Skip to main content

Livestock infected with Leishmania spp. in southern Iran

Abstract

Background

The magnitude of the health problems caused by leishmaniasis has been a major driving factor behind the development and implementation of leishmaniasis control programs by the national authorities in Iran, with a priority for health and environmental management. Such programs are not achievable unless all of the factors leading to the infection, including the parasite’s life-cycle, vectors and reservoirs, are recognized. So far in Iran, humans and rodents have been considered the principal reservoirs of Leishmania tropica and Leishmania major, respectively, both associated with cutaneous leishmaniasis (CL), with domestic dogs considered to be the main reservoir for Leishmania infantum, associated with visceral leishmaniasis (VL). The role of other mammals in maintaining the Leishmania parasite has remained unclear. This study aimed to investigate Leishmania infection among livestock in endemic areas of VL and CL in Fars province, southern Iran, using serological and molecular methods.

Methods

Blood samples from 181 clinically healthy livestock, including 49 sheep, 114 goats, 16 cattle and two donkeys, were screened to detect Leishmania DNA and anti-Leishmania antibodies using qPCR (quantitative PCR) and the direct agglutination test (DAT), respectively. Four qPCR-positive samples were amplified using the internal transcribed spacer one (ITS1) primers in conventional PCR and sent for directional sequencing.

Results

Of the 181 livestock tested, 51 (28.2%) were infected with Leishmania, using serological and molecular methods. Anti-Leishmania antibodies were detected in 70 (38.7%) (95% confidence interval [CI]: 31.5–46.2) and Leishmania DNA in 93 (51.4%) (95% CI: 43.9–58.9) livestock. The identified Leishmania spp. were L. infantum and L. major.

Conclusions

The findings of the present study show a relatively high prevalence of Leishmania infection among livestock in endemic areas of the disease, in Fars province, southern Iran. Given the large population of this group of animals and the fact that they live in the vicinity of the main reservoirs of the disease and vectors, it seems that sand flies regularly bite these animals. Further studies are needed to determine the role of livestock in the parasite’s life-cycle and the epidemiology of Leishmania infection.

Graphical Abstract

Background

Leishmaniasis is a vector-borne disease caused by obligate intracellular parasites in the genus Leishmania that can be classified by geographic occurrence into Old World and New World Leishmania species. Old World leishmaniasis can lead to cutaneous leishmaniasis (CL), caused by Leishmania majorL. tropica and L. aethiopica, and visceral leishmaniasis (VL), caused by Leishmania infantum and L. donovani [1]. The WHO estimated that 350 million persons in 98 countries, including Iran, are at the risk of developing leishmaniasis, with an annual incidence of around 1.5 million new cases [2].

Iran is one of the countries included in the WHO Eastern Mediterranean Region, and accounts for 54% of CL cases in that region [3]. Although the incidence of VL cases is lower than that of CL, VL remains a major health concern in the respective endemic areas. In Iran, CL is caused by L. major and L. tropica, whereas VL is caused by L. infantum and, to a lesser extent, L. tropica.

Leishmania infection can be anthroponotic, i.e., transmitted from human to human, or zoonotic, with wild or domestic animals as the reservoir [4]. Mammals that can sustain the population of the infectious agents are called reservoirs and play an essential role in maintaining the life-cycle of the organism and, thus, the continued survival of the infectious agents [5, 6]. In addition to the main reservoirs, some incidental hosts can temporarily maintain the parasite and play a role in transmitting the infection [5]. Determination of the life-cycle of parasites and identification of sand fly vectors and reservoirs play a crucial role in obtaining a better understanding of epidemiology and ultimately applying appropriate strategies to control and eliminate the disease. The magnitude of leishmaniasis-associated problems, a disease which mainly affects poor communities due to the lack of preventive measures, has made the characterization of the above-mentioned issues more necessary. The principal reservoirs of L. tropica and L. major are humans and rodents, respectively [7]; however, the zoonotic transmission of L. tropica has been reported in some regions [8, 9]. Domestic dogs are considered to be the main reservoir for L. infantum, but other mammals have also been shown to be able to sustain the parasite [10, 11]. Previous epidemiological studies have Leishmania infection in horses, goats, cattle and cows in endemic regions of the disease, and some studies have shown that Leishmania vectors (Phlebotomine) bite these groups of animals and feed on their blood [12,13,14,15,16,17]. However, the roles of such mammals in maintaining infection and functioning as reservoirs have not yet been investigated.

