- Open Access
Detection of Leishmania tarentolae in lizards, sand flies and dogs in southern Italy, where Leishmania infantum is endemic: hindrances and opportunities
Parasites & Vectors volume 14, Article number: 461 (2021)
Leishmania tarentolae is a protozoan isolated from geckoes (Tarentola annularis, Tarentola mauritanica), which is considered non-pathogenic and is transmitted by herpetophilic Sergentomyia spp. sand flies. This species occurs in sympatry with Leishmania infantum in areas where canine leishmaniasis is endemic. In the present study, we investigated the circulation of L. tarentolae and L. infantum in sand flies, dogs and lizards in a dog shelter in southern Italy, where canine leishmaniasis by L. infantum is endemic.
Sheltered dogs (n = 100) negative for Leishmania spp. (March 2020) were screened by immunofluorescence antibody test (IFAT) using promastigotes of both species at two time points (June 2020 and March 2021). Whole blood from dogs, tissues of Podarcis siculus lizards (n = 28) and sand flies (n = 2306) were also sampled and tested by a duplex real-time PCR (dqPCR). Host blood meal was assessed in sand flies by PCR.
Overall, 16 dogs became positive for L. infantum and/or L. tarentolae by IFAT at one or both sampling periods. One canine blood sample was positive for L. infantum, whilst two for L. tarentolae by dqPCR. At the cytology of lizard blood, Leishmania spp. amastigote-like forms were detected in erythrocytes. Twenty-two tissue samples, mostly lung (21.4%), scored molecularly positive for L. tarentolae, corresponding to 10 lizards (i.e., 35.7%). Of the female Sergentomyia minuta sampled (n = 1252), 158 scored positive for L. tarentolae, four for L. infantum, and one co-infected. Two Phlebotomus perniciosus (out of 29 females) were positive for L. tarentolae. Engorged S. minuta (n = 10) fed on humans, and one P. perniciosus, positive for L. tarentolae, on lagomorphs.
Dogs and lacertid lizards (Podarcis siculus) were herein found for the first time infected by L. tarentolae. The detection of both L. tarentolae and L. infantum in S. minuta and P. perniciosus suggests their sympatric circulation, with a potential overlap in vertebrate hosts. The interactions between L. tarentolae and L. infantum should be further investigated in both vectors and vertebrate hosts to understand the potential implications for the diagnosis and control of canine leishmaniasis in endemic areas.
Zoonotic visceral leishmaniasis, caused by Leishmania infantum (Kinetoplastida, Trypanosomatidae), is a neglected disease of medical and veterinary importance, which impacts health, society and the economy in many tropical, subtropical and temperate regions of the globe . Indeed, this disease affects mainly poor people  and may be fatal if not treated timely and properly. Infected dogs are the main reservoirs of L. infantum in the domestic and peri-domestic environments , with mainly subclinical presentation and only a small proportion manifesting overt clinical disease . The causative agent is transmitted by bites of phlebotomine sand flies of the genera Phlebotomus in the Old World [5, 6] and Lutzomyia in the New World [7, 8]. Meanwhile, phlebotomine sand flies of the genus Sergentomyia are known to feed primarily on cold-blooded animals  and are associated to Leishmania spp. in lizards [9,10,11,12]. Nonetheless, DNA of L. infantum has been detected in Sergentomyia minuta [13, 14], suggesting it can feed also on available endothermic tetrapod animals. This picture has also been corroborated by other reports of L. infantum DNA in several Sergentomyia spp., such as Sergentomyia dubia, Sergentomyia magna and Sergentomyia schewtzi in Africa , and S. minuta from endemic areas of canine leishmaniasis (CanL) in Europe [13, 14, 16,17,18,19,20,21]. Meanwhile, the DNA and/or amastigote forms of pathogenic Leishmania spp. (i.e., Leishmania donovani, Leishmania tropica and Leishmania turanica) have been detected in reptiles [22,23,24,25], therefore suggesting their potential role as reservoirs of mammalian pathogenic Leishmania spp. [24, 26, 27]. On the other hand, Leishmania tarentolae (subgenus Sauroleishmania) is a less regarded trypanosomatid infecting geckoes (e.g., Tarentola mauritanica), possibly transmitted by herpetophilic Sergentomyia spp. in Europe, North Africa and the Middle East [28, 29]. Incidentally, L. tarentolae is widely considered non-pathogenic. Nonetheless, some strains of this species (e.g., RTAR/FR/78/LEM125) may cause transient infection in mammals under laboratory conditions, as this species can differentiate into amastigote-like forms [30,31,32,33]. However, the molecular findings of L. tarentolae in a human mummy in Brazil , as well as in human blood  in central Italy, suggest its capacity to infect mammals. Nevertheless, the pathogenicity, virulence and overall deleterious effects of L. tarentolae in mammals are still unknown. In addition, given the high similarity in gene composition with L. infantum (i.e., 90%), L. tarentolae is considered a model for recombinant protein production and vaccine candidate [34,35,36], which could mean that natural infection with L. tarentolae may have a protective effect against L. infantum . Given the variations in dog antibody levels between seasons of sand fly activity and the sympatric occurrence of both Leishmania species, we investigated the circulation of L. tarentolae and L. infantum in sand flies, dogs and lizards in a dog shelter in southern Italy where CanL by L. infantum is endemic.
