
==== Front
Parasit Vectors
Parasit Vectors
Parasites & Vectors
1756-3305
BioMed Central London

39300564
6487
10.1186/s13071-024-06487-z
Research
Seasonal dynamics, Leishmania diversity, and nanopore-based metabarcoding of blood meal origins in Culicoides spp. in the newly emerging focus of leishmaniasis in Northern Thailand
Promrangsee Chulaluk 1
Sriswasdi Sira 234
Sunantaraporn Sakone 56
Savigamin Chatuthanai 6
Pataradool Thanapat 6
Sricharoensuk Chatchapon 6
Boonserm Rungfar 56
Ampol Rinnara 56
Pruenglampoo Pitchayaporn 7
Mungthin Mathirut 8
Schmidt-Chanasit Jonas 910
Siriyasatien Padet 56
Preativatanyou Kanok kanok.pr@chula.ac.th

56
1 https://ror.org/028wp3y58 grid.7922.e 0000 0001 0244 7875 Interdisciplinary Program of Biomedical Sciences, Graduate School, Chulalongkorn University, Bangkok, Thailand
2 https://ror.org/028wp3y58 grid.7922.e 0000 0001 0244 7875 Center of Excellence in Computational Molecular Biology, Faculty of Medicine, Chulalongkorn University, Bangkok, Thailand
3 https://ror.org/028wp3y58 grid.7922.e 0000 0001 0244 7875 Center for Artificial Intelligence in Medicine, Research Affairs, Faculty of Medicine, Chulalongkorn University, Bangkok, Thailand
4 https://ror.org/057zh3y96 grid.26999.3d 0000 0001 2169 1048 Department of Integrated Biosciences, Graduate School of Frontier Sciences, The University of Tokyo, Kashiwa, Chiba Japan
5 https://ror.org/028wp3y58 grid.7922.e 0000 0001 0244 7875 Center of Excellence in Vector Biology and Vector-Borne Disease, Chulalongkorn University, Bangkok, Thailand
6 https://ror.org/028wp3y58 grid.7922.e 0000 0001 0244 7875 Department of Parasitology, Faculty of Medicine, Chulalongkorn University, Bangkok, Thailand
7 https://ror.org/03rn0z073 grid.415836.d 0000 0004 0576 2573 Division of Medical Technical and Academic Affairs, Department of Medical Services, Ministry of Public Health, Nonthaburi, Thailand
8 https://ror.org/04md5yc36 0000 0004 0576 1116 Department of Parasitology, Phramongkutklao College of Medicine, Bangkok, Thailand
9 https://ror.org/01evwfd48 grid.424065.1 0000 0001 0701 3136 Bernhard-Nocht-Institute for Tropical Medicine, Bernhard-Nocht-Str. 74, Hamburg, Germany
10 https://ror.org/00g30e956 grid.9026.d 0000 0001 2287 2617 Faculty of Mathematics, Informatics and Natural Sciences, Universität Hamburg, Hamburg, Germany
19 9 2024
19 9 2024
2024
17 40026 7 2024
4 9 2024
© The Author(s) 2024
2024
https://creativecommons.org/licenses/by/4.0/ 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.
Background

Clinical cases of leishmaniasis caused by Leishmania (Mundinia) parasites have been increasingly reported in Southeast Asia, particularly Thailand. Recent evidence has shown that Leishmania (Mundinia) parasites successfully developed into infective metacyclic promastigotes in Culicoides biting midges, strongly supporting their putative role in disease transmission. However, Culicoides diversity, host preference, and Leishmania prevalence in endemic areas remain largely unknown.

Methods

We investigated the seasonal dynamics, infection prevalence, and blood meal identification of Culicoides collected from the emerging focus of visceral leishmaniasis in Lampang Province, Northern Thailand, during 2021–2023. Midge samples were molecularly screened for Leishmania using SSU rRNA-qPCR and ITS1-PCR, followed by Sanger plasmid sequencing, and parasite haplotype diversity was analyzed. Host blood meal origins were comparatively identified using host-specific Cytb-PCRs and a nanopore-based metabarcoding approach.

Results

A total of 501 parous and gravid females and 46 blood-engorged ones belonging to at least 17 species of five subgenera (Remmia, Trithecoides, Avaritia, Hoffmania, and Meijerehelea) and two species groups (Shortti and Calvipalpis) were collected with temporal differences in abundance. Leishmania was detected by SSU rRNA-qPCR in 31 samples of at least 11 midge species, consisting of Culicoides oxystoma, C. guttifer, C. orientalis, C. mahasarakhamense, C (Trithecoides) spp., C. innoxius, C. shortti, C. arakawae, C. sumatrae, C. actoni, and C. fulvus, with the overall infection prevalence of 5.7%. The latter six species represent the new records as putative leishmaniasis vectors in Northern Thailand. The ITS1-PCR and plasmid sequencing revealed that Leishmania martiniquensis was predominantly identified in all qPCR-positive species, whereas L. orientalis was identified only in three C. oxystoma samples. The most dominant haplotype of L. martiniquensis in Thailand was genetically intermixed with those from other geographical regions, confirming its globalization. Neutrality test statistics were also significantly negative on regional and country-wide scales, suggesting rapid population expansion or selective sweeps. Nanopore-based blood meal analysis revealed that most Culicoides species are mammalophilic, with peridomestic and wild mammals (cow, pig, deer, and goat-like species) and humans as hosts, while C. guttifer and C. mahasarakhamense fed preferentially on chickens.