To our knowledge, there has been no study in Iran investigating livestock infection caused by Leishmania spp. Therefore, the aim of the present study was to evaluate Leishmania infection in livestock in VL and CL endemic regions of southern Iran.

Methods

Study site and blood sampling

This seroepidemiological and molecular study of infection with Leishmania in livestock was carried out in three villages in Fars province, southern Iran, in VL and CL endemic regions. The study areas are located in a semi-arid region. Two villages, Jaydasht and Do-Ghalat (28°48′ N, 52°38ʹ E), are located in the central district of Firuzabad County, and the third village, Emam-shahr (28°27ʹ N, 53°07ʹ E), is located in the central district of Qir va Karzin County (Fig. 1). According to data obtained from the Iran Veterinary Organization at the time of the study, the livestock population in these three districts was 5711. A minimal sample size of 136 was estimated (95% confidence interval [CI], 5% margin of error), based on studies conducted in other endemic countries with a 10% infection rate. No particular skin manifestation related to leishmaniosis was observed in any of the livestock tested. Blood samples (8 ml) from jugular veins of 181 livestock, consisting of 114 goats, 16 cattle, 49 sheep and two donkeys, were collected in tubes containing K2EDTA and stored in a cold box before being transferred to the laboratory. Plasma and buffy coat were separated from the blood on the same days and stored at − 20 °C until use.

Fig. 1
figure 1

The geographical location of the study on a map of Fars province

Meanwhile, blood samples were collected from 30 goats, 30 sheep and 30 cattle in a non-endemic Leishmania region, Shiraz, to define the direct agglutination test (DAT) cut-off titer and to determine the validity of the qPCR assay. qPCR with kinetoplast DNA (kDNA) primers was used to detect the Leishmania kDNA and quantify the parasite load. In addition, bidirectional sequencing was used to identify the parasite species after performing conventional PCR with internal transcribed spacer 1 (ITS1) primers. In this study, infected livestock was defined as those with positive results in qPCR and DAT.

Direct agglutination test

All plasma samples were assessed for antibodies against Leishmania spp. by DAT, according to Harith et al. [18]. In summary, sera were serially diluted twofold in DAT diluent (0.1 M 2-mercaptoethanol in saline [0.9%]) to make dilutions of 100–102,400. Antigen was dispensed to each well (50 µl/well), and the plates were gently rotated and left overnight at room temperature. The DAT titer cut-off point was determined using samples from non-endemic regions. As a positive control, a confirmed human VL serum was applied.

Quantitative real-time PCR and conventional PCR

Buffy coats digested in 500 µl of lysis buffer (0.5% Tween 20, 0.5% Nonidet P-40, 10 mM NaOH, 10 mM Tris pH 7.2) containing 320 mg proteinase K/ml were incubated at 56 °C overnight. DNA was then extracted using the simplified phenol–chloroform extraction method [19].

The qPCR assay was performed using the TaqMan Gene Expression Assay (Applied Biosystems, Thermo Fisher Scientific, Waltham, MA, USA) with forward and reverse primers directed against kDNA (CTTTTCTGGTCCTCCGGGTAGG and CCACCCGGCCCTATTTTACACCAA, respectively) and FAM-labeled TaqMan probe (TTTTCGCAGAACGCCCCTACCCGC-BHQ1) [20]. The Applied Biosystems 7500 Real-Time PCR System was used for amplification and detection, with the following reaction conditions applied: 2 min at 50 °C, 10 min at 95 °C, followed by a two-step temperature (94 °C and 60 °C) cycling for 45 cycles. The qPCR assays were performed with 5 µl of DNA in a final reaction volume of 25 µl. The standard curve was created using Leishmania DNA extracted from 5 × 106 parasites, with serial dilutions of 5 µl, from 50,000 to 0.005 parasites prepared for each reaction tube. From among the samples that tested positive, we randomly chose 15 samples for conventional PCR using ITS1 primers to amplify the ITS1 gene of Leishmania species. Conventional PCR was done according to Tai et al. [21]. Four PCR products were randomly chosen and sent for bidirectional sequencing using ITS1 primers to confirm the amplified DNA corresponding to Leishmania species.