Study area and sample collection
One hundred dogs which scored negative to Leishmania spp. on molecular and serological tests in March 2020 were re-sampled in June 2020 and March 2021 in a shelter located in a CanL-endemic area in Apulia region, southern Italy (40.419326N, 18.165582E, Lecce) . The shelter is built in a dry and windy area 8.0 km from the nearest seaside (Fig. 1a). The environment around the shelter is characterized by few olive trees, withered grass, no water sources and surrounded by the typical muretti a secco (stone walls) where reptiles (i.e., Podarcis siculus lizards, Hierophis viridiflavus carbonarius snakes, and T. mauritanica geckoes) and rodents thrive. Dogs with a minimum age of 7 months were included in the study; signalment (i.e., age, sex, breed) and anamnestic data (i.e., previous protozoan and bacterial infection, and treatment) were recorded at time of enrollment. Moreover, a complete physical examination was performed by a veterinary clinician to assess the health status of the enrolled dogs. From each dog, whole blood was collected in vacuum containers with EDTA (2.5 ml) and serum collection tubes with clot activator (5 ml).
From May to November 2020, sand flies were collected biweekly using 64 sticky papers (21.0 cm × 29.7 cm, covering up to 4 m2) (Fig. 1b) and two CDC light traps were set from 5:00 p.m. to 8:00 a.m. Collections were carried out during the sand fly activity season  until the total disappearance/absence of sand flies (i.e., three consecutive negative captures). All specimens were stored in labeled glass vials containing 70% ethanol then morphologically identified using taxonomic keys and descriptions [39, 40].
Reptiles were captured in the area of the shelter, on the same walls where the sticky traps were placed (Fig. 1c), by lassoing or by hand. Species of reptiles were identified using reference keys , and then physically examined to assess their health status. Anamnestic data (e.g., species, biological stage, sex, physical abnormalities such as tail loss or predator-induced wounds) were recorded in each animal’s file. A small amount of blood was obtained via lizard tail fracture or by cardiocentesis when animals were adults and non-gravid females. Blood samples were stored at −20 °C and tail tissue in 70% ethanol. For each animal, blood smears were performed and then assessed for the presence of Leishmania parasites  using Diff-Quik stain . Smears were rinsed in tap water to remove excess stain, and later evaluated using an optical microscope (LEICA DM LB2, Germany). Fecal samples were also collected from each animal. Captured lizards were humanely euthanized according to protocols  and dissected. Intestine, heart, kidneys, liver, lungs, spleen and skeletal muscle were individually collected and frozen at −20 °C.
Serum samples from all enrolled dogs were tested to assess the exposure to L. infantum and L. tarentolae. An IFAT for the detection of IgG anti-L. infantum was performed as previously described (Fig. 2a) , whereas for antibodies against L. tarentolae, the IFAT was performed using promastigotes of L. tarentolae (strain RTAR/IT/81/ISS21-G.6c) as antigen (Fig. 2b) following the same procedure as for L. infantum IFAT. Serum samples from a dog positive for L. infantum by cytological and molecular analyses, and a healthy dog negative for L. infantum, were used as positive and negative controls, respectively, for both IFAT. Samples were scored as positive when they produced a clear cytoplasmic and membrane fluorescence of promastigotes from a cut-off dilution of 1:80 . Positive sera were titrated by serial dilutions until negative results were obtained.
Genomic DNA (gDNA) was extracted from the thorax and abdomen (heads and last segments were removed for morphological identification) of each female sand fly (n = 1281) using an in-house method as previously described . Lizard tissues (i.e., intestine, heart, kidneys, liver, lungs, spleen, skeletal muscle and tail), as well as blood samples from lizards and dogs, were extracted using two commercial kits, GenUP gDNA and GenUP Blood DNA kits (Biotechrabbit GmbH, Hennigsdorf, Germany), respectively, according to the manufacturer’s instructions. DNA from lizard fecal samples was extracted using a specific commercial kit (DNeasy PowerSoil Kit, QIAGEN, Hilden Germany) following the manufacturer’s instructions. All samples were tested by duplex real-time PCR (dqPCR) for detection of L. infantum and L. tarentolae (samples were considered positive with quantitation cycle (Cq) values up to 38.0 and 38.6, respectively), as previously described . Approximately 100 ng of gDNA (with the exception of the no-template control) was added to each dqPCR. gDNA from cultured promastigotes of L. infantum, originally retrieved from a dog living in Italy (zymodeme MON-1) (Fig. 2c), and L. tarentolae (strain RTAR/IT/81/ISS21-G.6c) (Fig. 2d) was used as positive controls. For sequences analyses, Leishmania dqPCR-positive samples were amplified by conventional PCR (cPCR) using primers L5.8S/LITSR targeting the partial region of the internal transcribed spacer 1 (ITS1, ~ 300 bp) and PCR protocol run as described elsewhere .