Conclusions

This study revealed seasonal dynamics and sympatric circulation of L. martiniquensis and L. orientalis in different species of Culicoides. Evidence of human blood feeding was also demonstrated, implicating Culicoides as putative vectors of human leishmaniasis in endemic areas. Further research is therefore urgently needed to develop vector control strategies and assess the infection status of their reservoir hosts to effectively minimize disease transmission.

Graphical Abstract

Supplementary Information

The online version contains supplementary material available at 10.1186/s13071-024-06487-z.

Keywords

Culicoides biting midges
Leishmania
Mundinia
Nanopore metabarcoding
Host preference
Killick-Kendrick’s criteria
Northern Thailand
the Ratchadapiseksompote Fund, Graduate Affairs, Faculty of Medicine, Chulalongkorn UniversityGA68/001 Preativatanyou Kanok issue-copyright-statement© BioMed Central Ltd., part of Springer Nature 2024
==== Body
pmcBackground

Autochthonous leishmaniasis is currently considered an important public health problem in Southeast Asia with Thailand being an endemic hotspot. Since 1996, an increasing number of clinical cases have been reported, particularly in the northern and southern provinces of the country [1]. This emerging disease has been proven to be caused by two Leishmania species of the newly classified subgenus Mundinia, namely Leishmania martiniquensis and L. orientalis [1–5]. Traditionally, leishmaniasis was known to be transmitted by phlebotomine sand flies [6]. Based on the positive detection of Leishmania DNA, several phlebotomine species of the genera Sergentomyia [7–9], Grassomyia [10], and Phlebotomus [11] were proposed as potential vectors of these two Leishmania (Mundinia) species. However, recent evidence indicates that several Leishmania (Mundinia) parasites could not successfully develop to the infective stage within the midgut of the sand fly [12–14]. In addition, the positive detection rates for Leishmania (Mundinia) species are relatively low in field-caught sand flies, suggesting that these emerging Leishmania (Mundinia) parasites may be transmitted by other blood-sucking dipteran vectors than sand flies [7, 9–11, 15].

In addition to sand flies, Culicoides biting midges are globally distributed hematophagous insects of the family Ceratopogonidae, order Diptera. Of medical and veterinary importance, species of this insect genus have been implicated as vectors of several human, domestic, and wildlife pathogens. They are involved in the transmission of several animal arboviruses, including African horse sickness, bluetongue, epizootic hemorrhagic disease, Oropouche, vesicular stomatitis, and Schmallenberg viruses [16]. Culicoides species have also been incriminated as vectors of protozoan parasites infecting mammals and birds, including Hepatocystis, Haemoproteus, Leucocytozoon, and Trypanosoma, as well as several species of filarial nematodes [17–20].

It has been previously noted that several Leishmania species, including Leishmania (Leishmania) amazonensis [21], L. (L.) infantum [22], L. (L.) mexicana [23], and L. (Viannia) braziliensis [21], have been molecularly detected in Culicoides spp. collected from different geographical origins. Nevertheless, the metacyclic development of these Leishmania species has never been demonstrated in dissections of natural Culicoides species or in laboratory experiments, which is an important criterion for confirming their vector competence [12–14, 24, 25]. However, recent experimental infections have shown that several strains of L. martiniquensis and L. orientalis can complete metacyclic development in Culicoides sonorensis and be successfully transmitted to mice [12, 14]. It is therefore highly likely that Culicoides biting midges play a role in the transmission of autochthonous leishmaniasis caused by these two Leishmania (Mundinia) species in endemic areas of Thailand.

Positive detection of two autochthonous Leishmania species, L. martiniquensis and L. orientalis, in certain Culicoides species has been previously reported in endemic areas of Thailand. In Northern Thailand, Culicoides mahasarakhamense has been identified as the predominant vector [26, 27], while in Southern Thailand, C. peregrinus and C. oxystoma are the main putative vectors [28]. However, more than 168 Culicoides species from different ecotypes have been described in Southeast Asian countries, and many of them feed on mammals, which may serve as parasite reservoirs [29]. Accordingly, it can be speculated that Leishmania parasites may exploit more Culicoides species as hosts than previously reported, with spatiotemporal differences in the vector species spectrum between leishmaniasis-affected localities in different geographical regions of Thailand.