Statistical analysis

The Spearman correlation test was used to determine a correlation between DAT titer and parasite load, using SPSS version 18 software (SPSS IBM Corp., Armonk, NY, USA). The level of agreement between qPCR and DAT was determined by calculating Cohen’s Kappa (k) and interpreted as follows: negligible (k = 0–0.20), weak (k = 0.21–0.40), moderate (k = 0.41–0.60), good (k = 0.61–0.80) and excellent (k = 0.81–1).

Results

A titer > 1600 was designated as the cut-off based on the results of the samples from non-endemic regions, which was also previously used for a study on livestock by Mukhtar et al. [22]. We detected Leishmania antibody at a titer > 1600 in 38.7% (70/181) (95% CI: 31.5–46.2) of the livestock tested, whereas Leishmania DNA was detected in 51.4% (93/181) (95% CI 43.9–58.9) of the livestock tested. DAT revealed that 11.5, 23.8 and 3.9% of the sheep, goats and cattle, respectively, were seropositive for infection with Leishmania. The titer distribution was: 36 animals with titer 3200, 25 with titer 6400, five with titer 12,800, one with titer 25,600 and three with titer 51,200.

According to the Spearman correlation test analysis, there was a significant relationship between the titer determined with the DAT and parasite load (P < 0.5). Three samples with high titers according to the DAT (51,200) were among those with the highest parasite load. Forty-two qPCR-positive samples which tested negative by the DAT had a low parasite load (1–40 parasites/ml; median: 7 parasites/ml). The level of agreement between the qPCR and DAT results was weak (k = 0.33; P < 0.000).

Leishmania DNA was found in 14.9, 29.3 and 6.6% of the sheep, goats and cattle, respectively, and in one of the two donkey’s buffy coat suspension. Among those samples with positive qPCR results, sample quantification yielded parasite loads ranging from 1 to 3000 parasites/ml (median: 100 parasite/ml). The 15 randomly selected samples with a positive qPCR result were also positive in the conventional PCR analyses using ITS1 primers.

Of the four ITS1-sequenced PCR products, two sequences revealed > 99% sequence identity with the ITS1 sequence of L. infantum with accession numbers MT497974 and AJ634370 (registered in the GenBank database). Of the remaining two products, one had an 86.32% sequence identity with the ITS1 sequence of Leishmania spp. with accession number MT302159, and the second had > 99% sequence identity with the ITS1 sequence of L. major with accession number AY260965 (both registered in the GenBank database). Of these four sequenced PCR products, two were related to cattle and two were related to sheep and goats. These four ITS1 DNA sequences were deposited in the GenBank database with accession numbers OM238068, OM238069, OM218660 and OM218661.

Both qPCR and DAT-positive results were found in 51 cases (28.2%). Tables 1 and 2 present the DAT and qPCR results in terms of livestock type, locations and tests.

Table 1 Direct agglutination test and qPCR results in different livestock from three districts of southern Iran
Table 2 qPCR and DAT results on the samples collected from livestock in southern Iran

Discussion

Firuzabad and Qir va Karzin counties in Fars province are endemic areas where both CL and VL are known to be actively transmitted. In Iran, dogs and rodents are the only confirmed primary reservoirs for VL and CL infections caused by L. infantum and L. major, respectively.

A few studies in Iran have found leishmaniosis in cats using both serological and molecular approaches, while the relevance of cats in the zoonotic transmission is still controversial [23]. In Iran’s VL and CL endemic regions, no attempts have been made to identify Leishmania infection in livestock that come into close contact with the principal reservoir host and graze near resting habitats of Leishmania-infected sand flies. In the present study, we found that 28.2% of the livestock tested were positive for Leishmania. It is unclear how these animals became infected, but given the importance of phlebotomine sand flies in the Leishmania infection cycle, it is likely that bites from infectious sand flies were the only route of transmission. Since dogs and rodents are the main reservoirs for VL and CL, and given the large population of rodents in these areas, as well as the presence of guard dogs for domestic herds, and assuming no preference for blood meal source for female sand flies, such livestock must be bitten by sand flies frequently and become easily infected.

Yard et al. found that the majority of blood meals consumed by sand flies in the VL endemic region of northeastern Ethiopia were from donkeys (33.9%), followed by cows (24.2%), humans (17.6%), dogs (11.8%) and goats or sheep (8.6%) [17]. Abbate et al. reported that the most prevalent blood sources observed in sand flies in the Leishmania endemic area of Sicily came from wild rabbits (n = 28), humans (n = 24), goats (n = 16), horses (n = 13), pigs (n = 9), dogs (n = 9), chickens (n = 3), cows (n = 3), cats (n = 1), donkeys (n = 1) and rats (n = 1) [24]. A remarkably high bovine blood feed (92%) was reported by Gebre-Michael et al. in their analyses of blood meals of 273 fresh-fed Phlebotomus orientalis females from northwest Ethiopia [25].