Engorged sand flies (n = 22) and all specimens that scored positive for Leishmania spp. were tested for blood-meal determination by cPCR using primers targeting the vertebrate host mitochondrial cytochrome b (350 bp), and a PCR protocol was run as previously described . All PCR reactions consisted of 4 μl of gDNA and 46 μl of PCR mix containing 3 mM MgCl2, 10 mM Tris–HCl (pH 8.3) and 50 mM KCl, 125 μM of each dNTP, 1 pmol/μl of each primer and 2 U of AmpliTaq Gold (Applied Biosystems, Foster City, CA, USA). Amplified products were examined on 2% agarose gels stained with GelRed (VWR International PBI, Milan, Italy) and visualized on a Gel Logic 100 gel documentation system (Kodak, NY, USA). Amplicons were purified and sequenced in both directions using the same primers as for PCR, employing the Big Dye Terminator v.3.1 chemistry in an automated sequencer (3130 Genetic Analyzer, Applied Biosystems, Foster City, CA, USA). All sequences were aligned using the ClustalW program  and compared with those available in GenBank using the BLASTn tool (http://blast.ncbi.nlm.nih.gov/Blast.cgi).
To determine genetic clustering of L. tarentolae, the representative ITS1 sequences obtained from lizard, sand fly and dog samples and from reference strains of L. tarentolae and L. infantum were phylogenetically analyzed along with those of other Leishmania spp. available in the GenBank database. Phylogenetic relationships were inferred using the maximum likelihood (ML) method based on the Kimura 2-parameter model , and discrete gamma distribution was used to model evolutionary rate differences among sites, selected by best-fit model analysis and based on the lowest score obtained by Bayesian information criterion (BCI) using MEGA6 software . Evolutionary analyses were conducted with 5000 bootstrap replications using MEGA6 software . The corresponding ITS1 sequence of Trypanosoma brucei (GenBank: KU552356.1) was used as outgroup.
Of 100 dogs serologically examined, 16 scored positive against promastigotes of L. infantum and/or L. tarentolae by IFAT at one or both sampling periods (June 2020 and March 2021; Table 1). In particular, three dogs scored positive only against promastigotes of L. infantum (titer of 1:80) and five of L. tarentolae (titer up to 1:160). Of the eight animals positive for both species, four were positive at both time points, the remaining with different combinations (Table 1). Of dog blood samples tested by dqPCR, one collected in March 2021 scored positive for L. infantum (Cq = 37.2), whilst two for L. tarentolae (one in June 2020, Cq = 36.2; one in March 2021, Cq = 36.9).
A total of 2306 phlebotomine sand flies (2138 S. minuta and 168 P. perniciosus) were collected, of which 1281 were females (i.e., 1252 S. minuta and 29 P. perniciosus). Of female sand flies, 161 scored positive for Leishmania spp. (12.6%) by dqPCR (Table 2). Among them, 155 S. minuta (95.7%) and two P. perniciosus (1.2%) were positive for L. tarentolae, whilst four S. minuta scored positive for L. infantum (2.5%), and only one was co-infected (0.6%) for both Leishmania species. In addition, of 22 engorged females tested (14 S. minuta and eight P. perniciosus), the host mitochondrial cytb was amplified from 10 specimens (45.4%, nine S. minuta and one P. perniciosus). Cytb sequences detected in S. minuta displayed 99.67% of nucleotide identity with that of Homo sapiens (GenBank: JN315800), whilst that from P. perniciosus showed 84.4% of identity with lagomorph species Ochotona cansus (GenBank: MN547415).
Podarcis siculus lizards (n = 28) were captured in the same study area (Fig. 1d), including 14 males and 14 females, whereas no snakes or geckoes were collected. Cytological blood smear examination revealed Leishmania spp. amastigote-like forms inside erythrocytes (Fig. 3a) and promastigote-like (Fig. 3b) forms in one lizard. Out of 224 lizard tissue samples examined by dqPCR, 22 samples (i.e., intestine, heart, kidneys, liver, lungs, spleen and skeletal muscle) scored positive for L. tarentolae, corresponding to 10 positive lizards (35.7%). Lungs had the highest number of positive samples (six, 21.4%), whereas the lowest Cq value (24.7) was recorded from liver (Table 3). Lizard blood, tails and fecal samples were all negative by dqPCR. BLASTn analysis of ITS1 sequences confirmed the L. tarentolae species identification showing a nucleotide identity of 98.7% with the reference sequence (GenBank: KU680858) available in the GenBank database and with L. tarentolae strain RTAR/IT/81/ISS21-G.6c. The phylogram of ITS1 showed a close phylogenetic relationship by clustering all L. tarentolae sequences herein obtained in a species-specific clade (Sauroleishmania), with the exclusion of the other Leishmania species (bootstrap value of 95%) (Fig. 4). Sequences obtained for L. tarentolae from lizards, dogs and sand flies were deposited in GenBank (MW832546, MW832547, MW832548).
Data herein presented suggests that dogs may be exposed to L. tarentolae, a species largely disregarded by the scientific community since it is merely considered a saurian-associated trypanosomatid, yet it occurs in sympatry with L. infantum. In addition, dogs after initial exposure against promastigotes of L. tarentolae may then seroconvert, remaining seropositive even during the non-transmission sand fly season, suggesting a persistent rather than transient presence of L. tarentolae in a non-permissive host. This event may happen in endemic areas where reptiles, herpetophilic sand flies and dogs share the same environment, and both Leishmania spp. occur in sympatry.
