https://orcid.org/0000-0002-7518-476X
Abstract
Equids may be infected by zoonotic Leishmania spp., including Leishmania infantum, in regions where canine leishmaniasis (CanL) is endemic, and Leishmania martiniquensis, which has been reported in horses from Central Europe. This study was designed to evaluate the occurrence of both Leishmania spp. among equids living in CanL endemic areas of Italy, as well as to identify dipteran vectors from the same habitats. From March to October 2023, blood, serum and tissue samples from skin lesions were collected from equids (n = 98; n = 56 donkeys and n = 42 horses) living in Italy, as well as sand flies and biting midges. Blood samples (n = 98) and skin lesions (n = 56) were tested for Leishmania spp. by conventional and real time PCRs and sera were tested by immunofluorescence antibody tests (IFAT) for both L. infantum and L. martiniquensis. Insects were morphologically identified, and female specimens (n = 268 sand flies, n = 7 biting midges) analyzed for Leishmania DNA, as well as engorged sand flies (n = 16) for blood-meal detection. Two animals with skin lesions (i.e., one donkey and one horse) scored positive for Leishmania spp. DNA, and 19 animals (i.e., 19.4%; n = 13 donkeys and n = 6 horses) were seropositive for L. infantum, with five of them also for L. martiniquensis. Most seropositive animals had no dermatological lesions (i.e., 68.4%) while both animals molecularly positive for Leishmania spp. scored seronegative. Of the 356 sand flies collected, 12 females (i.e., n = 8 Sergentomyia minuta; n = 3 Phlebotomus perniciosus, n = 1 Phlebotomus perfiliewi) were positive for Leishmania spp. DNA, and one out of seven biting midges collected was DNA-positive for L. infantum. Moreover, engorged sand flies scored positive for human and equine DNA. Data suggest that equids living in CanL endemic areas are exposed to Leishmania spp., but their role in the circulation of the parasite needs further investigations.
Author summary
Horses and donkeys seem to be infected by Leishmania infantum, causative agent of canine leishmaniasis (CanL), which is one of the most important zoonotic vector-borne disease. Moreover, Leishmania martiniquensis, causing visceral and mucocutaneus leishmaniasis in humans, was reported in horses from central Europe. Therefore, we designed this study to investigate the circulation of both Leishmania zoonotic species among equids living in CanL endemic areas and to access the presence of dipteran vectors from the same habitats. Horses and donkeys (n = 98) were sampled and analyzed for Leishmania spp., as well as sand flies and biting midges. Nineteen equids (19.4%) scored seropositive by immunofluorescence antibody test for L. infantum and one donkey and one horse, presenting skin lesions, scored molecularly positive for Leishmania spp. Twelve sand flies and one biting midge were positive for Leishmania spp. DNA. Overall, data herein presented suggest that equids living in CanL endemic areas are exposed to Leishmania spp., potentially contributing to the circulation of the parasite.
Figures
Citation: Carbonara M, Mendoza-Roldan JA, Bezerra-Santos MA, de Abreu Teles PP, Lia RP, Locantore F, et al. (2024) Leishmania spp. in equids and their potential vectors in endemic areas of canine leishmaniasis. PLoS Negl Trop Dis 18(7): e0012290. https://doi.org/10.1371/journal.pntd.0012290
Editor: Dhafer Laouini, Insitut Pasteur de Tunis, TUNISIA
Received: March 25, 2024; Accepted: June 11, 2024; Published: July 18, 2024
Copyright: © 2024 Carbonara 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 Leishmania sequences generated in this study were deposited in GenBank database (i.e., Leishmania sp. from horse, PP486329; Leishmania infantum from Sergentomyia minuta, PP488754; L. infantum from Culicoides imicola, PP505437; Leishmania tarentolae from S. minuta, PP505438).
Funding: D.O. and J.A.M.R. were partially supported by EU funding within the NextGeneration EU-MUR PNRR Extended Partnership initiative on Emerging Infectious Diseases (Project no. PE00000007, INF-ACT). 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.