Although several experimental and field studies have implicated Culicoides as putative vectors of autochthonous leishmaniasis [12, 14, 26–28], evidence of their feeding behaviors on human blood has never been demonstrated, particularly in endemic areas of the country. Therefore, the incrimination of Culicoides as true natural vectors of human leishmaniasis according to the Killick-Kendrick criteria [30] cannot be strengthened. Several attempts were made using Sanger sequencing to analyze blood meals in engorged Culicoides collected near human dwellings in leishmaniasis-endemic areas, but it was found that all specimens fed only on animals with domestic and peridomestic environments, without evidence of human blood [28, 31]. We speculated that these results were likely biased because Sanger sequencing can only generate a single chromatogram, representing a single blood meal source, and is therefore not suitable for characterizing multiple blood meal sources in a single specimen. Additionally, information on the composition of Culicoides with seasonal abundance, prevalence of infection, and sympatric occurrence of Leishmania parasites in the affected communities remains limited.

In the present study, we comparatively investigated the seasonal dynamics of Culicoides biting midges and the infection prevalence and genetic diversity of Leishmania in Culicoides samples collected from the residence of the leishmaniasis index case in Lampang Province, an emerging focus of visceral leishmaniasis in Northern Thailand, from 2021 to 2023. In addition, we demonstrated the application of a nanopore-based metabarcoding strategy to provide high-throughput data for the unbiased identification of multiple host species. Novel insights from this study will help us better understand the role of Culicoides in leishmaniasis transmission and facilitate effective vector management and control to interrupt disease transmission, especially in endemic areas of northern Thailand.

Methods

Investigation area, biting midge collection, and morphological identification

Culicoides biting midges were collected in Wang Nuea District, Lampang Province, Northern Thailand (19° 10′ 27.5″ N, 99° 38′ 53.0″ E), from November 2021 to May 2023 (Fig. 1). The midge collection was performed within a 50-m radius around the residence of the index case who had been previously diagnosed with visceral leishmaniasis. The Centers for Disease Control and Prevention miniature UV light traps were situated 1 m above the ground in different places near the household area and neighboring cattle sheds. The traps were installed from dusk to dawn for 3 consecutive days. All insect specimens collected the next day would be immobilized by being knocked out in the freezer for 30 min. At the field site, the collection was then inspected under a stereomicroscope (EZ4 HD, Leica, Germany) to sort female Culicoides individuals from males and other insect species, according to distinct morphological structures, i.e., non-plumose antennae and wing venation patterns. In this study, nulliparous females, which generally predominated in the traps and had never been exposed to an infectious blood meal, were discarded from the female collection. Only non-engorged (parous and gravid) and blood-engorged ones, which had fed on hosts and entered the gonotrophic cycle, were picked up for downstream analysis. All midge specimens were then morphologically identified using the illustrated keys to Culicoides of Southeast Asia formerly described by Wirth and Hubert [29] and the formal description of a new species, C. mahasarakhamense, recorded by Pramual et al. [32]. Then, the midge specimens were frozen at – 80 °C before genomic DNA isolation.Fig. 1 Map and geographic coordinates of Culicoides trapping site in Wang Nuea District, Lampang Province, Northern Thailand. The light traps were installed near the cattle pen and chicken coop within the patient’s housing area. The map of Thailand was obtained from the public domain (https://d-maps.com). The satellite image was modified from the Google Earth website (https://earth.google.com/web/search/Thailand)

Genomic DNA isolation from Culicoides samples

To preserve the morphological structure of the samples, each Culicoides individual was extracted for total genomic DNA (gDNA) without sample homogenization using a non-destructive protocol previously described by Santos et al. [33]. Briefly, each sample was digested in 200 µl lysis buffer containing Proteinase K and incubated at 50 °C overnight. The lysate was then purified using the magnetic bead-based system with the GENTi™ Advanced Genomic DNA Extraction Kit (GeneAll®, Seoul, South Korea). The quality and concentration of gDNA were assessed using the NanoDrop 2000c spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). The gDNA samples obtained were stored at – 20 °C until use. Post-extracted insect remains were stored in 70% ethanol at room temperature for further species reaffirmation.

Initial screening of Leishmania DNA by SSU rRNA-qPCR

All extracted gDNA samples were first tested for the presence of Leishmania DNA using TaqMan quantitative PCR (qPCR) targeting the conserved region of the small subunit ribosomal RNA (SSU rRNA) gene of all Leishmania species, as previously described [34]. The SSU rRNA-qPCR reactions were performed using a forward primer (5′-CCAAAGTGTGGAGATCGAAG-3′), a reverse primer (5′-GGCCGGTAAAGGCCGAATAG-3′), and a TaqMan probe (5′-6FAM-ACCATTGTAGTCCACACTGC-3′-MGB-NFQ). Reaction components were prepared in a total volume of 20 µl containing 10 µl TaqMan™ Fast Advanced Master Mix (Thermo Scientific, Waltham, MA, USA), 0.5 µM each of 10 µM primers, 0.5 µl 10 µM probe, and 2 µl gDNA. All reactions were performed on the QuantStudio™ 5 Real-Time PCR System (Applied Biosystems, CA, USA) with the following thermal conditions: initial denaturation at 95 °C, 2 min and 45 cycles of denaturation at 95 °C, 15 s and annealing/extension at 60 °C, 15 s. A Ct value < 40 was considered positive.

