y describes week post infection, and axis X shows the number codes of examined animals (each column represents one animal). Specimens used for xenodiagnoses trials are marked with * sign.

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Table 2. Infectiousness of tested steppe lemmings to sand flies during xenodiagnoses.

https://doi.org/10.1371/journal.pntd.0011897.t002

In all animals infected with L. enriettii, parasite DNA was detected in the inoculated ears and also in 30% of the draining lymph nodes of the inoculated ears. Parasite DNA was also present in 10% of the forepaws, hind paws, tails, and spleens (Fig 3). Although the number of parasites in infected ears was low or moderate (maximum 2.4x10⁴), a very high proportion of pools of sand flies used for xenodiagnoses were positive (more than 85% from week 10 p.i.) (Table 2). All animals that were used for xenodiagnoses developed cutaneous symptoms (swelling and dry lesions) on the ears from week 8 or 9 p.i., respectively, which persisted to the end of the experiment while other animals did not show any cutaneous symptoms. In one animal, which was not used for xenodiagnoses, we also observed hair loss in the abdomen from the 17^th week p.i. (Fig 3).

Animals infected with L. macropodum did not show any symptoms of infection during the experiment. We detected only low numbers of parasites (<1000) in 33% of inoculated ears (again, only in animals used for xenodiagnoses) and in the tail of one additional animal not exposed to sand flies (Fig 3). Steppe lemmings were infectious to sand flies only in 11% by week 5 p.i. and 33% by week 20 p.i. (Table 2).

In steppe lemmings infected with L. chancei, one animal died 2 weeks post infection from unknown reasons. The development of the lesion was observed in only one animal used for the xenodiagnosis. The lesion with swelling began to form by week 8, leading to partial destruction of the pinna by week 13 (Fig 4C and 4D). The loss of hair in the abdomen appeared in two specimens between 10 and 12 weeks (Fig 4F) and one of these animals died 3 weeks later. All other 6 animals showed no external signs of infection during the entire experiment. However, parasites disseminated throughout the body and were found in all tested tissues except lymphatic nodes draining non-inoculated ears, although one of the tested non-inoculated ears tested was positive for the presence of parasites (Fig 3). The positivity rate of inoculated ears, lymph nodes that drain them, hind paws and spleen was 37.5%. Although the positivity and infectiousness of the animals were high, only low or moderate numbers of parasites (<2500 per tested tissue) were detected by qPCR (S3 Table).

Among the animals infected with L. orientalis, 4/10 specimens died during the experiment, by weeks 7, 8, 13 and 15 p.i. One specimen that died by week 13 showed hair loss and weight loss, while no external signs of infection were recorded in others. Nested PCR revealed that 75% of the animals were positive for Leishmania DNA at the time of dissection. Parasites spread into all tested tissues except the tail (Fig 3), most frequently they were present in the lymph nodes (75%), forepaws (62.5%), spleen (50%) and liver (37.5%). Similarly to animals infected with L. chancei, the positivity and infectiousness of the animals were high, but parasite numbers were only low or moderate (<2500) (S3 Table).

Large differences were observed in animals infected with various strains of L. martiniquensis. Although L. martiniquensis strains Mar1, Cu1R1, and Aig1 did not cause any pathological signs of infection, animals infected with L. martiniquensis Cu2 showed the most severe cutaneous signs of infection of all tested groups. All four animals that were used for xenodiagnoses developed pathological changes in the pinnae, while none of the five animals that were not used for xenodiagnoses developed lesions. After the first round of xenodiagnoses 5 weeks p.i., swelling and lesions began to appear (Fig 4B) and continued to grow in two animals, leading to the destruction of the pinna by week 12 (Fig 4E). In another animal, the lesion started to form by week 12 and the pinna was completely destructed by week 19, and one animal lost the entire pinna between weeks 16 to 19, while no pathological signs were observed before that (Fig 5). One animal did not show any lesions, but weight loss was observed from week 10 and its overall health status degraded until the end of the experiment. Four other animals (that have not been xenodiagnosed) showed no external signs of infection and one animal died 6 weeks p.i. Parasites were detected by PCR in 88.9% (8/9) of the animals. The parasites often spread into all tissues tested (Fig 3), most frequently to lymph nodes (88.9%), forepaws (66.7%), hind paws (55.6%), and spleen (55.6%) (Fig 3). Quantitative PCR revealed only low numbers of Leishmania in most tissues tested, but higher numbers were present in 6/9 inoculated ears (7x10³–3.16x10⁵) and 4x10⁴ parasites were detected in the spleen of the animal with the highest parasite load in the inoculated ear (S3 Table). Interestingly, this specimen did not show signs of infection (S2 Table). Animals infected with L. martiniquensis Mar1 were infectious to sand flies from week 5 p.i. to the end of experiment (Table 2). Parasite DNA was found in all tested tissues except of contralateral ears, most often in inoculated ears (89%), draining lymph nodes of inoculated ears (78%), spleen (66%) and forepaws (44%) (Fig 3). Low or moderate numbers of parasites were detected in inoculated ears (maximum 3x10⁴) and only low numbers of parasites (<1000) were detected in other tested tissues (S3 Table). In animals infected with L. martiniquensis Cu1R1, parasites were detected in inoculated ears (88%), draining lymph nodes of inoculated ears (50%), draining lymph nodes of contralateral ears and tail (13%) (Fig 3). The animals were infectious to sand flies throughout the course of experiment, but only in a low proportion compared to other strains of L. martiniquensis with positive pools ratio slowly increasing over time (Table 2). In steppe lemmings infected with the Aig1 strain, parasites were found in 56% of inoculated ears and 11% of forepaws and tails (Fig 3). Although only low numbers of parasites were detected (S3 Table), the xenodiagnoses proved that animals were infectious to sand flies, even in 50% by week 20 p.i. (Table 2).

4. Discussion

Leishmaniases caused by Mundinia species are an emerging health problem that cannot be ignored. While cases of animal leishmaniasis caused by L. enriettii, L. macropodum, and L. procaviensis have been described only sporadically, human disease caused by L. martiniquensis and L. orientalis is on the rise in Southeast Asia, especially in Thailand [9,43], and cutaneous leishmaniasis caused by L. chancei is emerging in Ghana [15,25]. Since the knowledge of natural reservoirs is limited, it is necessary to develop a reliable model for experimental research on the biology of these parasites.

Among the classical laboratory models, guinea pigs (Cavia porcellus) were first used for Mundinia research. These animals have been shown to be susceptible to L. enriettii [9,44–47], however, infections with other Mundinia species were lost; only guinea pigs infected with L. orientalis and L. martiniquensis showed temporary pathological changes in the ear pinnae [47]. In golden hamsters (Mesocricetus auratus), L. enriettii showed only mild symptoms with temporary lesions at the site of inoculation and their subsequent healing [9,43] while L. martiniquensis disseminated, causing signs of VL [48]. Recently, L. chancei was tested in Mastomys natalensis and Arvicanthis niloticus as these two rodents inhabit Ghana and could potentially serve as reservoir hosts of the parasite. However, Leishmania survival in both hosts was limited, making them unsuitable experimental models [36].