Introduction
The leishmaniases include a diverse group of protozoan vector-borne diseases, with around 30 species of Leishmania infecting animals and humans [1, 2]. Among them, Leishmania infantum is an important species of veterinary and medical relevance, widely distributed in the Mediterranean basin, the Middle East, western Asia and Brazil [3, 4]. This protozoan primarily infects dogs but also a large variety of domestic and wild mammals [4–6], including horses [7]. Indeed, after the first description of Leishmania spp. in a horse from Argentina, in the beginning of the 20th century [8], several cases of equine leishmaniasis (EL) have been reported worldwide [9]. Based on results of serological tests, L. infantum-prevalence ranging from 0.3% to 36% were detected in healthy equids living in countries where canine leishmaniasis (CanL) is endemic, such as Portugal, Italy and Greece [10–13]. In addition, a few cases of cutaneous leishmaniasis due to L. infantum have been described in horses from Spain, Portugal and Italy [14–16] suggesting that, in areas with prevalent infections in dogs, other animal species may be exposed and/or infected by these parasites [17–19]. Moreover, autochthonous cases of Leishmania (Mundinia) martiniquensis, previously known as ‘Leishmania siamensis’, were molecularly detected from skin lesions of horses living in the United States of America [20], Switzerland and Germany [21], where CanL by L. infantum is not endemic [2]. Importantly, this Mundinia species was isolated from human patients in south-east Asia [22,23], being a zoonotic agent of visceral and mucocutaneus leishmaniasis in Thailand [24]. Differently from L. infantum, many knowledge gaps persist regarding the biology of L. martiniquensis and their vectors. Culicoides peregrinus specimens naturally infected by L. martiniquensis were collected in southern Thailand [25], along with laboratory experimental studies demonstrating successful transmission of Mundinia species by Culicoides sonorensis [26], hence suggesting that biting midges are responsible for L. martiniquensis transmission in Europe.
Descriptive clinical data about EL are scant, being the infections usually subclinical or characterized by local inflammatory response, with single or multiple papules or ulcerated nodules localized around the eyes, in muzzle, neck, pinnae, scrotum and legs, where biting insects commonly feed [7,9,27]. The diagnosis of EL relies on microscopical observation of amastigotes, usually coupled with the molecular identification of Leishmania at the species level [9]. On the other hand, immunofluorescence antibody test (IFAT) and enzyme-linked immunosorbent assay (ELISA) have been employed to detect Leishmania exposure in equids [9].
Given all the above, this study aimed to assess the circulation of L. infantum and L. martiniquensis among equids living in geographical areas of Italy, endemic for human and CanL, as well as to identify potential dipteran vectors from the same habitats.
Methods
Ethics statement
Animals were handled with regard to their well-being and the protocol of this study was approved by the ethical committee of the Department of Veterinary Medicine, University of Bari, Italy (Prot. Uniba 2/24).
Study areas and equid sample collection
From March to October 2023, blood and serum were sampled from n = 98 equids (i.e., n = 56 donkeys and n = 42 horses) living in endemic areas for CanL [28], specifically in 10 sites from Apulia, four sites from Basilicata, one from Sicily and one from Veneto regions (Fig 1). Of the 16 sampling sites, 13 were private stables hosting 2–9 animals each, and three were large breeding herds (up to 90 animals each) (Table 1). In addition, when animals presented dermatological signs, biopsies (Fig 2A) and at least three cytological slides were taken from skin lesions (Fig 2B). At the enrollment, animal data (i.e., species, sex, microchip code) along with a brief skin lesion description were recorded in individual files. The location of equid sampling was geo-referenced using a geographical information system (QGIS software, Buenos Aires version).
Map of study areas (colored green) indicated by provinces, showing the distribution of equids sampled (red circles). Map prepared using QGIS software—Buenos Aires version (link source of the shapefile: https://www.eea.europa.eu/data-and-maps/data/eea-reference-grids-2/gis-files/italy-shapefile).