While L. tarentolae has been previously reported exclusively infecting geckoes (i.e., Tarentola annularis and T. mauritanica) [53,54,55], the detection in lacertid lizards, P. siculus, is a new finding, which could be of major importance to better understand the epidemiology and host preference of this protozoan. The occurrence of L. tarentolae in lizards was confirmed both by the detection of Leishmania amastigote-like forms in erythrocytes (Fig. 3a) and by cPCR and dqPCR. At cytology, the Leishmania forms differ from those of L. infantum in that amastigote-like forms infected erythrocytes rather than leucocytes, with possible promastigote-like forms circulating freely in blood (Fig. 3b). The life cycle of L. tarentolae is yet to be fully unraveled, though promastigotes and amastigote-like forms have been previously recorded in blood and intestinal lumen from geckoes [10, 22]. Unexpectedly, rather than blood and feces, results of the dqPCR indicated that organs, such as lungs and liver of lizards, are the preferential samples for diagnosing the infection by L. tarentolae, probably due to a low parasitic load of promastigotes/amastigotes-like forms in blood. Indeed, parenchymatous organs showed higher amount of L. tarentolae DNA, which agrees with higher parasitic loads detected. Blood is not considered the ideal sample for the molecular detection of Leishmania spp. due to the low circulation of the parasite . This could represent a hindrance for the molecular identification in mammals, despite the detection of two samples positive with high Cq values.
Podarcis lacertid lizards (commonly known as wall lizards) are synanthropic reptiles, which may play a role as reservoirs of other pathogens causing zoonotic diseases such as Lyme disease and rickettsiosis [56, 57]. These reptiles live in wall cracks, under stones and anywhere they find shelter and food, with a wide distribution throughout the Mediterranean basin . The microhabitats where lizards live are similar to that of breeding and resting sites of sand flies . The high prevalence of infection in lizards overlaps the abundance of herpetophilic S. minuta and of P. perniciosus, which is the main vector of L. infantum . Nonetheless, the finding of S. minuta as the most abundant species (92.7%) compared to P. perniciosus (7.3%) was already observed in other dog shelters from southern Italy where L. infantum is prevalent, such as in Apulia , Sicily [60,61,62,63], as well as Morocco , Portugal  and Spain . In addition, the low number of P. perniciosus collected may be correlated to the species phenology and environmental preferences. Indeed, P. perniciosus is more abundant in domestic or peri-urban settings, and S. minuta in rural or wild areas, similar to the characteristics of the studied shelter . As for many other species of phlebotomine sand flies, S. minuta displays a rather catholic feeding behavior  depending on host availability. The detection of human blood in S. minuta suggests the opportunistic attitude of this species, as already demonstrated in Sicily where 64% of engorged sand flies scored positive for human blood . The ectoparasiticide treatment of dogs could have affected the sand fly species composition, similarly to a previous study from a dog shelter where a group of animals were treated with a combination of 10% imidacloprid/4.5% flumethrin collar, and the remaining were left untreated . In that study, S. minuta was the most common sand fly species identified (66.6%) throughout a collection period of 2 years, followed by P. perniciosus (15.1%), Phlebotomus neglectus (8.8%) and Phlebotomus papatasi (0.23%). Although S. minuta has been found molecularly positive for L. tarentolae, the vector capacity has never been demonstrated. However, transmission of this Leishmania sp. most likely occurs as described for mammalian Leishmania, through a pool feeding mechanism . Also, the direct ingestion of the sand fly by lizards cannot be ruled out . Given that the dog population was under an ectoparasiticide treatment and considering the high abundance of S. minuta, dogs could have ingested infected L. tarentolae sand flies. Another peculiar result of this study is the lack of reptile blood in the engorged S. minuta analyzed. This can agree with the hypothesis of a reduced density of preferred reptile hosts in the shelter area, as a consequence of the high predatory pressure exerted by dogs. Hence, further studies are advocated to better elucidate the reptilian and mammalian interactions in the life cycle of L. tarentolae.
The molecular detection of L. tarentolae in the blood of two dogs is unprecedented, and the exposure to this parasite was confirmed by the seropositivity in 16 dogs, of which eight scored positive for both Leishmania species and five against promastigotes of L. tarentolae only. This result is new to science, since IFAT using promastigotes of L. tarentolae was herein described for the first time. Although the IFAT method reported should be further validated using serum samples of animals purposely infected with both Leishmania spp., L. tarentolae exposure has been previously demonstrated to be associated with transient infections in mammals [30,31,32,33]. Positive dqPCR blood samples for both species of Leishmania were from seronegative dogs at both time points, suggesting a recent or transient presence of the parasite for which the animal had not yet seroconverted. In addition, the exposure of animals to this protozoon is also supported by the detection of L. tarentolae in two P. perniciosus, which usually feed on dogs. The association of this Leishmania species to sand flies of the genus Phlebotomus was already described in 6.6% of Phlebotomus perfiliewi examined in Central Italy .