In the present study, we tested three rodent species: BALB/c mouse (M. musculus), Chinese hamster (C. griseus), and steppe lemming (L. lagurus). BALB/c mice were chosen as the most commonly studied murine model, highly susceptible to cutaneous leishmaniasis caused by L. major as well as visceral leishmaniasis (reviewed by [49]). Chinese hamsters and steppe lemmings were used as these animals have been shown to be more susceptible to L. donovani than golden hamsters, the most common rodent model for visceral leishmaniasis: In experiments where the three rodent species were directly compared after intradermal inoculation of L. donovani, both Chinese hamsters and steppe lemmings showed more extensive spread of parasites over the body and higher parasite loads and infectiousness to sand flies then golden hamsters. In addition, both Chinese hamsters and steppe lemmings were also highly susceptible to cutaneous L. major [33]. We inoculated culture-derived parasites intradermally into rodent pinnae to simulate the natural mode of infection to the best of our knowledge. Indeed, the use of vector-derived parasites together with homogenate of salivary glands would be a more appropriate way to simulate natural mode of transmission [50], however, this experimental scheme was not applicable in the study since the natural vectors of these parasites are unknown.

BALB/c mice were proved to be resistant to infection with all five Mundinia species tested. At the end of the experiment, by week 20 p.i., Leishmania DNA was not detected in any of the tissues tested, and the rodents were not infectious to sand flies throughout the whole experiment. Our results were consistent with previous observations: BALB/c mice infected with L. martiniquensis strains MAR1 and MAR2 were asymptomatic and no mortality, weight loss, or clinical signs were observed in mice infected with these strains, regardless of the route of inoculation. But parasite dissemination and infection kinetics varied by inoculation method and inoculum size [51]. Somboonpoonpol (2016) [52] described the poor development of Thai L. martiniquensis after subcutaneous inoculation whereas high parasite burdens were observed after i.v. application of the parasite. Intakhan et al. (2020) [48] used intraperitoneal infection and detected parasite DNA only in the liver and spleen of infected animals in low numbers at 8 weeks p.i. followed by clearance at week 16 p.i. in most animals. These results suggest that BALB/c mice are not a suitable laboratory model to study the pathology of Mundinia infection at least after intradermal inoculation. However, there are other inbred mouse models that differ in their immune response to leishmaniasis from the BALB/c strain. It may be helpful to study the development of Mundinia, for example, in C57BL/6 mice, which, unlike the BALB/c strain, are resistant to L. major but are susceptible to L. amazonensis or L. mexicana (reviewed by [4]). Another option is to study Mundinia in outbred mouse strains that demonstrated an intermediate phenotype between resistant and susceptible inbred strains [53].

Chinese hamsters (C. griseus) were demonstrated as susceptible to L. enriettii, L. chancei, L. orientalis and three strains of L. martiniquensis (MAR1, Cu1R1, and Cu2), even though no external signs of disease were observed. All hamsters were in perfect health until the end of the experiment, a low number of parasites were localized in their inoculated ears, and the animals were sporadically infectious to sand flies. Very rarely, parasites were detectable in lymph nodes draining the inoculated ear and hind paws, but also in low numbers. These data suggest that Chinese hamsters have the potential to mimic asymptomatic infection. These asymptomatic infections seem to play a major role in the circulation of Mundinia, as no symptomatic wild hosts have yet been found in nature [5] and most human infections in Thailand are also asymptomatic [54]. On the other hand, except for L. martiniquensis Cu2 (80% of positive animals) and L. orientalis (50% of positive animals), the proportion of animals with detectable parasites was very low, which would be problematic for more robust and detailed studies.

Steppe lemmings (Lagurus lagurus) were shown to be highly susceptible to all Mundinia species tested, with severe signs of infection observed throughout the experiment in human infecting species. Animals were infectious to sand flies and parasites were detected in all tissues tested. Interestingly, only lemmings used for xenodiagnoses developed pathological signs on the ears. This is consistent with previously stated hypothesis of enhancing effect of sand fly bites to the course of infection by Vojtkova et al. (2021), who observed significantly faster and to a larger size lesion development in L. major infected BALB/c mice exposed to repeated bites of P. dubosqci compared to the unexposed group of mice, and the tissue parasite load was higher in rodents exposed to repeated bites [55]. Also in hamster model, increased transmissibility of L. donovani was observed following multiple exposures of hamsters to Lutzomyia longipalpis bites [56].

The present study demonstrated that steppe lemmings support the survival and development of L. enriettii, which caused small dry skin lesions in animals exposed to sand flies, while other animals, although positive for parasites at the end of the experiment, remained asymptomatic. Interestingly, a very high percentage of positive sand fly pools (over 90%) from xenodiagnoses was observed, although we found only a low or moderate number of parasites in the inoculated ears. Also, unlike guinea pigs, where lesions heal and reduction of infectiousness to sand flies with time was observed [47], steppe lemmings did not heal lesions over time and animals remained infectious to sand flies until the end of the experiment which brings the potential to study the non-healing phenotype of cutaneous leishmaniasis and supports our hypothesis that in nature, animals infected with Mundinia show mild signs of infection or remain asymptomatic and may serve as reservoirs for a long term.

Up to date, it was thought that L. macropodum is restricted to kangaroos and circulate in nature in these marsupials being transmitted by biting midges [7,12]. Here we prove these parasites can infect, survive and be infectious to vectors even in rodents. Steppe lemmings were infectious to sand flies in 11% by week 5 p.i., then any of the tested pools was positive for presence of parasites for 10 weeks, but even 33% of pools were positive by week 20 p.i. By this time, parasites were present in inoculated ears and rarely in the tail. Due to these findings, we hypothesise parasites can develop well in ears for limited period of time, then they are suppressed by immune system, but they can survive, slowly overcome immunological barriers, and start to multiply again. It would be interesting to perform long term experiments and observe dynamics of the infection in several time intervals.

Interestingly, only animals exposed to sand flies remained infected, whereas no parasites were observed in other animals, which also supports the hypothesis of sand fly bites influence on the course of infection.

Animals infected with L. chancei showed a mixture of cutaneous and visceral symptoms, with 7/8 animals positive in various tissues and organs. On the other hand, the xenodiagnoses were positive only sporadically. The visceralisation observed in steppe lemmings is unparalleled to human infections described in Ghana and neighbouring countries. Here, among 8000 cases of leishmaniasis none of the cases was described as visceral, only skin symptoms were observed [15]. However, the results of current study suggest that L. chancei could have the potential to visceralise, for example, in immunocompromised individuals such as HIV-positive people.

Leishmania chancei is most closely related to L. orientalis from Thailand [23], where variable human disease manifestations correlated with patients’ immunocompetence status can be observed. The type strain LSCM4 [22] used also in our study and the strain CULE5 [23] caused simple skin lesions in immunocompetent patients while in AIDS patients, the strain PCM2 Trang and the anonymous isolate from Kanchanaburi caused disseminated cutaneous leishmaniasis and visceral disease [16,21]. Steppe lemmings infected with L. orientalis displayed no skin lesions, but 4/10 of the animals died during the experiment and one more suffered from hair loss, which is a usual symptom of visceral leishmaniasis similar to their spread widely into various body tissues and organs (38% of liver and 50% of spleen were positive by PCR). Previously, a high susceptibility of steppe lemmings to progressive VL caused by L. donovani has been reported [33,34], so the present study extends the spectrum of Leishmania species for which steppe lemmings may be a suitable animal model for future studies.