A) Donkey presenting two alopecic lesions with epithelial discontinuity. B) Apposition cytology made on the skin lesion (*), with an intense mixed inflammatory infiltrate composed predominantly by neutrophils, activated macrophages phagocytosing neutrophils and apoptotic bodies, and eosinophils. Absence of amastigote forms of Leishmania spp. (Diff Quik stain, 100X).
Serological testing
Serum samples were tested to assess the exposure to L. infantum and L. martiniquensis. An IFAT for the detection of IgG anti-L. infantum was performed adapting the procedures previously described for dogs [29]. Promastigotes of L. infantum zymodeme MON-1 were used as antigens, while rabbit-anti-horse-IgG FITC (Sigma-Aldrich Chemical, Darmstadt, Germany) diluted at 1:50 was used as conjugate. Each horse serum sample was centrifugated at 1200 rpm for 15 min before starting the IFAT procedure, to reduce the unspecific fluorescence associated with physiological jaundice. To detect the antibodies against L. martiniquensis, the IFAT was performed using promastigotes of L. martiniquensis (strain MHOM/TH/2011/CU1) as antigens, following the same procedure for L. infantum IFAT, except for the use of goat-anti-horse-IgG FITC (Thermo Fisher Scientific, Rockford, USA) diluted 1:1000. For both IFAT tests, serum samples from a horse positive for L. infantum by molecular analyses (i.e., cPCR), and a healthy horse negative for L. infantum, were used as positive and negative controls, respectively. Samples were scored as positive when they produced a clear cytoplasmic and membrane fluorescence of promastigotes from a dilution of 1:80, while those that produced promastigote fluorescence at 1:40 were considered as exposed. Positive sera were titrated by serial dilutions until negative results were obtained [30].
Entomological sampling
Sand flies and biting midges were collected from September to October 2023 in two sites (Sites A and B) where Leishmania-seropositive equids were found. Site A (Crispiano, Apulia, 40°36′N 17°14′E) was a large farm (i.e., about 80 donkeys and one dog) and had a typical Mediterranean environment characterized by olive trees, the presence of “muretti a secco” (stone walls) where sand flies thrive (Fig 3A); while Site B (Matera, Basilicata, 40°40′N 16°36′E) was a small farm, close to the urban center, with about 7 horses, a donkey and one dog (Fig 3B). Insects were collected on weekly or bi-weekly basis, depending on weather conditions and the availability of the owners, setting from 5:00 p.m. to 8:00 a.m. two CDC light traps with dry ice per site, both outdoor (Fig 3C) and indoor (Fig 3D). Collections were carried out until the total disappearance/absence of insects (i.e., two consecutive negative captures), as previously described [31]. After separating sand flies and biting midges under a stereomicroscope, alive sand fly females were dissected in a drop of saline solution and the gut was observed under an optical microscope to determine the presence of flagellates [32].
A) Large farm with the typical Mediterranean environment (Crispiano, Apulia). B) Small stable, close to the city center (Matera, Basilicata) provided with stalls for equids. C) “Muretto a secco” where the CDC light traps were placed in the Site A. D) Placement of the CDC light traps at the entrance of the stable, Site B.