Though the seropositivity of dogs against promastigotes of L. tarentolae does not imply the reservoir competence of canids, these data are of medical and veterinary relevance. Indeed, the detection of a significant reduction in anti-L. infantum antibody titers in 55.4% of L. infantum-seropositive and clinically healthy dogs from the same shelter was recently demonstrated after sampling one year apart . A large proportion of these animals (44.4%) became seronegative (i.e., below the cut-off value of 1:80), further suggesting a possible L. tarentolae transient exposure. Indeed, although the IFAT is considered the gold standard for the diagnosis of L. infantum, as it is based on the visualization of the immunofluorescence on the whole promastigotes, cross-reactions with highly similar species of Leishmania may occur. This event was observed in eight dogs which had titers for both species. However, co-infections could also have caused cross-reactivity, given the discrepancies in titers for both species (e.g., dog positive for L. infantum with titers 1:1280 and to L. tarentolae with 1:160). Given the relevance of serology in epidemiological studies and in the management of diseased patients, the variations in antibody titers requires careful examination. Under the above circumstances, considering that the IFAT for the detection of antibodies against Leishmania promastigotes represents the reference serological method for CanL diagnosis and screening, as well as for clinical staging and therapeutic purposes [69, 70], the cross-reactivity between the two species of Leishmania might directly impact the interpretation of CanL-related clinical signs, prognosis and treatment. Finally, the sympatric occurrence of L. infantum and L. tarentolae in sand flies (e.g., co-infection in S. minuta) in the specific epidemiological context herein studied could result in hybridization events between these two species. This event has been previously experimentally confirmed for L. infantum and Leishmania major in Lutzomyia longipalpis . The possibility of genetic exchange and hybridization events could have implications for the pathogenicity and visceralization capacity of an otherwise innocuous species such as L. tarentolae. However, these hypotheses need further research.
Under specific epidemiological contexts where canids, reptiles, herpetophilic sand flies, L. infantum and L. tarentolae occur in sympatry, dogs may be exposed to L. tarentolae. Results of this study further suggest the low host specificity of L. tarentolae in the ability to infect other reptiles (i.e., lacertid lizards) and likely mammals on which S. minuta may feed. Serological findings indicate that a cross-reactivity for both species of Leishmania may occur, having diagnostic and clinical implications for seropositive healthy dogs. Future studies should focus on determining the prevalence of L. tarentolae infection in dogs and its possible interactions with L. infantum in areas where they are sympatric.
Availability of data and materials
All data generated or analyzed during this study are included in this published article. The sequences generated in this study were deposited in GenBank (MW832546, MW832547, MW832548).
Centers for Disease Control and Prevention
- cytb :
Duplex real-time PCR
Ethylenediamine tetraacetic acid
Immunofluorescence antibody test
Polymerase chain reaction
Otranto D, Dantas-Torres F. The prevention of canine leishmaniasis and its impact on public health. Trends Parasitol. 2013;29:339–45.
Okwor I, Uzonna J. Social and economic burden of human leishmaniasis. Am J Trop Med Hyg. 2016;94(3):489–93.
Dantas-Torres F. The role of dogs as reservoirs of Leishmania parasites, with emphasis on Leishmania (Leishmania) infantum and Leishmania (Viannia) braziliensis. Vet Parasitol. 2007;149(3–4):139–46.
Solano-Gallego L, Miró G, Koutinas A, Cardoso L, Pennisi MG, Ferrer L, et al. LeishVet guidelines for the practical management of canine leishmaniosis. Parasit Vectors. 2011;4:86.
Maroli M, Feliciangeli MD, Bichaud L, Charrel RN, Gradoni L. Phlebotomine sandflies and the spreading of leishmaniases and other diseases of public health concern. Med Vet Entomol. 2013;27(2):123–47.
Alten B, Maia C, Afonso MO, Campino L, Jiménez M, González E, et al. Seasonal dynamics of phlebotomine sand fly species proven vectors of Mediterranean leishmaniasis caused by Leishmania infantum. PLoS Negl Trop Dis. 2016;10(2):e0004458.
Dantas-Torres F, Solano-Gallego L, Baneth G, Ribeiro VM, de Paiva-Cavalcanti M, Otranto D. Canine leishmaniosis in the Old and New Worlds: unveiled similarities and differences. Trends Parasitol. 2012;28(12):531–8.
Silva MDD, Galvis-Ovallos F, Casanova C, Silva VGD, Leonel JAF, Oliveira TMFS, et al. Natural infection of Lutzomyia longipalpis (Cembrene-1 population) with Leishmania infantum in a new visceral leishmaniasis focus in the eastern region of São Paulo State, Brazil. Rev Soc Bras Med Trop. 2021;54:e05862020.
Lewis DJ. The phlebotomine sandflies (Diptera: Psychodidae) of the Oriental Region. Syst Entomol. 1987;12:163–80.
Killick-Kendrick R, Lainson R, Rioux JA, Saf'janova VM. The taxonomy of Leishmania-like parasites of reptiles. In: Rioux JA. Leishmania: Taxonomie et Phylogenèse. Application Éco-epidemiologiques (Colloque International du CNRS/INSERM, 1984), MEE, Montpellier. 1986; 143–8.
Noyes HA, Arana BA, Chance ML, Maingon R. The Leishmania hertigi (Kinetoplastida; Trypanosomatidae) complex and the lizard Leishmania: their classification and evidence for a neotropical origin of the Leishmania-Endotrypanum clade. J Eukaryot Microbiol. 1997;44(5):511–7.