This study presents a unique comparison of four strains of L. martiniquensis originating from various continents and hosts. While the strains Cu1R1 and Cu2 came from humans living in Thailand, Mar1 originates from human in Martinique and Aig1 was isolated from a horse in the Czech Republic. Leishmania martiniquensis is probably the most widespread species of Leishmania, which was hidden from scientists until the 1990s and causes various symptoms in different hosts. While the Cu2 strain was destructive to ear pinnae of animals and was present in moderate or high numbers, other strains did not cause any symptoms for the whole course of experiment and they were present only in low numbers in tissues. Interestingly, strains Cu1R1 and Cu2 coming from the same area developed differently in steppe lemmings: while Mar1 and Cu2 visceralised, strains Cu1R1 and Aig1 remained mainly in inoculated ears or their draining lymph nodes. These differences point to polymorphisms between strains within the species similar to those observed in L. major [57].

Mundinia is remarkable, important and poorly described subgenus of the genus Leishmania. Its worldwide distribution, wide spectrum of hosts, various clinical manifestations, and probable involvement of unusual vectors add more mystery to the whole story. Unravelling questions concerning vectors, reservoir hosts, etc. is a prerequisite for describing the life cycle of these parasites. The use of new animal models, such as steppe lemmings and Chinese hamsters, can contribute significantly to this goal. In general, wild rodents represent a novel approach to the study of host competence and host-parasite relationships [4]. The use of only one inbred genotype may not correspond to the natural variability in parasite’s tissue tropism, the dynamics and duration of infection or the infectiousness to vectors. Since the manifestations of human diseases caused by Mundinia are variable and show almost as polymorphic disease course as those caused by Leishmania and Viannia subspecies, it would be optimal to introduce a wider range of model animals. Conversely, for studies aimed at drug testing or the development of diagnostic methods, it would be preferable to use less genetically variable models, either obtained by long-term inbreeding of these rodent species or by searching among existing inbred mouse models different from BALB/c. The results of this work must be considered pioneering and indicative of the direction of future research.

Supporting information

S1 Table. Representation of metacyclic promastigotes in inocula.

https://doi.org/10.1371/journal.pntd.0011897.s001

(XLSX)

S2 Table. Weight changes of infected animals during the experiment.

https://doi.org/10.1371/journal.pntd.0011897.s002

(XLSX)

S3 Table. Parasite load results from various tissues measured by qPCR.

https://doi.org/10.1371/journal.pntd.0011897.s003

(XLSX)

Acknowledgments

We would like to thank prof. Padet Siriyasatien, prof. Paul Bates and doc. Jan Votýpka and prof. David Modrý for providing Leishmania isolates. Also, we would like to thank our technical support stuff Kristyna Srstkova, Lenka Krejcirikova, and Lenka Hlubinkova.