Insect identification and host blood meal detection
Specimens were stored in individual vials containing 70% ethanol and then morphologically identified using taxonomic keys. Heads and last segments of sand flies were removed using sterile tips and mounted on a glass slide using Hoyer’s medium; biting midges were morphologically evaluated based on their wing patterns, head palps and antennae [33–39]. Genomic DNA (gDNA) was extracted from the thorax and abdomen of sand flies (n = 268 females and n = 5 males) using the GenUP DNA Kit (Biotechrabbit, Berlin, Germany), following the producer’s recommendations. Similarly, the head-thorax of biting midge female specimens (n = 7, single sex recorded) were separated from the abdomen using sterile tips and gDNA was extracted using the DNeasy Blood and Tissue Kit (Qiagen GmbH, Hilden, Germany). When sand fly morphological identification was not possible (n = 25, n = 20 females and n = 5 males) due to the absence of morphological relevant segments, molecular identification was carried out by conventional PCR (cPCR), amplifying the mitochondrial DNA fragment encompassing cytb and nd1 regions (∼500 bp) using the primers PhleF/R, as previously described [40]. Finally, engorged female sand flies (n = 16) were tested for blood-meal by cPCR using the primers cytB1-F/B2-R, targeting the vertebrate host mitochondrial cytb (350 bp) [41] (Tables 2 and 3). Amplified PCR products were visualized by gel-electrophoresis in 2% agarose gel containing GelRed nucleic acid gel stain (VWR International PBI, Milan, Italy) and viewed on a GelLogic 100 gel documentation system (Kodak, New York, USA). All the positive cPCR products were purified and sequenced in forward direction using the same primers, employing the Big Dye Terminator v.3.1 chemistry in a 3130 Genetic analyzer (Applied Biosystems, Foster city, California, USA) in an automated sequencer (ABI-PRISM 377). Nucleotide sequences were edited, aligned, and analyzed using the Geneious platform version 9.0 (Biomatters Ltd., Auckland, New Zealand) [42], and compared with available sequences in the GenBank database, using the Basic Local Alignment Search Tool (BLAST; http://blast.ncbi.nlm.nih.gov/Blast.cgi) for species identification.
n: number of specimens collected.
Molecular detection of Leishmania
Genomic DNA was extracted from skin lesions and whole blood (200 μl) using a commercial kit (QIAampDNA Blood Tissue, Qiagen, Hilden, Germany), according to the manufacturer’s instructions. Skin lesion (n = 56) and whole blood (n = 98) DNA samples, as well as female insect DNA (n = 268 sand flies; n = 7 biting midges), were tested by real time PCR (qPCR) for the detection of a L. infantum kDNA minicircle fragment (120 bp), using primers, probes and protocol described elsewhere [43]. In addition, given the sympatric occurrence of L. infantum and Leishmania tarentolae in the areas investigated [31], a duplex real-time PCR (dqPCR) for detection of both Leishmania spp. DNA was performed as previously described [44]. Finally, three cPCRs targeting ITS1 (L5.8S/LITSR, ~ 300 bp), 18S rRNA (18SN1F/18SN1R, ~ 290 bp; 18SN2F/18SN2R, ~ 190 bp), and HSP 70 genes (F25/R617, ~ 500 bp) were performed to evaluate the presence of other Leishmania species DNA, as previously reported [45–47]. The cPCR protocol by [47] was slightly modified as follows: 95°C for 10 min initial denaturation, followed by 35 cycles of 94°C for 30 sec, 62°C for 30 sec, and 72°C for 30 sec, and then 72°C for 7 min for final elongation. Negative (i.e., male P. perniciosus specimen) and positive controls (i.e., L. infantum, L. tarentolae) were included in all PCR runs. Amplified PCR products were visualized as mentioned above; all the positive cPCR products were purified and sequenced in forward and reverse directions, following the procedures stated overhead.
Results
Leishmania spp. in equids
Of the sampled equids, 57.1% (i.e., 56/98) presented at least one skin lesion (Fig 4), with hyperkeratotic and alopecic nodules on the face, being the most common dermatological signs (Table 1). While on cytological smear examination, no Leishmania spp. amastigotes were observed, one donkey, presenting small nodules on the muzzle, scored positive to L. infantum by qPCR (i.e., Ct: 33 for both skin lesion tissue and blood), and one horse with hyperkeratotic alopecic area on the tail was positive for Leishmania spp. by cPCR (i.e., skin lesion tissue, 18S rRNA, 100% nucleotide identity with GenBank sequences, accession n. MT560279-Leishmania major, MK495994-L. infantum) (Table 1).