It should be noted that in these three studies, the authors noted that a considerable percentage of phlebotomine blood meals came from livestock.

An animal is considered to be a reservoir of Leishmania infection when: (i) it is long-lived and is present in relative abundance; (ii) it is responsible for the long-term maintenance of parasites; (iii) it remains infected and apparently healthy for a long time; (iv) it bears high levels of parasites in the superficial skin vessels for availability to sand flies; and (v) sand flies depend on feeding on them [5, 6].

In VL, the parasites spread through skin macrophages and affect organs such as lymph nodes, bone marrow and spleen. Therefore, biopsies of these tissues for direct parasite detection are required to confirm parasite multiplication and survival in these organs. It is also recommended that samples be taken from the infected animals at different intervals to see how long they can stay infected. Another option is to look into the blood meal source of Leishmania-infected sand flies in these regions. Consequently, consideration of these mammals as reservoirs demands adequate and consistent evidence. Performing xenodiagnoses on cases with high parasite burdens is crucial to proving that these mammals are infectious to laboratory-reared sand flies.

To determine whether the same parasite clone is present, the Leishmania ITS1 DNA sequences of VL and CL patients in this study area should be compared to that extracted from infected dogs, rodents, sand flies and livestock. Several previous studies in Leishmania endemic regions around the world have reported Leishmania infection in livestock, consistent with present study findings, although these studies differ in terms of sample size, diagnostic method applied, types of samples, presence of heterogeneous parasite populations and immune responses to the Leishmania parasite [12,13,14, 16, 26]. In the present study, some samples were positive by both molecular and serological methods (51/181; 28.2%), while others were positive by only one method. Our data show a direct correlation between parasite load and antibody titer determined by DAT; accordingly, some qPCR-positive samples with a low parasite load were negative by DAT (42/181; 23.2%), suggesting the onset of infection. According to a recent study, qPCR has a higher sensitivity than serological tests in identifying asymptomatic leishmaniasis in whole blood; however, the qPCR test could not detect all infected cases [27, 28]. Although it is unclear whether these results might be extended to the diagnosis of the infection with Leishmania in livestock, a combination of several methods may be the best strategy to diagnose Leishmania infection in livestock [28].

DAT results were positive in 38.7% (70/181) of the livestock tested, with approximately 10.5% (19/181) of the livestock being only positive by DAT, and the rest positive by both techniques. The antibodies detected in this 10.5% of the livestock may be associated with an old infection; nevertheless, the antigenic similarity between Leishmania and Trypanosoma infecting livestock should not be overlooked [29].

More than half (54%) of CL cases in WHO Eastern Mediterranean Region countries occur in Iran, with a quarter of these cases occurring in Fars province, with the predominant form of zoonotic CL caused by L. major [3, 30]In the current investigation, the sequence of one of the randomly selected cases was L. major. The high percentage of infected livestock in this study is significant, emphasizing the need to pay attention to such animals for their ability to maintain and transmit Leishmania parasites.

Given the data above and the lack of clinical manifestations in the studied livestock and their large population, these animals may serve as a potential Leishmania reservoir. However, further research is needed to determine not only how these animals are infected and but also the feeding behavior of the sand fly in order to clarify the sand fly’s role in the epidemiology of Leishmania infection.

Availability of data and materials

Data supporting the conclusions of this article are included within the article. The raw data obtained during the present study are available upon request.

Abbreviations

CL:

Cutaneous leishmaniasis

DAT:

Direct agglutination test

ITS1:

Internal transcribed spacer 1

VL:

Visceral leishmaniasis

References

  1. WHO. The leishmaniasis: report of a WHO expert committee. World Health Organ Tech Rep Ser. 1984;701:1–140.

    Google Scholar 

  2. WHO. Working to overcome the global impact of neglected tropical diseases: first WHO report on neglected tropical diseases. 2010. https://apps.who.int/iris/handle/10665/44440. Accessed 22 May 2022.

  3. Postigo JAR. Leishmaniasis in the World Health Organization Eastern Mediterranean Region. Int J Antimicrob. 2010;36:S62–5.