Conventional LeishmaniaITS1-PCR

The gDNA samples with positive qPCR results were used as templates for conventional PCR targeting all Leishmania species' internal transcribed spacer-1 (ITS1) region. ITS1-PCR reactions were performed using forward primer LITSR2 (5’-CTGGATCATTTTCCGATGATT-3’) and reverse primer L5.8Sinner (5’-GTTATGTGAGCCGTTATCC-3’) to generate an amplicon of approximately 272–280 bp [35]. Reaction components consisted of 4 µl gDNA, 1.5 µl each of 10 µM primers, 25 µl 2X KAPA HiFi HotStart ReadyMix (Roche, Basel, Switzerland), and nuclease-free water added to a final volume of 50 µl. Thermal conditions included initial denaturation at 95 °C, 5 min, followed by 35 cycles of denaturation at 98 °C, 30 s, annealing at 53 °C, 30 s, extension at 72 °C, 30 s, and final extension at 72 °C, 10 min. Amplification products were verified on 1.5% (w/v) agarose gel electrophoresis stained with ethidium bromide and then visualized using the Gel Doc XR + Gel Documentation System (Bio-Rad, Hercules, CA, USA).

Parasite identification by Sanger plasmid DNA sequencing

All Leishmania ITS1 amplicons were ligated into the pGEM® T-Easy cloning vector (Promega Corp., Madison, WI, USA) according to the manufacturer's specifications. The ligation products were then chemically transformed into competent Escherichia coli strain DH5α cells. Transformants were transferred onto the Luria Bertani (LB) agar plates containing ampicillin, IPTG, and X-Gal for blue/white colony selection. Five white colonies suspected of harboring the recombinant plasmids were confirmed by PCR and subsequently cultured overnight at 37 °C in LB broth medium containing ampicillin. Chimeric plasmids were extracted using the Exprep™ Plasmid SV DNA Purification Kit (GeneAll®, Seoul, Korea) and sequenced using T7 promoter primer by Macrogen, Inc. (Seoul, Korea). To identify the parasite species, the ITS1 sequences obtained were compared with GenBank references in the nucleotide collection (nr/nt) database using BLASTn (http://blast.ncbi.nlm.nih.gov/Blast.cgi) optimized for highly similar sequences.

Haplotype network analysis

The genetic variability of L. martiniquensis and L. orientalis ITS1 sequences identified in this study and those from GenBank was investigated by haplotype network analysis based on single nucleotide variation and small insertions and deletions (indels). A minimum-spanning network was constructed in RStudio version 2024.04.2+764 [36]. Briefly, the ITS1 sequences in FASTA format were converted into a DNA binary file required for network construction using the package “adegenet” version 2.1.10 [37]. The package “pegas” version 1.3 was then used to create the haplotype network from a list of DNA sequences in DNA binary format [38]. DNA sequences were compared, and each unique sequence was assigned to a specific haplotype using the 'haplotype' function in the pegas package. Genetic diversity statistics, including the number of haplotypes, haplotype diversity (Hd), nucleotide diversity (π), average number of nucleotide differences (k), and Tajima’s D statistic, were calculated using the package “pegas” version 1.3. Fu and Li’s D statistics were calculated using the package “PopGenome” version 2.7.5 [39]. P-values < 0.05 were considered statistically significant.

PCR amplification of mammalian and avian cytochrome b regions

The gDNA extracted from engorged specimens was used as a template for PCR targeting the vertebrate cytochrome b (Cytb) region for mammalian and avian host identification as described elsewhere. For mammalian hosts, multiplex PCR reactions were set up in a total volume of 25 µl consisting of 3 µl gDNA template, 0.9 µl 10 µM forward primer (Human741F, Cow121F, Dog368F, Pig573F), and universal reverse primer (UNREV1025) [40], 12.5 µl 2X KAPA HiFi HotStart ReadyMix, and 5 µl nuclease-free water. Additionally, singleplex PCR reactions were performed for the avian host using L15557 and H16065 primers [41]. Thermal conditions included initial denaturation at 95 °C, 5 min; 35 cycles of denaturation at 95 °C, 1 min, annealing at 58 °C (for mammals) and 50 °C (for avian), 1 min, extension at 72 °C, 1 min, and final extension at 72 °C, 7 min. PCR products were verified by 1.5% (w/v) agarose gel electrophoresis with staining and visualization as previously described. The expected amplicon sizes were 334, 561, 680, 453, and 508 bp for human, bovine, canine, porcine, and avian, respectively.