References

1. Espinosa OA, Serrano MG, Camargo EP, Teixeira MMG, Shaw JJ. An appraisal of the taxonomy and nomenclature of trypanosomatids presently classified as Leishmania and Endotrypanum. Parasitology. 2018; 145(4), 430–442.
- View Article
- Google Scholar
2. Leishmaniasis [Internet]. [cited 2023 Feb 28]. Available from: https://www.who.int/news-room/fact-sheets/detail/leishmaniasis
3. Akilov OE, Khachemoune A, Hasan T. Clinical manifestations and classification of Old World cutaneous leishmaniasis. International Journal of Dermatology. 2007, 46(2), 132–142. pmid:17269962
- View Article
- PubMed/NCBI
- Google Scholar
4. Loría-Cervera EN, Andrade-Narváez FJ. Review: Animal models for the study of leishmaniasis immunology. Rev Inst Med Trop Sao Paulo. 2014; 56(1):1–11.
- View Article
- Google Scholar
5. Sadlova J, Becvar T, Volf P. Transmission of Enigmatic Mundinia Parasites Journal of Infectious Diseases & Therapy. 2022; 10(5).
- View Article
- Google Scholar
6. Kwakye-Nuako G, Mosore MT, Boakye D, Bates PA. Description, Biology, and Medical Significance of Leishmania (Mundinia) chancei N. Sp. (Kinetoplastea: Trypanosomatidae) From Ghana and Leishmania (Mundinia) procaviensis N. Sp. (Kinetoplastea: Trypanosomatidae) From Namibia. J Parasitol. 2023; 109(1): 43–50. pmid:36848641
- View Article
- PubMed/NCBI
- Google Scholar
7. Dougall AM, Alexander B, Holt DC, Harris T, Sultan AH, Bates PA, et al. Evidence incriminating midges (Diptera: Ceratopogonidae) as potential vectors of Leishmania in Australia. Int J Parasitol. 2011; 41(5), 571–579. pmid:21251914
- View Article
- PubMed/NCBI
- Google Scholar
8. Becvar T, Vojtkova B, Siriyasatien P, Votypka J, Modry D, Jahn P, et al. Experimental transmission of Leishmania (Mundinia) parasites by biting midges (Diptera: Ceratopogonidae). PLoS Pathog. 2021; 17(6), e1009654. pmid:34115806
- View Article
- PubMed/NCBI
- Google Scholar
9. Seblova V, Sadlova J, Vojtkova B, Votypka J, Carpenter S, Bates PA, et al. The Biting Midge Culicoides sonorensis (Diptera: Ceratopogonidae) Is Capable of Developing Late Stage Infections of Leishmania enriettii. PLoS Negl Trop Dis. 2015; 9(9), e0004060. pmid:26367424
- View Article
- PubMed/NCBI
- Google Scholar
10. Chanmol W, Jariyapan N, Somboon P, Bates MD, Bates PA. Development of Leishmania orientalis in the sand fly Lutzomyia longipalpis (Diptera: Psychodidae) and the biting midge Culicoides soronensis (Diptera: Ceratopogonidae). Acta Trop. 2019; 199, 105157. pmid:31491400
- View Article
- PubMed/NCBI
- Google Scholar
11. Muniz J, Medina H. Leishmaniose tegumentar do cobaio, Leishmania enriettii n. sp. Hospital (Rio J). 1948; 33(1), 7–25.
- View Article
- Google Scholar
12. Rose K, Curtis J, Baldwin T, Mathis A, Kumar B, Sakthianandeswaren A, et al. Cutaneous leishmaniasis in red kangaroos: Isolation and characterisation of the causative organisms. Int J Parasitol. 2004; 34(6), 655–664. pmid:15111087
- View Article
- PubMed/NCBI
- Google Scholar
13. Dougall A, Shilton C, Low Choy J, Alexander B, Walton S. New reports of Australian cutaneous leishmaniasis in Northern Australian macropods. Epidemiol Infect. 2009; 45(11), 679–684. pmid:19288959
- View Article
- PubMed/NCBI
- Google Scholar
14. Grove SS. Leishmaniasis in South West Africa/Namibia to date. South African Med J. 1989;75(6):290–2. pmid:2648612
- View Article
- PubMed/NCBI
- Google Scholar
15. Kwakye-Nuako G, Mosore MT, Duplessis C, Bates MD, Puplampu N, Mensah-Attipoe I, et al. First isolation of a new species of Leishmania responsible for human cutaneous leishmaniasis in Ghana and classification in the Leishmania enriettii complex. Int J Parasitol. 2015; 45(11), 679–684.
- View Article
- Google Scholar
16. Bualert L, Charungkiattikul W, Thongsuksai P, Mungthin M, Siripattanapipong S, Khositnithikul R, et al. Case report: Autochthonous disseminated dermal and visceral leishmaniasis in an AIDS patient, Southern Thailand, caused by Leishmania siamensis. Am J Trop Med Hyg. 2012; 86(5), 821–824.
- View Article
- Google Scholar
17. Pothirat T, Tantiworawit A, Chaiwarith R, Jariyapan N, Wannasan A, Siriyasatien P, et al. First Isolation of Leishmania from Northern Thailand: Case Report, Identification as Leishmania martiniquensis and Phylogenetic Position within the Leishmania enriettii Complex. PLoS Negl Trop Dis. 2014; 8(12), e3339. pmid:25474647
- View Article
- PubMed/NCBI
- Google Scholar
18. Lobsiger L, Müller N, Schweizer T, Frey CF, Wiederkehr D, Zumkehr B, et al. An autochthonous case of cutaneous bovine leishmaniasis in Switzerland. Vet Parasitol. 2010; pmid:20153118
- View Article
- PubMed/NCBI
- Google Scholar
19. Müller N, Welle M, Lobsiger L, Stoffel MH, Boghenbor KK, Hilbe M, et al. Occurrence of Leishmania sp. in cutaneous lesions of horses in Central Europe. Vet Parasitol. 2009; 169(3–4), 408–414.
- View Article
- Google Scholar
20. Reuss SM, Dunbar MD, Calderwood Mays MB, Owen JL, Mallicote MF, Archer LL, et al. Autochthonous Leishmania siamensis in horse, Florida, USA. Emerging Infectious Diseases. 2012. 18(9), 1545–1547.
- View Article
- Google Scholar
21. Supsrisunjai C, Kootiratrakarn T, Puangpet P, Bunnag T, Chaowalit P, Wessagowit V. Case report: Disseminated autochthonous dermal leishmaniasis caused by Leishmania siamensis (PCM2 Trang) in a patient from central Thailand infected with human immunodeficiency virus. Am J Trop Med Hyg. 2017; 96(5), 1160–1163.
- View Article
- Google Scholar
22. Jariyapan N, Daroontum T, Jaiwong K, Chanmol W, Intakhan N, Sor-Suwan S, et al. Leishmania (Mundinia) orientalis n. sp. (Trypanosomatidae), a parasite from Thailand responsible for localised cutaneous leishmaniasis. Parasites and Vectors. 2018; 11, 1–9.
- View Article
- Google Scholar
23. Anugulruengkitt S, Songtaweesin WN, Thepnarong N, Tangthanapalakul A, Sitthisan M, Chatproedprai S, et al. Case Report: Simple Nodular Cutaneous Leishmaniasis Caused by Autochthonous Leishmania (Mundinia) orientalis in an 18-Month-Old Girl: The First Pediatric Case in Thailand and Literature Review. Am J Trop Med Hyg. 2023; 108(1):44–50. pmid:36410322
- View Article
- PubMed/NCBI
- Google Scholar
24. Manomat J, Leelayoova S, Bualert L, Tan-ariya P, Siripattanapipong S, Mungthin M, et al. Prevalence and risk factors associated with Leishmania infection in Trang Province, southern Thailand. PLoS Negl Trop Dis. 2017; 11(11), e0006095. pmid:29155831
- View Article
- PubMed/NCBI
- Google Scholar
25. Srivarasat S, Brownell N, Siriyasatien P, Noppakun N, Asawanonda P, Rattanakorn K, et al. Case Report: Autochthonous Disseminated Cutaneous, Mucocutaneous, and Visceral Leishmaniasis Caused by Leishmania martiniquensis in a Patient with HIV/AIDS from Northern Thailand and Literature Review. Am J Trop Med Hyg. 2022;107(6):1196–1202.
- View Article
- Google Scholar
26. Kweku MA, Odoom S, Puplampu N, Desewu K, Nuako GK, Gyan B, et al. An outbreak of suspected cutaneous leishmaniasis in Ghana: lessons learnt and preparation for future outbreaks. Global health action. 2011, 4(1), 5527. pmid:21765823
- View Article
- PubMed/NCBI
- Google Scholar