    CAS  Article  Google Scholar 

  4. Desjeux P. Leishmaniasis: public health aspects and control. Clin Dermatol. 1996;14:417–23.

    CAS  Article  Google Scholar 

  5. Ashford RW. Leishmaniasis reservoirs and their significance in control. Clin Dermatol. 1996;14:523–32.

    CAS  Article  Google Scholar 

  6. WHO Expert Committee on the Control of the Leishmaniases & WHO. Control of the leishmaniases: report of a meeting of the WHO Expert Commitee on the Control of Leishmaniases, Geneva, 22–26 Mar 2010. 2010. https://apps.who.int/iris/handle/10665/44412. Accessed 22 May 2022.

  7. González U, Pinart M, Sinclair D, Firooz A, Enk C, Vélez ID, et al. Vector and reservoir control for preventing leishmaniasis. Cochrane Database Syst Rev. 2015;2015(8):CD008736. https://doi.org/10.1002/14651858.CD008736.pub2.

  8. Kassahun A, Sadlova J, Dvorak V, Kostalova T, Rohousova I, Frynta D, et al. Detection of Leishmania Donovani and L. tropica in Ethiopian wild rodents. Acta Trop. 2015;145:39–44.

    Article  Google Scholar 

  9. Svobodova M, Votypka J, Peckova J, Dvorak V, Nasereddin A, Baneth G, et al. Distinct transmission cycles of Leishmania tropica in 2 adjacent foci, Northern Israel. Emerg Infect Dis. 2006;12:1860.

    CAS  Article  Google Scholar 

  10. Pennisi M-G, Cardoso L, Baneth G, Bourdeau P, Koutinas A, Miró G, et al. LeishVet update and recommendations on feline leishmaniosis. Parasit Vectors. 2015;8:1–18.

    Article  Google Scholar 

  11. Alcover MM, Ribas A, Guillén MC, Berenguer D, Tomás-Pérez M, Riera C, et al. Wild mammals as potential silent reservoirs of Leishmania infantum in a Mediterranean area. Prev Vet Med. 2020;175:104874.

    Article  Google Scholar 

  12. Bhattarai NR, Van der Auwera G, Rijal S, Picado A, Speybroeck N, Khanal B, et al. Domestic animals and epidemiology of visceral leishmaniasis, Nepal. Emerg Infect Dis. 2010;16:231.

    Article  Google Scholar 

  13. Kenubih A, Dagnachew S, Almaw G, Abebe T, Takele Y, Hailu A, et al. Preliminary survey of domestic animal visceral leishmaniasis and risk factors in northwest Ethiopia. Trop Med Int Health. 2015;20:205–10.

    Article  Google Scholar 

  14. Labony SS, Begum N, Rima UK, Chowdhury G, Hossain M, Habib M, et al. Apply traditional and molecular protocols for the detection of carrier state of visceral leishmaniasis in black Bengal goat. J Agric Vet Sci. 2014;7:13–8.

    Google Scholar 

  15. Rohousova I, Talmi-Frank D, Kostalova T, Polanska N, Lestinova T, Kassahun A, et al. Exposure to Leishmania spp. and sand flies in domestic animals in northwestern Ethiopia. Parasit Vectors. 2015;8:1–10.

    CAS  Article  Google Scholar 

  16. Singh N, Mishra J, Singh R, Singh S. Animal reservoirs of visceral leishmaniasis in India. J Parasitol. 2013;99:64–7.

    Article  Google Scholar 

  17. Yared S, Gebresilassie A, Abbasi I, Aklilu E, Kirstein OD, Balkew M, et al. A molecular analysis of sand fly blood meals in a visceral leishmaniasis endemic region of northwestern Ethiopia reveals a complex host–vector system. Heliyon. 2019;5:e02132.

    Article  Google Scholar 

  18. Harith AE, Kolk AHJ, Leeuwenburg J, Muigai R, Huigen E, Jelsma T, et al. Improvement of a direct agglutination test for field studies of visceral leishmaniasis. J Clin Microbiol. 1988;26:1321–5.

    Article  Google Scholar 

  19. Sambrook J, Fritsch EF, Maniatis T. Preparation and analysis of eukaryotic genomic DNA. Molecular cloning: a laboratory manual. 3rd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, Academic Press. 1989. p. 6.8–6.9.

  20. Mary C, Faraut F, Lascombe L, Dumon H. Quantification of Leishmania infantum DNA by a real-time PCR assay with high sensitivity. J Clin Microbiol. 2004;42:5249–55.