Vertebrate COI-PCR and MinION® amplicon sequencing

To identify additional hosts not detected by Cytb-PCR, nanopore amplicon sequencing was performed on all engorged samples. The partial cytochrome c oxidase subunit I (COI) gene of approximately 395 bp was amplified using VertCOI_7194_F (5’-CGMATRAAYAAYATRAGCTTCTGAY-3’) and Mod_RepCOI_R (5’-TTCDGGRTGNCCRAARAATCA-3’) [42]. The amplification reactions were set up in a total volume of 25 µl consisting of 3 µl gDNA template, 0.75 µl of each 10 µM primer, and 12.5 µl 2X KAPA HiFi HotStart ReadyMix. Thermal conditions were programmed as follows: 95 °C for 3 min, 40 cycles of denaturation at 95 °C, 40 s, annealing at 48.5 °C, 30 s, extension at 72 °C, 1 min, and final extension at 72 °C for 7 min. Each amplicon sample was then purified using Agencourt AMPure XP beads (Beckman Coulter, CA, USA).

For DNA library preparation, 200 fmol purified amplicon per sample was end-repaired using the NEBNext Ultra II End Repair/dA-Tailing Module (New England Biolabs, MA, USA). End-prepped DNA samples were ligated with native barcodes provided in the Native Barcoding Kit 96 V14 (Cat. No. SQK-NBD114.96, Oxford Nanopore Technologies, Didcot, UK) using NEB Blunt/TA Ligase Master Mix (New England Biolabs). For multiplexing, barcoded samples were pooled and ligated to the sequencing adaptor using the NEBNext Quick Ligation Module (New England Biolabs). After removing the excess adaptor, 50 fmol of the final preparation was loaded into the MinION® R10.4.1 flow cell for 10 h. The Dorado version 7.3.11 was used for super-accuracy base calling using the dna_r10.4.1_e8.2_400bps_sup@v.4.3.0 model, demultiplexing, and adaptor-barcode trimming. Sequenced reads with Q scores < 20 were filtered out using Chopper version 0.7.0 [43], and reads between 300–500 bp were analyzed. Reads of similar sequence and length were clustered using amplicon_sorter.py (version 2024_02_20) [44] and then error-polished with raw reads using Medaka version 1.11.3 to generate all accurate consensus sequences for each sample. For the taxonomic assignment, the obtained consensus sequences of each sample were compared to GenBank references using BLASTn optimized for highly similar sequences.

Results

Index case description

On 11 January 2021, a 60-year-old male living in Wang Nuea District, Lampang Province, presented with a 1-month history of fatigue and leg myalgia. Physical examination revealed markedly pale conjunctiva, consistent with anemia. Hematological investigations revealed pancytopenia with a hemoglobin level of 5.0 g/dl, hematocrit of 16%, white blood cell count of 2040 cells/mm3 (neutrophil, 38%; lymphocyte, 56%; monocyte, 3%; eosinophil, 2%; basophil, 1%), and platelet count of 72,000 platelets/mm3. His HIV serology was negative. He then received several blood transfusions without significant improvement in his symptoms, and huge splenomegaly developed. In August 2021, a bone marrow biopsy showed several macrophages full of intracellular amastigotes. ITS1-PCR and Sanger sequencing confirmed the final diagnosis of visceral leishmaniasis caused by L. martiniquensis. The DNA sequence obtained was submitted to GenBank under accession no. OR917763. The patient improved clinically following a 1-week course of intravenous amphotericin B (1 mg/kg/day) with seven doses of filgrastim (300 mcg/day) by subcutaneous injection every 2 days.

Seasonal abundance of Culicoides species

Culicoides biting midges were caught three times around the patient’s household in November 2021, March 2022, and May 2023. In total, our collection consisted of 501 non-engorged (parous and gravid) females and 51 blood-engorged females caught only in November 2021. All Culicoides samples were taxonomically identified as at least 17 different species belonging to five subgenera, namely Remmia, Trithecoides, Avaritia, Hoffmania, Meijerehelea, and two species groups, namely Shortti and Calvipalpis, mainly based on the characteristic wing pigmentation patterns (Table 1, Fig. 2). With temporal variability, C. (Trithecoides) spp. were prevalent in November 2021, whereas C. innoxius and C. oxystoma were the most abundant species in March 2022 and May 2023, respectively. Only a single specimen of Culicoides imicola, C. huffi, and C. liui were found sporadically during these three collection periods. The three most abundant species of the entire collection were C. oxystoma, C. (Trithecoides) spp., and C. shortti, representing 29.3%, 15.5%, and 11.9% of all Culicoides midges, respectively.Table 1 Species diversity with temporal and overall variability in the relative abundance of Culicoides biting midges collected from the vicinity of the patient’s residence during three collection periods from November 2021 to May 2023