27. Chusri S, Thammapalo S, Silpapojakul K, Siriyasatien P. Animal reservoirs and potential vectors of Leishmania siamensis in southern Thailand. Southeast Asian J Trop Med Public Health. 2014; 45(1), 13.
- View Article
- Google Scholar
28. Chaves LF, Hernandez MJ, Dobson AP, Pascual M. Sources and sinks: revisiting the criteria for identifying reservoirs for American cutaneous leishmaniasis. Trends Parasitol. 2007; 23(7), 311–316. pmid:17524806
- View Article
- PubMed/NCBI
- Google Scholar
29. Smyly HJ, Young CW. The experimental transmission of leishmaniasis to animals. Proc Soc Exp Biol Med. 1924;21:354–6.
- View Article
- Google Scholar
30. Hindle E, Patton WS. Reports from the Royal Society’s Kala Azar Commission in China. No. 2.—Experiments Bearing on the Susceptibility of the Striped Hamster (Cricetulus griseus) to Leishmania of Chinese Kala Azar. Proc R Soc B Biol Sci. 1926;374–9.
- View Article
- Google Scholar
31. Young CW, Smyly HJ, Brown C. Experimental kala azar in a hamster, Cricetulus griseus M.Edw. Am J Hyg. 1926;6(2):254–75.
- View Article
- Google Scholar
32. Meleney HE. The Histopathology of Kala-Azar in the Hamster, Monkey, and Man. Am J Pathol. 1925; 1(2), 147. pmid:19969637
- View Article
- PubMed/NCBI
- Google Scholar
33. Vojtkova B, Spitzova T, Votypka J, Lestinova T, Kominkova I, Hajkova M, et al. Central Asian rodents as model animals for Leishmania major and Leishmania donovani research. Microorganisms. 2020; 8(9), 1440. pmid:32962237
- View Article
- PubMed/NCBI
- Google Scholar
34. Lun ZR, Wu MS, Chen YF, Wang JY, Zhou XN, Liao LF, et al. Visceral leishmaniasis in China: An endemic disease under Control. Clin Microbiol Rev. 2015; 28(4), 987–1004. pmid:26354822
- View Article
- PubMed/NCBI
- Google Scholar
35. Sadlova J, Vojtkova B, Lestinova T, Becvar T, Frynta D, Benallal KE, et al. Infectiousness of Asymptomatic Meriones shawi, Reservoir Host of Leishmania major. Pathogens. 2023;12(4):1–10. pmid:37111500
- View Article
- PubMed/NCBI
- Google Scholar
36. Sadlova J, Vojtkova B, Becvar T, Lestinova T, Spitzova T, Bates P, et al. Host competence of the African rodents Arvicanthis neumanni, A. niloticus and Mastomys natalensis for Leishmania donovani from Ethiopia and L. (Mundinia) sp. from Ghana. Int J Parasitol Parasites Wildl. 2020; 11, 40–45. pmid:31879594
- View Article
- PubMed/NCBI
- Google Scholar
37. Pruzinova K, Sadlova J, Myskova J, Lestinova T, Janda J, Volf P. Leishmania mortality in sand fly blood meal is not species-specific and does not result from direct effect of proteinases. Parasites and Vectors. 2018;11(1):1–9.
- View Article
- Google Scholar
38. Dostálová A, Volf P. Leishmania development in sand flies: Parasite-vector interactions overview. Parasites and Vectors. 2012, 5(1), 1–12.
- View Article
- Google Scholar
39. Sadlova J, Volf P. Peritrophic matrix of Phlebotomus duboscqi and its kinetics during Leishmania major development. Cell Tissue Res. 2009;337:313–25.
- View Article
- Google Scholar
40. Walters LL. Leishmania Differentiation in Natural and Unnatural Sand Fly Hosts. J Eukaryot Microbiol. 1993; 40(2), 196–206.
- View Article
- Google Scholar
41. Čiháková J, Volf P. Development of different Leishmania major strains in the vector sandflies Phlebotomus papatasi and P. duboscqi. Ann Trop Med Parasitol. 1997; 91(3), 267–27.
- View Article
- Google Scholar
42. Sadlova J, Bacikova D, Becvar T, Vojtkova B, England M, Shaw J, et al. Porcisia transmission by prediuresis of sand flies. Front Cell Infect Microbiol. 2022; 12:1–14.
- View Article
- Google Scholar
43. Belehu A, Turk JL. Establishment of cutaneous Leishmania enriettii infection in hamsters. Infect Immun. 1976; 13(4), 1235–1241
- View Article
- Google Scholar
44. Paranaiba LF, Pinheiro LJ, Macedo DH, Menezes-Neto A, Torrecilhas AC, Tafuri WL, et al. An overview on Leishmania (Mundinia) enriettii: Biology, immunopathology, LRV and extracellular vesicles during the host-parasite interaction. Parasitology. 2018; 145(10), 1265–1273. pmid:29223169
- View Article
- PubMed/NCBI
- Google Scholar
45. Thomaz-Soccol V. New isolation of Leishmania enriettii Muniz and Medina, 1948 in Parana State, Brazil, 50 years after the first description, and isoenzymatic polymorphism of the L. enriettii taxon. Ann Trop Med Parasitol. 1996; 90(5), 491–495.
- View Article
- Google Scholar
46. Paraense WL. The spread of Leishmania enriettii through the body of the guinea pig. Trans R Soc Trop Med Hyg. 1953; 47(6), 556–60.
- View Article
- Google Scholar
47. Becvar T, Siriyasatien P, Bates P, Volf P, Sádlová J. Development of Leishmania (Mundinia) in guinea pigs. Parasites and Vectors. 2020; 13(1), 1–6.
- View Article
- Google Scholar
48. Intakhan N, Chanmol W, Kongkaew A, Somboon P, Bates MD, Bates PA, et al. Experimental infection of Leishmania (Mundinia) martiniquensis in BALB/c mice and Syrian golden hamsters. Parasitol Res. 2020; 19, 3041–3051. pmid:32779021
- View Article
- PubMed/NCBI
- Google Scholar
49. Wilson ME, Jeronimo SMB, Pearson RD. Immunopathogenesis of infection with the visceralizing Leishmania species. Microb Pathog. 2005; 38(4), 147–160.
- View Article
- Google Scholar
50. Sadlova J, Seblova V, Votypka J, Warburg A, Volf P. Xenodiagnosis of Leishmania donovani in BALB/c mice using Phlebotomus orientalis: A new laboratory model. Parasites and Vectors. 2015; 2015; 8, 1–8. pmid:25881258
- View Article
- PubMed/NCBI
- Google Scholar
51. Garin YJF, Sulahian A, Méneceur P, Pratlong F, Prina E, Gangneux JP, et al. Experimental pathogenicity of a presumed monoxenous trypanosomatid isolated from humans in a murine model. J Eukaryot Microbiol. 2001; 48(2), 170–176. pmid:12095104
- View Article
- PubMed/NCBI
- Google Scholar
52. Somboonpoonpol N. Parasite burden, distribution and imunopathology of Leishmania martiniquensis—infected BALB/c mice in different routes and time points. Chulalongkorn University; 2016.
- View Article
- Google Scholar
53. Falú MA, Fernanda M, Bustos G, María C, Ramoneda P, Raspi EM De, et al. Susceptibility of different mouse strains to Leishmania amazonensis infection. 2007; 15(5):334–33.
- View Article
- Google Scholar
54. Sriwongpan P, Nedsuwan S, Manomat J, Charoensakulchai S, Lacharojana K, Sankwan J, et al. Prevalence and associated risk factors of leishmania infection among immunocompetent hosts, a community based study in Chiang Rrai, Tthailand. PLoS Negl Trop Dis. 2021; 15(7), e0009545.
- View Article
- Google Scholar
55. Vojtkova B, Frynta D, Spitzova T, Lestinova T, Votypka J, Volf P, et al. Repeated Sand Fly Bites of Infected BALB/c Mice Enhance the Development of Leishmania lesions. Front Trop Dis. 2021; 2, 745104.
- View Article
- Google Scholar
56. Valverde JG, Paun A, Inbar E, Romano A, Lewis M, Ghosh K, et al. Increased transmissibility of Leishmania donovani from the mammalian host to vector sand flies after multiple exposures to sand fly bites. J Infect Dis. 2017;215(8):1285–93.
- View Article
- Google Scholar
57. Sadlova J, Vojtkova B, Hrncirova K, Lestinova T, Spitzova T, Becvar T, et al. Host competence of African rodents Arvicanthis neumanni, A. niloticus and Mastomys natalensis for Leishmania major. Int J Parasitol Parasites Wildl. 2019; 8, 118–126.
- View Article
- Google Scholar