    CAS  Article  Google Scholar 

  21. El Tai N, Osman O, El Fari M, Presber W, Schönian G. Genetic heterogeneity of ribosomal internal transcribed spacer in clinical samples of Leishmania donovani spotted on filter paper as revealed by single-strand conformation polymorphisms and sequencing. Trans R Soc Trop Med Hyg. 2000;94:575–9.

    Article  Google Scholar 

  22. Mukhtar MM, Sharief AH, el Saffi SH, Harith AE, Higazzi TB, Adam AM, et al. Detection of antibodies to Leishmania donovani in animals in a kala-azar endemic region in eastern Sudan: a preliminary report. Trans R Soc Trop Med Hyg. 2000;94:33–6.

    CAS  Article  Google Scholar 

  23. Hatam GR, Adnani SJ, Asgari Q, Fallah E, Motazedian MH, Sadjjadi SM, et al. First report of natural infection in cats with Leishmania infantum in Iran. Vector Borne Zoonotic Dis. 2010;10:313–6.

    Article  Google Scholar 

  24. 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:e0229536.

    CAS  Article  Google Scholar 

  25. Gebre-Michael T, Balkew M, Berhe N, Hailu A, Mekonnen Y. Further studies on the phlebotomine sandflies of the kala-azar endemic lowlands of Humera-Metema (northwest Ethiopia) with observations on their natural blood meal sources. Parasit Vectors. 2010;3:1–7.

    Article  Google Scholar 

  26. Mutinga MJ, Mutero C, Ngindu A, Amimo F. The isolation of leishmanial parasites from domestic goats and wild hosts in Kenya and the possible role of goats as reservoirs of leishmaniases. Int J Trop Insect Sci. 1988;9:347–9.

    Article  Google Scholar 

  27. Alborzi A, Pourabbas B, Shahian F, Mardaneh J, Pouladfar GR, Ziyaeyan M. Detection of Leishmania infantum kinetoplast DNA in the whole blood of asymptomatic individuals by PCR-ELISA and comparison with other infection markers in endemic areas, southern Iran. Am J Trop Med. 2008;79:839–42.

    Article  Google Scholar 

  28. Rezaei Z, Pourabbas B, Asaei S, Kühne V, Sepehrpour S, Pouladfar G, et al. Pediatric visceral leishmaniasis: a retrospective study to propose the diagnostic tests algorithm in southern Iran. Parasitol Res. 2021;120:1447–53.

    Article  Google Scholar 

  29. Camargo ME, Rebonato C. Cross-reactivity in fluorescence tests for Trypanosoma and Leishmania antibodies. A simple inhibition procedure to ensure specific results. Am J Trop Med. 1969;18:500–5.

    CAS  Article  Google Scholar 

  30. Norouzinezhad F, Ghaffari F, Norouzinejad A, Kaveh F, Gouya MM. Cutaneous leishmaniasis in Iran: results from an epidemiological study in urban and rural provinces. Asian Pac J Trop Biomed. 2016;6:614–9.

    Article  Google Scholar 

Download references

Acknowledgements

Hassan Khajehei is thanked for copy editing the manuscript.

Funding

This project was financially supported by Professor Alborzi Clinical Microbiology Research Center, Shiraz University of Medical Sciences, Iran.

Author information

Authors and Affiliations

Authors

Contributions

AA, ZR, and BP conceived and designed the experiments. SA, SS, PP, SAM and SAK performed the experiments. ZR, analyzed the data. ZR and BP wrote the paper. All authors read and approved the final manuscript.

Corresponding author

Correspondence to Bahman Pourabbas.

Ethics declarations

Ethics approval and consent to participate

This study received ethical approvals from the Ethics Committee of the Professor Alborzi Clinical Microbiology Research Center (no. 1/98/30.11.97- 2019) and the Ethics Committees of Shiraz University of Medical Sciences (SUMS), Shiraz, Iran (IR.SUMS.REC.1400.856).

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Rezaei, Z., Pourabbas, B., Asaei, S. et al. Livestock infected with Leishmania spp. in southern Iran. Parasites Vectors 15, 215 (2022). https://doi.org/10.1186/s13071-022-05313-8

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s13071-022-05313-8

Keywords

  • Iran
  • Leishmaniasis
  • Leishmania infection
  • Livestock
  • Reservoirs