Genus (subgenus/species group)	Species name	November 2021 (winter)	March 2022 (summer)	May 2023 (rainy)	Total	
Unfed	Engorged	Unfed	Unfed	
C. (Remmia)	oxystoma	12	–	9	139	160 (29.3%)	
C. (Trithecoides)	spp.	66	5	8	6	85 (15.5%)	
C. (Shortti group)	shortti	11	18	16	20	65 (11.9%)	
C. (Avaritia)	orientalis	46	5	–	4	55 (10.1%)	
C. (Hoffmania)	innoxius	5	–	31	5	41 (7.5%)	
C. (Meijerehelea)	arakawae	1	–	1	32	34 (6.2%)	
C. (Hoffmania)	peregrinus	22	–	3	5	30 (5.5%)	
C. (Hoffmania)	sumatrae	19	–	1	–	20 (3.7%)	
C. (Meijerehelea)	guttifer	–	2	1	14	17 (3.1%)	
C. (Meijerehelea)	mahasarakhamense	–	2	1	11	14 (2.6%)	
C. (Hoffmania)	insignipennis	5	1	2	1	9 (1.6%)	
C. (Avaritia)	fulvus	–	7	–	–	7 (1.3%)	
C. (Avaritia)	actoni	–	5	–	–	5 (0.9%)	
C. (Avaritia)	jacobsoni	1	1	–	–	2 (0.4%)	
C. (Avaritia)	imicola	–	–	1	–	1 (0.2%)	
C. (Clavipalpis group)	huffi	–	–	–	1	1 (0.2%)	
C. (Hoffmania)	liui	–	–	1	–	1 (0.2%)	
Total	188	46	75	238	547	

Fig. 2 Wing morphology and pigmentation patterns of Culicoides species from the trap collections, shown in descending order of total relative abundance. The scale bar represents 200 µM

Infection prevalence in Culicoides and parasite identification

Of 501 non-engorged and 46 blood-engorged Culicoides samples tested, Leishmania SSU rRNA-qPCR was positive in 27 non-engorged and 4 engorged individuals, consisting of C. oxystoma (n = 6), C. innoxius (n = 4), C. shortti (n = 4), C. guttifer (n = 3), C. orientalis (n = 3), C. mahasarakhamense (n = 3), C. arakawae (n = 2), C. sumatrae (n = 2), C. (Trithecoides) spp. (n = 2), C. actoni (n = 1), and C. fulvus (n = 1), with an overall prevalence of 5.7%. Leishmania was temporally detected with the highest frequency in C. orientalis, C. innoxius, and C. oxystoma in November 2021, March 2022, and May 2023, respectively, as shown in Table 2. Conventional ITS1-PCR successfully amplified all qPCR-positive samples. Plasmid DNA sequencing and BLASTn analysis revealed that L. martiniquensis was commonly detected in 24 samples of these positive Culicoides species, whereas L. orientalis was only amplified in three samples of C. oxystoma. None of the samples showed co-infection with these two Leishmania (Mundinia) species. The Leishmania ITS1 sequences obtained from non-engorged and blood-engorged samples were deposited in GenBank under accession nos. OR917764-OR917790 and PQ014661-PQ014664, respectively.Table 2 Parasite identification and accession numbers of Leishmania sequences obtained from the patient and field-caught Culicoides biting midges by conventional ITS1-PCR and plasmid sequencing in this study

Collection date	Host species	Sample code	Parasite identification	Accession no.	BLASTn result and % identity	
August 2021	Homo sapiens (patient)	LP-VL	L. martiniquensis	OR917763	L. martiniquensis 770,605 (KY982650), 100%	
November

2021

	C. orientalis	LP002	L. martiniquensis	OR917764	L. martiniquensis SK4-1 (MK603826), 99.2%	
C. orientalis	LP118	L. martiniquensis	OR917765	L. martiniquensis 770,605 (KY982650), 100%	
C. orientalis	LP148	L. martiniquensis	OR917766	L. martiniquensis 770,605 (KY982650), 100%	
C. (Trithecoides) sp.	LP036	L. martiniquensis	OR917767	L. martiniquensis 770,605 (KY982650), 100%	
C. (Trithecoides) sp.	LP094	L. martiniquensis	OR917768	L. martiniquensis SK4-1 (MK603826), 98.5%	
C. sumatrae	LP048	L. martiniquensis	OR917769	L. martiniquensis SK4-1 (MK603826), 98.1%	
C. sumatrae	LP129	L. martiniquensis	OR917770	L. martiniquensis SK4-1 (MK603826), 98.5%	
C. innoxius	LP156	L. martiniquensis	OR917771	L. martiniquensis SK4-1 (MK603826), 99.2%	
C. fulvus	BF02	L. martiniquensis	PQ014661	L. martiniquensis 770,605 (KY982650), 100%	
C. actoni	BF03	L. martiniquensis	PQ014662	L. martiniquensis 770,605 (KY982650), 100%	
C. shortti	BF10	L. martiniquensis	PQ014663	L. martiniquensis 770,605 (KY982650), 100%	
C. shortti	BF21	L. martiniquensis	PQ014664	L. martiniquensis 770,605 (KY982650), 100%	
March 2022	C. innoxius	LP260	L. martiniquensis	OR917772	L. martiniquensis 770,605 (KY982650), 100%	
C. innoxius	LP272	L. martiniquensis	OR917773	L. martiniquensis TR17 (OR077858), 100%	
C. innoxius	LP286	L. martiniquensis	OR917774	L. martiniquensis TR17 (OR077858), 100%	
C. shortti	LP309	L. martiniquensis	OR917775	L. martiniquensis 770,605 (KY982650), 100%	
May 2023	C. oxystoma	LP364	L. martiniquensis	OR917776	L. martiniquensis 770,605 (KY982650), 100%	
C. oxystoma	LP478	L. martiniquensis	OR917777	L. martiniquensis 770,605 (KY982650), 100%	
C. oxystoma	LP496	L. martiniquensis	OR917778	L. martiniquensis 770,605 (KY982650), 98.9%	
C. oxystoma	LP408	L. orientalis	OR917779	L. orientalis 609,106 (KY982677), 99.2%	
C. oxystoma	LP419	L. orientalis	OR917780	L. orientalis PCM2 (JX195640), 100%	
C. oxystoma	LP443	L. orientalis	OR917781	L. orientalis PCM2 (JX195640), 100%	
C. mahasarakhamense	LP367	L. martiniquensis	OR917782	L. martiniquensis 770,605 (KY982650), 100%	
C. mahasarakhamense	LP374	L. martiniquensis	OR917783	L. martiniquensis 770,605 (KY982650), 99.6%	
C. mahasarakhamense	LP495	L. martiniquensis	OR917784	L. martiniquensis 770,605 (KY982650), 99.2%	
C. arakawae	LP429	L. martiniquensis	OR917785	L. martiniquensis 770,605 (KY982650), 100%	
C. arakawae	LP459	L. martiniquensis	OR917786	L. martiniquensis 770,605 (KY982650), 100%	
C. guttifer	LP446	L. martiniquensis	OR917787	L. martiniquensis 770,605 (KY982650), 100%	
C. guttifer	LP508	L. martiniquensis	OR917788	L. martiniquensis 770,605 (KY982650), 99.6%	
C. guttifer	LP532	L. martiniquensis	OR917789	L. martiniquensis 770,605 (KY982650), 100%	
C. shortti	LP474	L. martiniquensis	OR917790	L. martiniquensis 770,605 (KY982650), 100%	