[ref1] 1. Espinosa OA, Serrano MG, Camargo EP, Teixeira MMG, Shaw JJ. An appraisal of the taxonomy and nomenclature of trypanosomatids presently classified as Leishmania and Endotrypanum. Parasitology. 2018; 145(4), 430–442.
View Article
Google Scholar

[2] View Article

[3] Google Scholar

[ref2] 2. Leishmaniasis [Internet]. [cited 2023 Feb 28]. Available from: https://www.who.int/news-room/fact-sheets/detail/leishmaniasis

[ref3] 3. Akilov OE, Khachemoune A, Hasan T. Clinical manifestations and classification of Old World cutaneous leishmaniasis. International Journal of Dermatology. 2007, 46(2), 132–142. pmid:17269962
View Article
PubMed/NCBI
Google Scholar

[6] View Article

[7] PubMed/NCBI

[8] Google Scholar

[ref4] 4. Loría-Cervera EN, Andrade-Narváez FJ. Review: Animal models for the study of leishmaniasis immunology. Rev Inst Med Trop Sao Paulo. 2014; 56(1):1–11.
View Article
Google Scholar

[10] View Article

[11] Google Scholar

[ref5] 5. Sadlova J, Becvar T, Volf P. Transmission of Enigmatic Mundinia Parasites Journal of Infectious Diseases & Therapy. 2022; 10(5).
View Article
Google Scholar

[13] View Article

[14] Google Scholar

[ref6] 6. Kwakye-Nuako G, Mosore MT, Boakye D, Bates PA. Description, Biology, and Medical Significance of Leishmania (Mundinia) chancei N. Sp. (Kinetoplastea: Trypanosomatidae) From Ghana and Leishmania (Mundinia) procaviensis N. Sp. (Kinetoplastea: Trypanosomatidae) From Namibia. J Parasitol. 2023; 109(1): 43–50. pmid:36848641
View Article
PubMed/NCBI
Google Scholar

[16] View Article

[17] PubMed/NCBI

[18] Google Scholar

[ref7] 7. Dougall AM, Alexander B, Holt DC, Harris T, Sultan AH, Bates PA, et al. Evidence incriminating midges (Diptera: Ceratopogonidae) as potential vectors of Leishmania in Australia. Int J Parasitol. 2011; 41(5), 571–579. pmid:21251914
View Article
PubMed/NCBI
Google Scholar

[20] View Article

[21] PubMed/NCBI

[22] Google Scholar

[ref8] 8. Becvar T, Vojtkova B, Siriyasatien P, Votypka J, Modry D, Jahn P, et al. Experimental transmission of Leishmania (Mundinia) parasites by biting midges (Diptera: Ceratopogonidae). PLoS Pathog. 2021; 17(6), e1009654. pmid:34115806
View Article
PubMed/NCBI
Google Scholar

[24] View Article

[25] PubMed/NCBI

[26] Google Scholar

[ref9] 9. Seblova V, Sadlova J, Vojtkova B, Votypka J, Carpenter S, Bates PA, et al. The Biting Midge Culicoides sonorensis (Diptera: Ceratopogonidae) Is Capable of Developing Late Stage Infections of Leishmania enriettii. PLoS Negl Trop Dis. 2015; 9(9), e0004060. pmid:26367424
View Article
PubMed/NCBI
Google Scholar

[28] View Article

[29] PubMed/NCBI

[30] Google Scholar

[ref10] 10. Chanmol W, Jariyapan N, Somboon P, Bates MD, Bates PA. Development of Leishmania orientalis in the sand fly Lutzomyia longipalpis (Diptera: Psychodidae) and the biting midge Culicoides soronensis (Diptera: Ceratopogonidae). Acta Trop. 2019; 199, 105157. pmid:31491400
View Article
PubMed/NCBI
Google Scholar

[32] View Article

[33] PubMed/NCBI

[34] Google Scholar

[ref11] 11. Muniz J, Medina H. Leishmaniose tegumentar do cobaio, Leishmania enriettii n. sp. Hospital (Rio J). 1948; 33(1), 7–25.
View Article
Google Scholar

[36] View Article

[37] Google Scholar

[ref12] 12. Rose K, Curtis J, Baldwin T, Mathis A, Kumar B, Sakthianandeswaren A, et al. Cutaneous leishmaniasis in red kangaroos: Isolation and characterisation of the causative organisms. Int J Parasitol. 2004; 34(6), 655–664. pmid:15111087
View Article
PubMed/NCBI
Google Scholar

[39] View Article

[40] PubMed/NCBI

[41] Google Scholar

[ref13] 13. Dougall A, Shilton C, Low Choy J, Alexander B, Walton S. New reports of Australian cutaneous leishmaniasis in Northern Australian macropods. Epidemiol Infect. 2009; 45(11), 679–684. pmid:19288959
View Article
PubMed/NCBI
Google Scholar

[43] View Article

[44] PubMed/NCBI

[45] Google Scholar

[ref14] 14. Grove SS. Leishmaniasis in South West Africa/Namibia to date. South African Med J. 1989;75(6):290–2. pmid:2648612
View Article
PubMed/NCBI
Google Scholar

[47] View Article

[48] PubMed/NCBI

[49] Google Scholar

[ref15] 15. Kwakye-Nuako G, Mosore MT, Duplessis C, Bates MD, Puplampu N, Mensah-Attipoe I, et al. First isolation of a new species of Leishmania responsible for human cutaneous leishmaniasis in Ghana and classification in the Leishmania enriettii complex. Int J Parasitol. 2015; 45(11), 679–684.
View Article
Google Scholar

[51] View Article

[52] Google Scholar

[ref16] 16. Bualert L, Charungkiattikul W, Thongsuksai P, Mungthin M, Siripattanapipong S, Khositnithikul R, et al. Case report: Autochthonous disseminated dermal and visceral leishmaniasis in an AIDS patient, Southern Thailand, caused by Leishmania siamensis. Am J Trop Med Hyg. 2012; 86(5), 821–824.
View Article
Google Scholar

[54] View Article

[55] Google Scholar

[ref17] 17. Pothirat T, Tantiworawit A, Chaiwarith R, Jariyapan N, Wannasan A, Siriyasatien P, et al. First Isolation of Leishmania from Northern Thailand: Case Report, Identification as Leishmania martiniquensis and Phylogenetic Position within the Leishmania enriettii Complex. PLoS Negl Trop Dis. 2014; 8(12), e3339. pmid:25474647
View Article
PubMed/NCBI
Google Scholar

[57] View Article

[58] PubMed/NCBI

[59] Google Scholar

[ref18] 18. Lobsiger L, Müller N, Schweizer T, Frey CF, Wiederkehr D, Zumkehr B, et al. An autochthonous case of cutaneous bovine leishmaniasis in Switzerland. Vet Parasitol. 2010; pmid:20153118
View Article
PubMed/NCBI
Google Scholar

[61] View Article

[62] PubMed/NCBI

[63] Google Scholar

[ref19] 19. Müller N, Welle M, Lobsiger L, Stoffel MH, Boghenbor KK, Hilbe M, et al. Occurrence of Leishmania sp. in cutaneous lesions of horses in Central Europe. Vet Parasitol. 2009; 169(3–4), 408–414.
View Article
Google Scholar

[65] View Article

[66] Google Scholar

[ref20] 20. Reuss SM, Dunbar MD, Calderwood Mays MB, Owen JL, Mallicote MF, Archer LL, et al. Autochthonous Leishmania siamensis in horse, Florida, USA. Emerging Infectious Diseases. 2012. 18(9), 1545–1547.
View Article
Google Scholar