Tuon FF, Neto VA, Amato VS. Leishmania: origin, evolution and future since the Precambrian. FEMS Immunol Med Microbiol. 2008;54(2):158–66.
Iatta R, Zatelli A, Laricchiuta P, Legrottaglie M, Modry D, Dantas-Torres F, et al. Leishmania infantum in tigers and sand flies from a leishmaniasis-endemic area, Southern Italy. Emerg Infect Dis. 2020;26(6):1311–4.
Pombi M, Giacomi A, Barlozzari G, Mendoza-Roldan J, Macrì G, Otranto D, et al. Molecular detection of Leishmania (Sauroleishmania) tarentolae in human blood and Leishmania (Leishmania) infantum in Sergentomyia minuta: unexpected host-parasite contacts. Med Vet Entomol. 2020;34(4):470–5.
Senghor MW, Niang AA, Depaquit J, Ferté H, Faye MN, Elguero E, et al. Transmission of Leishmania infantum in the canine leishmaniasis focus of Mont-Rolland, Senegal: ecological, parasitological and molecular evidence for a possible role of Sergentomyia sand flies. PLoS Negl Trop Dis. 2016;10(11):e0004940.
Tarallo VD, Dantas-Torres F, Lia RP, Otranto D. Phlebotomine sand fly population dynamics in a leishmaniasis endemic peri-urban area in southern Italy. Acta Trop. 2010;116(3):227–34.
Campino L, Cortes S, Dionísio L, Neto L, Afonso MO, Maia C. The first detection of Leishmania major in naturally infected Sergentomyia minuta in Portugal. Mem Inst Oswaldo Cruz. 2013;108(4):516–8.
Bravo-Barriga D, Parreira R, Maia C, Blanco-Ciudad J, Afonso MO, et al. First molecular detection of Leishmania tarentolae-like DNA in Sergentomyia minuta in Spain. Parasitol Res. 2016;115(3):1339–44.
Maia C, Parreira R, Cristóvão JM, Freitas FB, Afonso MO, Campino L. Molecular detection of Leishmania DNA and identification of blood meals in wild caught phlebotomine sand flies (Diptera: Psychodidae) from southern Portugal. Parasit Vectors. 2015;8:173.
Latrofa MS, Iatta R, Dantas-Torres F, Annoscia G, Gabrielli S, Pombi M, et al. Detection of Leishmania infantum DNA in phlebotomine sand flies from an area where canine leishmaniosis is endemic in southern Italy. Vet Parasitol. 2018;253:39–42.
González E, Molina R, Aldea I, Iriso A, Tello A, Jiménez M. Leishmania sp. detection and blood-feeding behaviour of Sergentomyia minuta collected in the human leishmaniasis focus of southwestern Madrid, Spain (2012–2017). Transbound Emerg Dis. 2020;67(3):1393–400.
Simpson L, Holz G Jr. The status of Leishmania tarentolae/Trypanosoma platydactyli. Parasitol Today. 1988;4(4):115–8.
Belova EM. Reptiles and their importance in the epidemiology of leishmaniasis. Bull World Health Organ. 1971;44(4):553–60.
Zhang JR, Guo XG, Liu JL, Zhou TH, Gong X, Chen DL, et al. Molecular detection, identification and phylogenetic inference of Leishmania spp. in some desert lizards from Northwest China by using internal transcribed spacer 1 (ITS1) sequences. Acta Trop. 2016;162:83–94.
Chen H, Li J, Zhang J, Guo X, Liu J, He J, et al. Multi-locus characterization and phylogenetic inference of Leishmania spp. in snakes from Northwest China. PLoS ONE. 2019;14(4):e0210681.
Mendoza-Roldan JA, Modry D, Otranto D. Zoonotic parasites of reptiles: a crawling threat. Trends Parasitol. 2020;36(8):677–87.
Mendoza-Roldan JA, Mendoza-Roldan MA, Otranto D. Reptile vector-borne diseases of zoonotic concern. Int J Parasitol Parasites Wildl. 2021;15:132–42 (Published 2021 Apr 22).
Telford SR. A review of trypanosomes of gekkonid lizards, including the description of five new species. Syst Parasitol. 1995;31:37–52.
Halla U, Korbel R, Mutschmann F, Rinder M. Blood parasites in reptiles imported to Germany. Parasitol Res. 2014;113(12):4587–99.
Adler S. The behavior of a lizard Leishmania in hamsters and baby mice. Rev Inst Med Trop Sao Paulo. 1962;4:61–4.
Breton M, Tremblay MJ, Ouellette M, Papadopoulou B. Live nonpathogenic parasitic vector as a candidate vaccine against visceral leishmaniasis. Infect Immun. 2005;73:6372–82.
Taylor VM, Muñoz DL, Cedeño DL, Vélez ID, Jones MA, Robledo SM. Leishmania tarentolae: utility as an in vitro model for screening of antileishmanial agents. Exp Parasitol. 2010;126(4):471–5.
Novo SP, Leles D, Bianucci R, Araujo A. Leishmania tarentolae molecular signatures in a 300 hundred-years-old human Brazilian mummy. Parasit Vectors. 2015;8:72.