Overall, 42.8% (i.e., 42/98) animals had IgG anti-Leishmania spp. titers of 1:40 (i.e., n = 25 for L. infantum, n = 30 for L. martiniquensis, of which n = 13 for both species). In addition, 19 equids (i.e., n = 13 donkeys and n = 6 horses; 19.4%) were seropositive for L. infantum (titers ≥ 1:80) and five of them (i.e., n = 1 horse and n = 4 donkeys) also for L. martiniquensis (titers = 1:80) (Table 1). Most seropositive animals had no dermatological lesions (i.e., 68.4%, 12/19) while both animals molecularly positive for Leishmania spp. scored seronegative.
Insect population and Leishmania spp. detection
Out of 356 sand flies collected (i.e., n = 160 Phlebotomus perniciosus, n = 92 Phlebotomus perfiliewi, n = 16 Phlebotomus neglectus, n = 88 Sergentomyia minuta; n = 142 from Site A and n = 214 from Site B), 284 were females (i.e., n = 134 P. perniciosus, n = 66 P. perfiliewi, n = 7 P. neglectus, and n = 77 S. minuta) (Table 2). While flagellates were not observed during dissection of alive female sand flies, 12 specimens (i.e., 4.5%, 12/268) were molecularly positive for Leishmania spp. (i.e., n = 7 for L. tarentolae and n = 5 for L. infantum) (Table 3, Fig 5). Particularly, one S. minuta from site A scored positive for L. tarentolae by cPCR (i.e., F25/R617, 100% nucleotide identity with GenBank sequence CP119852) and dqPCR (Ct: 24), being also blood fed on humans (i.e., Homo sapiens DNA was detected, 99.7% nucleotide identity with GenBank sequence MG660591). On the other hand, one non-blood fed S. minuta from the same site was positive for L. infantum by cPCR (L5.8S/LITSR, 100% nucleotide identity with GenBank sequence MN503527). Similarly, one P. perniciosus from site B scored positive for L. infantum by qPCR (Ct: 34), being also blood fed on horse (98.3% nucleotide identity with GenBank sequence MN187576) (Table 3). Of the seven female biting midges collected from sites A and B (i.e., n = 1 Culicoides imicola, n = 1 Culicoides circumscriptus, n = 2 Culicoides catanei, n = 1 Culicoides pulicaris complex, n = 1 Culicoides obsoletus complex and n = 1 Culicoides sp.) (Table 2), one C. imicola scored positive for L. infantum by cPCR and qPCR (i.e., F25/R617, 98.83% nucleotide identity with GenBank sequence MW410135 and 99.66% with MH703538; qPCR, Ct: 35) (Table 3).
A) Duplex qPCR (qdPCR) results with blue colored asterisk and dark green asterisk indicating Leishmania tarentolae and Leishmania infantum positive controls, respectively. B) qPCR results with blue colored asterisk indicating L.infantum positive control.
Discussion
Horses and donkeys living in endemic areas for CanL are exposed to Leishmania spp., being bitten by infected insect vectors. Leishmania infantum seroprevalence in equids here recorded with a 1:80 IFAT cut-off value (i.e., 19.4%) was higher than that reported previously in horses from northern/central Italy (i.e., 6.4–13.9%, IFAT, 1:40 dilution cut-off) [12,13], probably because of the higher prevalence of L. infantum in southern Italian regions [28]. In addition, the high percentage of equids exposed to Leishmania spp. (i.e., 42.8%), indicate that the IFAT cut-off of 1:40 is not a reliable positive threshold in Leishmania endemic areas, though employed previously [9]. Thus, a 1:80 cut-off value should be recommended as proper IFAT serum dilution to score equids as seropositive to Leishmania spp., like previously indicated for cats [48].
Seropositivity for L. infantum in animals presenting from none to mild-moderate skin disorders, confirms that the infection may cause moderate skin lesions, suggesting that equid immune response is generally effective and prevents the development of systemic disease [49]. Accordingly, in vitro cellular immune response demonstrated that horses may develop specific lymphocyte proliferation towards L. infantum, with limited antibody production, resulting in an effective control of the infection [14,49]. The above might justify the seronegative results in animals scoring molecularly positive to Leishmania spp., as previously reported [50]. However, the possibility that equids, at the time of sampling, had already downgraded their Leishmania-IgG levels cannot be ruled out, given that the humoral response to L. infantum in horses can be transient [49].