[68] View Article

[69] Google Scholar

[ref21] 21. Supsrisunjai C, Kootiratrakarn T, Puangpet P, Bunnag T, Chaowalit P, Wessagowit V. Case report: Disseminated autochthonous dermal leishmaniasis caused by Leishmania siamensis (PCM2 Trang) in a patient from central Thailand infected with human immunodeficiency virus. Am J Trop Med Hyg. 2017; 96(5), 1160–1163.
View Article
Google Scholar

[71] View Article

[72] Google Scholar

[ref22] 22. Jariyapan N, Daroontum T, Jaiwong K, Chanmol W, Intakhan N, Sor-Suwan S, et al. Leishmania (Mundinia) orientalis n. sp. (Trypanosomatidae), a parasite from Thailand responsible for localised cutaneous leishmaniasis. Parasites and Vectors. 2018; 11, 1–9.
View Article
Google Scholar

[74] View Article

[75] Google Scholar

[ref23] 23. Anugulruengkitt S, Songtaweesin WN, Thepnarong N, Tangthanapalakul A, Sitthisan M, Chatproedprai S, et al. Case Report: Simple Nodular Cutaneous Leishmaniasis Caused by Autochthonous Leishmania (Mundinia) orientalis in an 18-Month-Old Girl: The First Pediatric Case in Thailand and Literature Review. Am J Trop Med Hyg. 2023; 108(1):44–50. pmid:36410322
View Article
PubMed/NCBI
Google Scholar

[77] View Article

[78] PubMed/NCBI

[79] Google Scholar

[ref24] 24. Manomat J, Leelayoova S, Bualert L, Tan-ariya P, Siripattanapipong S, Mungthin M, et al. Prevalence and risk factors associated with Leishmania infection in Trang Province, southern Thailand. PLoS Negl Trop Dis. 2017; 11(11), e0006095. pmid:29155831
View Article
PubMed/NCBI
Google Scholar

[81] View Article

[82] PubMed/NCBI

[83] Google Scholar

[ref25] 25. Srivarasat S, Brownell N, Siriyasatien P, Noppakun N, Asawanonda P, Rattanakorn K, et al. Case Report: Autochthonous Disseminated Cutaneous, Mucocutaneous, and Visceral Leishmaniasis Caused by Leishmania martiniquensis in a Patient with HIV/AIDS from Northern Thailand and Literature Review. Am J Trop Med Hyg. 2022;107(6):1196–1202.
View Article
Google Scholar

[85] View Article

[86] Google Scholar

[ref26] 26. Kweku MA, Odoom S, Puplampu N, Desewu K, Nuako GK, Gyan B, et al. An outbreak of suspected cutaneous leishmaniasis in Ghana: lessons learnt and preparation for future outbreaks. Global health action. 2011, 4(1), 5527. pmid:21765823
View Article
PubMed/NCBI
Google Scholar

[88] View Article

[89] PubMed/NCBI

[90] Google Scholar

[ref27] 27. Chusri S, Thammapalo S, Silpapojakul K, Siriyasatien P. Animal reservoirs and potential vectors of Leishmania siamensis in southern Thailand. Southeast Asian J Trop Med Public Health. 2014; 45(1), 13.
View Article
Google Scholar

[92] View Article

[93] Google Scholar

[ref28] 28. Chaves LF, Hernandez MJ, Dobson AP, Pascual M. Sources and sinks: revisiting the criteria for identifying reservoirs for American cutaneous leishmaniasis. Trends Parasitol. 2007; 23(7), 311–316. pmid:17524806
View Article
PubMed/NCBI
Google Scholar

[95] View Article

[96] PubMed/NCBI

[97] Google Scholar

[ref29] 29. Smyly HJ, Young CW. The experimental transmission of leishmaniasis to animals. Proc Soc Exp Biol Med. 1924;21:354–6.
View Article
Google Scholar

[99] View Article

[100] Google Scholar

[ref30] 30. Hindle E, Patton WS. Reports from the Royal Society’s Kala Azar Commission in China. No. 2.—Experiments Bearing on the Susceptibility of the Striped Hamster (Cricetulus griseus) to Leishmania of Chinese Kala Azar. Proc R Soc B Biol Sci. 1926;374–9.
View Article
Google Scholar

[102] View Article

[103] Google Scholar

[ref31] 31. Young CW, Smyly HJ, Brown C. Experimental kala azar in a hamster, Cricetulus griseus M.Edw. Am J Hyg. 1926;6(2):254–75.
View Article
Google Scholar

[105] View Article

[106] Google Scholar

[ref32] 32. Meleney HE. The Histopathology of Kala-Azar in the Hamster, Monkey, and Man. Am J Pathol. 1925; 1(2), 147. pmid:19969637
View Article
PubMed/NCBI
Google Scholar

[108] View Article

[109] PubMed/NCBI

[110] Google Scholar

[ref33] 33. Vojtkova B, Spitzova T, Votypka J, Lestinova T, Kominkova I, Hajkova M, et al. Central Asian rodents as model animals for Leishmania major and Leishmania donovani research. Microorganisms. 2020; 8(9), 1440. pmid:32962237
View Article
PubMed/NCBI
Google Scholar

[112] View Article

[113] PubMed/NCBI

[114] Google Scholar

[ref34] 34. Lun ZR, Wu MS, Chen YF, Wang JY, Zhou XN, Liao LF, et al. Visceral leishmaniasis in China: An endemic disease under Control. Clin Microbiol Rev. 2015; 28(4), 987–1004. pmid:26354822
View Article
PubMed/NCBI
Google Scholar

[116] View Article

[117] PubMed/NCBI

[118] Google Scholar

[ref35] 35. Sadlova J, Vojtkova B, Lestinova T, Becvar T, Frynta D, Benallal KE, et al. Infectiousness of Asymptomatic Meriones shawi, Reservoir Host of Leishmania major. Pathogens. 2023;12(4):1–10. pmid:37111500
View Article
PubMed/NCBI
Google Scholar

[120] View Article

[121] PubMed/NCBI

[122] Google Scholar

[ref36] 36. Sadlova J, Vojtkova B, Becvar T, Lestinova T, Spitzova T, Bates P, et al. Host competence of the African rodents Arvicanthis neumanni, A. niloticus and Mastomys natalensis for Leishmania donovani from Ethiopia and L. (Mundinia) sp. from Ghana. Int J Parasitol Parasites Wildl. 2020; 11, 40–45. pmid:31879594
View Article
PubMed/NCBI
Google Scholar

[124] View Article

[125] PubMed/NCBI

[126] Google Scholar

[ref37] 37. Pruzinova K, Sadlova J, Myskova J, Lestinova T, Janda J, Volf P. Leishmania mortality in sand fly blood meal is not species-specific and does not result from direct effect of proteinases. Parasites and Vectors. 2018;11(1):1–9.
View Article
Google Scholar

[128] View Article

[129] Google Scholar

[ref38] 38. Dostálová A, Volf P. Leishmania development in sand flies: Parasite-vector interactions overview. Parasites and Vectors. 2012, 5(1), 1–12.
View Article
Google Scholar

[131] View Article

[132] Google Scholar

[ref39] 39. Sadlova J, Volf P. Peritrophic matrix of Phlebotomus duboscqi and its kinetics during Leishmania major development. Cell Tissue Res. 2009;337:313–25.
View Article
Google Scholar