Mizbani A, Taheri T, Zahedifard F, Taslimi Y, Azizi H, Azadmanesh K, et al. Recombinant Leishmania tarentolae expressing the A2 virulence gene as a novel candidate vaccine against visceral leishmaniasis. Vaccine. 2009;28(1):53–62.
Niimi T. Recombinant protein production in the eukaryotic protozoan parasite Leishmania tarentolae: a review. Methods Mol Biol. 2012;824:307–15.
Klatt S, Simpson L, Maslov DA, Konthur Z. Leishmania tarentolae: Taxonomic classification and its application as a promising biotechnological expression host. PLoS Negl Trop Dis. 2019;13(7):e0007424.
Saljoughian N, Taheri T, Rafati S. Live vaccination tactics: possible approaches for controlling visceral leishmaniasis. Front Immunol. 2014;5:134 (Published 2014 Mar 31).
Panarese R, Iatta R, Beugnet F, Otranto D. Incidence of Dirofilaria immitis and Leishmania infantum infections in sheltered dogs from Southern Italy. Transbound Emerg Dis. 2021. https://doi.org/10.1111/tbed.14025.
Killick-Kendrick R, Tang Y, Killick-Kendrick M, et al. The identification of female sandflies of the subgenus Larroussius by the morphology of the spermathecal ducts. Parassitologia. 1991;33(Suppl):335–47.
Dantas-Torres F, Tarallo VD, Otranto D. Morphological keys for the identification of Italian phlebotomine sand flies (Diptera: Psychodidae: Phlebotominae). Parasit Vectors. 2014;7:479.
Arnold EN. Reptiles and amphibians of Europe. 2nd ed. Princeton: Princeton University Press; 2002.
Telford SR. Hemoparasites of the Reptilia. Boca Raton: CRC Press; 2009.
Skipper R, Destephano D. A rapid stain for Campylobacter pylori in gastrointestinal tissue sections using Diff-Quik®. J Histotechnol. 1989;4:303–4.
Warren K. Reptile euthanasia—no easy solution? Pac Conserv Biol. 2014;20:25–7.
Otranto D, Testini G, Dantas-Torres F, et al. Diagnosis of canine vector-borne diseases in young dogs: a longitudinal study. J Clin Microbiol. 2010;48(9):3316–24.
Otranto D, Paradies P, de Caprariis D, Stanneck D, Testini G, Grimm F, et al. Toward diagnosing Leishmania infantum infection in asymptomatic dogs in an area where leishmaniasis is endemic. Clin Vaccine Immunol. 2009;16(3):337–43.
Sangioni LA, Horta MC, Vianna MC, et al. Rickettsial infection in animals and Brazilian spotted fever endemicity. Emerg Infect Dis. 2005;11(2):265–70.
Latrofa MS, Mendoza-Roldan JA, Manoj RRS, Pombi M, Dantas-Torres F, Otranto D. A duplex real-time PCR assay for the detection and differentiation of Leishmania infantum and Leishmania tarentolae in vectors and potential reservoir hosts. Entomol Gen. 2021. in press
El Tai NO, El Fari M, Mauricio I, Miles MA, Oskam L, El Safi SH, et al. Leishmania donovani: intraspecific polymorphisms of Sudanese isolates revealed by PCR-based analyses and DNA sequencing. Exp Parasitol. 2001;97(1):35–44.
Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H, et al. Clustal W and Clustal X version 2.0. Bioinformatics. 2007;23(21):2947–8.
Kimura M. A simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences. J Mol Evol. 1980;16(2):111–20.
Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. MEGA6: molecular evolutionary genetics analysis version 6.0. Mol Biol Evol. 2013;30(12):2725–9.
Pozio E, Gramiccia M, Gradoni L, Maroli M. Hemoflagellates in Cyrtodactylus kotschyi (Steindachner, 1870) (Reptilia, Gekkonidae) in Italy. Acta Trop. 1983;40(4):399–400.
Elwasila M. Leishmania tarentolae Wenyon, 1921 from the gecko Tarentola annularis in the Sudan. Parasitol Res. 1988;74(6):591–2.
Simpson L, Frech GC, Maslov DA. RNA editing in trypanosomatid mitochondria. Methods Enzymol. 1996;264:99–121.
Mendoza-Roldan JA, Colella V, Lia RP, Nguyen VL, Barros-Battesti DM, Iatta R, et al. Borrelia burgdorferi (sensu lato) in ectoparasites and reptiles in southern Italy. Parasit Vectors. 2019;12(1):35.
Mendoza-Roldan JA, Ravindran Santhakumari Manoj R, Latrofa MS, Iatta R, Annoscia G, Lovreglio P, et al. Role of reptiles and associated arthropods in the epidemiology of rickettsioses: a one health paradigm. PLoS Negl Trop Dis. 2021;15(2):e0009090.
Kaliontzopoulou A, Brito JC, Carretero MA, Larbes S, Harris DJ. Modelling the partially unknown distribution of wall lizards (Podarcis) in North Africa: ecological affinities, potential areas of occurrence, and methodological constraints. Can J Zool. 2008;86:992–1001.