Since L. martiniquensis was not molecularly detected in skin lesions of seropositive animals, the IFAT positive results might be due to a cross-reaction between L. infantum and L. martiniquensis, as suggested in areas where L. infantum occurs in sympatry with other species, such as L. tarentolae [51]. However, the higher exposure recorded for L. martinquensis than L. infantum (i.e., 30 animals vs 25 animals) may deserve further investigations, also considering that low titers are difficult to be interpreted. Under the above circumstances, the exposure of equids to biting midge saliva, through serological assays, might be important to estimate the risk of L. martiniquensis infection, as demonstrated for sand fly bites, which are correlated to the risk of L. infantum infection in dogs and cats [52,53]. Nevertheless, data advocate for molecular tests for EL diagnosis, especially in equids with dermatological lesions, living in or travelled to CanL endemic areas.
The overall Leishmania molecular prevalence recorded in sand flies (i.e., 4.5%) falls in ranges of previous records from the same region (i.e., from 4.7% to 12%) [18,31] and it depends on many epidemiological factors at each sampling site as well as on the collecting efforts. The sand fly species composition found is in agreement with data from previous entomological surveys in southern Italy [54,55]. Likewise, the predominance of S. minuta in Site A and P. perniciosus in Site B, is consistent with the ecology of these two insect species, the first being more abundant in rural areas and the second in peri-urban settings [56,57]. The finding of human and equine DNA in S. minuta and P. perniciosus specimens, respectively, suggest that host choice of these sand flies may depend on hosts’ availability [58]. Accordingly, P. perniciosus was previously demonstrated to feed on horses in Central Italy [58]. Though the L. infantum-positive P. perniciosus found in this study was blood fed on horse, the possibility that the sand fly previously acquired the infection from another host cannot be ruled out. Albeit S. minuta prefers to feed on reptiles [59,60], its feeding on humans was demonstrated experimentally [59]. The finding of L. infantum DNA in S. minuta, the proven vector of L. tarentolae [60], was previously discussed [18,31] and advocates for future researches aiming to isolate L. infantum from “unconventional” hosts (i.e., S. minuta and reptiles), to better understand the epidemiological dynamics driven by the sympatric occurrence of L. infantum and L. tarentolae.
The variety of biting midge species identified (i.e., six different species) is consistent with that reported in Italy and other Mediterranean countries (e.g., Morocco, Spain) [38,39,61]. Similarly, L. infantum DNA was already detected in a wild-caught C. imicola specimen in Tunisia [62], as well as other species of Leishmania from biting midges [63,64]. Noteworthy, the PCR detection of Leishmania spp. does not imply the vector capability of any blood sucking insect [65], since only the observation of metacyclic promastigotes in the stomodeal valve or transmission experiments can ensure the vector competence [66].
Conclusions
Data suggest that equids living in CanL endemic areas are often exposed to Leishmania spp., with minor clinical impact. In such areas, horses or donkeys presenting skin lesions should be searched for Leishmania spp. molecularly, as this diagnostic method should be preferred to serological ones, due to the cross reactions (e.g., L. infantum vs L. martiniquensis) that may occur at IFAT and ELISA tests. Although L. martiniquensis was not detected in equids and biting insects in Italy yet, data advocate for a more in-depth evaluation of Leishmania species among equids, and in the vector populations from the same geographical areas.
Acknowledgments
The authors would like to thank all the practitioners (i.e., Dr. Vito Ricci, Dr.ssa Anna De Leo Ratti, Dr.ssa Stefania Martiradonna) and the staff of “Masseria Russoli” in Martina Franca, as the headquarters of the Center for the Conservation of the Genetic Heritage of the Donkey Breed of Martina Franca of the Apulia region, who contributed to the collection of samples that were analyzed in this study. Authors also thank Professor Canio Buonavoglia for his advice in setting of serological tests, and Dr.ssa Jovana Sadlova for support in the culture of L. martiniquensis.
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