Haplotype diversity and neutrality statistics of detected Leishmania

A total of 167 Leishmania ITS1 sequences, including 29 L. martiniquensis and 3 L. orientalis sequences amplified from the index case and Culicoides samples in our collection and 135 pre-exisitng sequences from the GenBank database, consisting of L. martiniquensis (n = 88) and L. orientalis (n = 47), were included in the haplotype diversity analysis. As shown in Fig. 3, the star-like network consisted of 32 and 27 unique haplotypes of L. martiniquensis and L. orientalis, respectively, circulating in Culicoides biting midges, animal hosts (cow, horse, and black rat), and leishmaniasis patients from Thailand and other geographical regions, as detailed in Supplementary File 1.Fig. 3 Haplotype diversity of Leishmania (Mundinia) species from different geographical origins based on ITS1 sequences. The size of the circle indicates the number of each unique haplotype. Hatch marks between the circles represent the number of nucleotide polymorphisms between haplotypes. Blue-green circles represent unique haplotypes of L. martiniquensis and L. orientalis identified in the present study

For L. martiniquensis, H01 was the most common haplotype shared by 72 sequences worldwide from several leishmaniasis-affected areas of Thailand, including Chanthaburi, Chiang Mai, Chiang Rai, Lampang (in this study), Lamphun, Nakhon Si Thammarat, Satun, Trang, and Songkhla Provinces, and other countries, namely Martinique Island, Myanmar, and the USA. The remaining descendant haplotypes were less common with varying degrees of regional area specificity. As shown in Tables 3 and 4, eight unique haplotypes with 14 polymorphic sites were identified from all L. martiniquensis sequences in our collection, with most sequences grouped into the dominant haplotype H01. The haplotype diversity for our collection and populations of L. martiniquensis in the northern provinces and the whole country was relatively high with values of 0.5556–0.6133. In contrast, nucleotide diversity for these three populations was exceptionally low with values of 0.0050–0.0063. In addition, neutrality tests showed significantly negative Tajima’s D values of −2.3353 and −2.5642 for the latter two populations and a significantly negative Fu and Li’s D value of −4.8110 for the whole country population.Table 3 Haplotypes of 32 Leishmania ITS1 sequences amplified from the index case and Culicoides biting midges collected in this study

Leishmania species	Haplotypes	No.	Accession numbers of associated Leishmania ITS1 sequences with human or Culicoides hosts	
L. martiniquensis	H01	18	OR917763 (patient),

OR917765 (C. orientalis), OR917766 (C. orientalis), OR917767 (C. (Trithecoides) sp.),

OR917772 (C. innoxius), OR917775 (C. shortti),

OR917776 (C. oxystoma), OR917777 (C. oxystoma), OR917782 (C. mahasarakhamense),

OR917785 (C. arakawae), OR917786 (C. arakawae),

OR917787 (C. guttifer), OR917789 (C. guttifer),

OR917790 (C. shortti)

PQ014661 (C. fulvus), PQ014662 (C. actoni)

PQ014663 (C. shortti), PQ014664 (C. shortti)

	
	H02	1	OR917778 (C. oxystoma)	
	H03	2	OR917773 (C. innoxius), OR917774 (C. innoxius)	
	H04	2	OR917783 (C. mahasarakhamense),