[134] View Article

[135] Google Scholar

[ref40] 40. Walters LL. Leishmania Differentiation in Natural and Unnatural Sand Fly Hosts. J Eukaryot Microbiol. 1993; 40(2), 196–206.
View Article
Google Scholar

[137] View Article

[138] Google Scholar

[ref41] 41. Čiháková J, Volf P. Development of different Leishmania major strains in the vector sandflies Phlebotomus papatasi and P. duboscqi. Ann Trop Med Parasitol. 1997; 91(3), 267–27.
View Article
Google Scholar

[140] View Article

[141] Google Scholar

[ref42] 42. Sadlova J, Bacikova D, Becvar T, Vojtkova B, England M, Shaw J, et al. Porcisia transmission by prediuresis of sand flies. Front Cell Infect Microbiol. 2022; 12:1–14.
View Article
Google Scholar

[143] View Article

[144] Google Scholar

[ref43] 43. Belehu A, Turk JL. Establishment of cutaneous Leishmania enriettii infection in hamsters. Infect Immun. 1976; 13(4), 1235–1241
View Article
Google Scholar

[146] View Article

[147] Google Scholar

[ref44] 44. Paranaiba LF, Pinheiro LJ, Macedo DH, Menezes-Neto A, Torrecilhas AC, Tafuri WL, et al. An overview on Leishmania (Mundinia) enriettii: Biology, immunopathology, LRV and extracellular vesicles during the host-parasite interaction. Parasitology. 2018; 145(10), 1265–1273. pmid:29223169
View Article
PubMed/NCBI
Google Scholar

[149] View Article

[150] PubMed/NCBI

[151] Google Scholar

[ref45] 45. Thomaz-Soccol V. New isolation of Leishmania enriettii Muniz and Medina, 1948 in Parana State, Brazil, 50 years after the first description, and isoenzymatic polymorphism of the L. enriettii taxon. Ann Trop Med Parasitol. 1996; 90(5), 491–495.
View Article
Google Scholar

[153] View Article

[154] Google Scholar

[ref46] 46. Paraense WL. The spread of Leishmania enriettii through the body of the guinea pig. Trans R Soc Trop Med Hyg. 1953; 47(6), 556–60.
View Article
Google Scholar

[156] View Article

[157] Google Scholar

[ref47] 47. Becvar T, Siriyasatien P, Bates P, Volf P, Sádlová J. Development of Leishmania (Mundinia) in guinea pigs. Parasites and Vectors. 2020; 13(1), 1–6.
View Article
Google Scholar

[159] View Article

[160] Google Scholar

[ref48] 48. Intakhan N, Chanmol W, Kongkaew A, Somboon P, Bates MD, Bates PA, et al. Experimental infection of Leishmania (Mundinia) martiniquensis in BALB/c mice and Syrian golden hamsters. Parasitol Res. 2020; 19, 3041–3051. pmid:32779021
View Article
PubMed/NCBI
Google Scholar

[162] View Article

[163] PubMed/NCBI

[164] Google Scholar

[ref49] 49. Wilson ME, Jeronimo SMB, Pearson RD. Immunopathogenesis of infection with the visceralizing Leishmania species. Microb Pathog. 2005; 38(4), 147–160.
View Article
Google Scholar

[166] View Article

[167] Google Scholar

[ref50] 50. Sadlova J, Seblova V, Votypka J, Warburg A, Volf P. Xenodiagnosis of Leishmania donovani in BALB/c mice using Phlebotomus orientalis: A new laboratory model. Parasites and Vectors. 2015; 2015; 8, 1–8. pmid:25881258
View Article
PubMed/NCBI
Google Scholar

[169] View Article

[170] PubMed/NCBI

[171] Google Scholar

[ref51] 51. Garin YJF, Sulahian A, Méneceur P, Pratlong F, Prina E, Gangneux JP, et al. Experimental pathogenicity of a presumed monoxenous trypanosomatid isolated from humans in a murine model. J Eukaryot Microbiol. 2001; 48(2), 170–176. pmid:12095104
View Article
PubMed/NCBI
Google Scholar

[173] View Article

[174] PubMed/NCBI

[175] Google Scholar

[ref52] 52. Somboonpoonpol N. Parasite burden, distribution and imunopathology of Leishmania martiniquensis—infected BALB/c mice in different routes and time points. Chulalongkorn University; 2016.
View Article
Google Scholar

[177] View Article

[178] Google Scholar

[ref53] 53. Falú MA, Fernanda M, Bustos G, María C, Ramoneda P, Raspi EM De, et al. Susceptibility of different mouse strains to Leishmania amazonensis infection. 2007; 15(5):334–33.
View Article
Google Scholar

[180] View Article

[181] Google Scholar

[ref54] 54. Sriwongpan P, Nedsuwan S, Manomat J, Charoensakulchai S, Lacharojana K, Sankwan J, et al. Prevalence and associated risk factors of leishmania infection among immunocompetent hosts, a community based study in Chiang Rrai, Tthailand. PLoS Negl Trop Dis. 2021; 15(7), e0009545.
View Article
Google Scholar

[183] View Article

[184] Google Scholar

[ref55] 55. Vojtkova B, Frynta D, Spitzova T, Lestinova T, Votypka J, Volf P, et al. Repeated Sand Fly Bites of Infected BALB/c Mice Enhance the Development of Leishmania lesions. Front Trop Dis. 2021; 2, 745104.
View Article
Google Scholar

[186] View Article

[187] Google Scholar

[ref56] 56. Valverde JG, Paun A, Inbar E, Romano A, Lewis M, Ghosh K, et al. Increased transmissibility of Leishmania donovani from the mammalian host to vector sand flies after multiple exposures to sand fly bites. J Infect Dis. 2017;215(8):1285–93.
View Article
Google Scholar

[189] View Article

[190] Google Scholar

[ref57] 57. Sadlova J, Vojtkova B, Hrncirova K, Lestinova T, Spitzova T, Becvar T, et al. Host competence of African rodents Arvicanthis neumanni, A. niloticus and Mastomys natalensis for Leishmania major. Int J Parasitol Parasites Wildl. 2019; 8, 118–126.
View Article
Google Scholar

[192] View Article

[193] Google Scholar

Figures

Abstract

Author summary

1. Introduction

2. Materials and methods

2.1. Ethics statement

2.2. Parasites and rodents

2.3. Experimental infections and xenodiagnoses of rodents

2.4. Morphometry of parasites

2.5. DNA isolation and nested PCR of sand fly and rodent samples

2.6. Quantitative PCR

2.7. Statistical analysis

3. Results

3.1. Development of Mundinia in BALB/c mice

3.2. Development of Mundinia in Chinese hamsters (Cricetulus griseus)

3.3. Development of Mundinia in steppe lemmings (Lagurus lagurus)

4. Discussion

Supporting information

S1 Table. Representation of metacyclic promastigotes in inocula.

S2 Table. Weight changes of infected animals during the experiment.

S3 Table. Parasite load results from various tissues measured by qPCR.

Acknowledgments

References