Jiménez M, González E, Iriso A, Marco E, Alegret A, Fúster F, et al. Detection of Leishmania infantum and identification of blood meals in Phlebotomus perniciosus from a focus of human leishmaniasis in Madrid, Spain. Parasitol Res. 2013;112(7):2453–9.
Gaglio G, Brianti E, Napoli E, Falsone L, Dantas-Torres F, Tarallo VD, et al. Effect of night time-intervals, height of traps and lunar phases on sand fly collection in a highly endemic area for canine leishmaniasis. Acta Trop. 2014;133:73–7.
Lisi O, D’Urso V, Vaccalluzzo V, Bongiorno G, Khoury C, Severini F, et al. Persistence of phlebotomine Leishmania vectors in urban sites of Catania (Sicily, Italy). Parasit Vectors. 2014;7:560.9.
Brianti E, Napoli E, Gaglio G, Falsone L, Giannetto S, Solari Basano F, et al. Field evaluation of two different treatment approaches and their ability to control fleas and prevent canine leishmaniosis in a highly endemic area. PLoS Negl Trop Dis. 2016;10(9):e0004987 (Published 2016 Sep 15).
Abbate JM, Maia C, Pereira A, Arfuso F, Gaglio G, Rizzo M, et al. Identification of trypanosomatids and blood feeding preferences of phlebotomine sand fly species common in Sicily, Southern Italy. PLoS ONE. 2020;15(3):e0229536 (Published 2020 Mar 10).
Daoudi MM, Boussaa S, Boumezzough A. Modeling spatial distribution of Sergentomyia minuta (Diptera: Psychodidae) and Its potential implication in leishmaniasis transmission in Morocco. J Arthropod Borne Dis. 2020;14(1):17–28 (Published 2020 Mar 31).
Pereira S, Pita-Pereira D, Araujo-Pereira T, Britto C, Costa-Rego T, Ferrolho J, et al. First molecular detection of Leishmania infantum in Sergentomyia minuta (Diptera, Psychodidae) in Alentejo, southern Portugal. Acta Trop. 2017;174:45–8.
Rossi E, Bongiorno G, Ciolli E, Di Muccio T, Scalone A, Gramiccia M, et al. Seasonal phenology, host-blood feeding preferences and natural Leishmania infection of Phlebotomus perniciosus (Diptera, Psychodidae) in a high-endemic focus of canine leishmaniasis in Rome province. Italy Acta Trop. 2008;105(2):158–65.
Otranto D, Dantas-Torres F, de Caprariis D, Di Paola G, Tarallo VD, Latrofa MS, et al. Prevention of canine leishmaniosis in a hyper-endemic area using a combination of 10% imidacloprid/4.5% flumethrin. PLoS ONE. 2013;8(2):e56374.
Cavalera MA, Iatta R, Panarese R, Mendoza-Roldan JA, Gernone F, Otranto D, et al. Seasonal variation in canine anti-Leishmania infantum antibody titres. Vet J. 2021;271:105638.
Oliva G, Roura X, Crotti A, Maroli M, Castagnaro M, Gradoni L, et al. Guidelines for treatment of leishmaniasis in dogs. J Am Vet Med Assoc. 2010;236(11):1192–8.
Paltrinieri S, Gradoni L, Roura X, Zatelli A, Zini E. Laboratory tests for diagnosing and monitoring canine leishmaniasis. Vet Clin Pathol. 2016;45(4):552–78.
Romano A, Inbar E, Debrabant A, Charmoy M, Lawyer P, Ribeiro-Gomes F, et al. Cross-species genetic exchange between visceral and cutaneous strains of Leishmania in the sand fly vector. Proc Natl Acad Sci U S A. 2014;111(47):16808–13.
The authors thank the veterinarian (Oana Gusatoaia) and the staff of the dog shelter who collaborated during field activities. The authors also thank Riccardo Paolo Lia (Department of Veterinary Medicine, University of Bari, Italy) for optical microscopy images used in figures, Viviana Domenica Tarallo (Department of Veterinary Medicine, University of Bari, Italy) for assistance with sand fly morphological identification and serological testing, and Margherita Ceccoli (Department of Veterinary Medicine, University of Bari, Italy) for assisting with laboratory activities.
This study was partially funded by Boehringer-Ingelheim.
Ethics approval and consent to participate
Protocols for collection of dog samples were approved by the ethical committee of the Department of Veterinary Medicine of the University of Bari, Italy (Prot. Uniba 12/20). Protocols for lizard collection and sampling were authorized by the Ministry for Environment, Land and Sea Protection of Italy (Approval Number 0073267/2019), the Societas Herpetologica Italica and the Istituto Superiore per la Protezione e la Ricerca Ambientale (Approval Number 71216).
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The sponsor played no role in the study design, data interpretation or conclusions. Fred Beugnet is a Boehringer-Ingelheim employee. The authors declare that they have no competing interests.
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Mendoza-Roldan, J.A., Latrofa, M.S., Iatta, R. et al. Detection of Leishmania tarentolae in lizards, sand flies and dogs in southern Italy, where Leishmania infantum is endemic: hindrances and opportunities. Parasites Vectors 14, 461 (2021). https://doi.org/10.1186/s13071-021-04973-2
- Canine leishmaniasis
- Leishmania infantum
- Leishmania tarentolae
- Sergentomyia minuta