OR917788 (C. guttifer)

	
	H05	2	OR917764 (C. orientalis), OR917771 (C. innoxius)	
	H14	1	OR917784 (C. mahasarakhamense)	
	H15	1	OR917769 (C. sumatrae)	
	H17	2	OR917768 (C. (Trithecoides) sp.),

OR917770 (C. sumatrae)

	
L. orientalis	H41	1	OR917779 (C. oxystoma)	
	H44	2	OR917780 (C. oxystoma), OR917781 (C. oxystoma)	

Table 4 Genetic diversity and neutrality statistics of Leishmania martiniquensis and L. orientalis populations identified in Culicoides vectors and humans in different geographic localities of Thailand based on ITS1 region

	Population	Total sample size (n)	No. haplotypes (H)	No. polymorphic sites (S)	Average no. of nucleotide difference (k)	Haplotype diversity (Hd) ± SD	Nucleotide diversity (π) ± SD	Tajima's D	Fu and Li's D	
L. martiniquensis	Lampang Province - Culicoides (28) and the index case (1)	29	8	14	1.6256	0.6133 ± 0.1025	0.0063 ± 0.0042	−1.8282NS	−1.3159NS	
Northern Region - Culicoides (76) and humans (5)	81	20	27	1.3222	0.5556 ± 0.0680	0.0051 ± 0.0035	−2.3353*	−3.5626**	
Thailand - Culicoides (82), black rat (1), and humans (24)	107	29	39	1.2756	0.5842 ± 0.0585	0.0050 ± 0.0034	−2.5642*	−4.8110**	
L. orientalis	Lampang Province - Culicoides (3)	3	2	5	2.0000	0.6667 ± 0.2722	0.0080 ± 0.0073	NA	NA	
Northern Thailand - Culicoides (25) and humans (1)	26	19	25	2.0800	0.9569 ± 0.0259	0.0082 ± 0.0051	−2.4975*	−1.8429NS	
Thailand - Culicoides (26) and humans (24)	50	27	31	2.1396	0.9094 ± 0.0318	0.0086 ± 0.0052	−2.3052*	−2.9282**	
*p-value < 0.05; **p-value < 0.01; NS not significant; NA not analyzed

For L. orientalis, the H33 haplotype was the most dominant with 14 sequences from Chiang Rai and Nan in Northern Thailand and Trang Province in Southern Thailand. Two haplotypes (H41 and H44) with five polymorphic sites were identified in our collection. Two haplotypes, H44 and H49, could be found in different geographical locations. H44 was shared by four individuals from Lampang, Nakhon Si Thammarat, and Trang Provinces, while H49 was shared by five individuals from Chiang Rai, Trang, and Songkhla Provinces. This suggests possible gene flow across these sites. The haplotype diversity value for our collection was 0.6667, while those of L. orientalis populations in the northern region and the whole country were remarkably higher with values of 0.9569 and 0.9094, respectively. The lower degree of genetic differentiation of L. orientalis in our collection was probably due to the considerably small effective population size. As with L. martiniquensis, nucleotide diversity values for these groups ranged from 0.0080 to 0.0086. Neutrality tests were also significantly negative, with Tajima’s D values of − 2.4975 and − 2.3052 for the northern region and the whole country populations, respectively, and Fu and Li’s D value of − 2.9282 for the whole country population.

Blood meal source identification by host-specific Cytb-PCRs

A total of 46 blood-engorged females of nine Culicoides species were collected in November 2021 as detailed in Table 1. The Cytb-PCR reactions targeting mammalian and avian DNA were successfully performed on all engorged samples. Of these, four kinds of vertebrate hosts were identified with the following detection frequencies: cow (n = 41), human (n = 7), bird (n = 4), and pig (n = 1). Forty-one samples with cows as hosts consisted of Culicoides shortti (n = 18), C. fulvus (n = 7), C. orientalis (n = 4), C. actoni (n = 5), C. (Trithecoides) spp. (n = 5), C. jacobsoni (n = 1), and C. insignipennis (n = 1). Of these 41 samples, six samples of four species, namely C. shortti (n = 3), C. fulvus (n = 1), C. orientalis (n = 1), and C. actoni (n = 1), were found to feed on both cows and humans. Pig blood was detected in only one sample of C. orientalis, while none of the samples tested positive for dogs. Four samples of two Culicoides species, C. guttifer (n = 2) and C. mahasarakhamense (n = 2), were recorded as feeding on birds, and one of these two C. guttifer samples also fed on humans.

MinION® amplicon sequencing

A metabarcoding approach using nanopore-based COI amplicon sequencing was also used to generate high-throughput data, greatly facilitating the identification of multiple host species. COI-PCR was also successfully performed on all the engorged samples in which host-specific Cytb-PCRs were obtained. After filtering out low-quality reads, 739,245 high-quality reads with Q scores ≥ 20 were obtained from all individuals in the engorged collection. The resulting high-quality reads from each sample were clustered and error-polished to generate highly accurate consensus sequences. In total, 71 consensus sequences were generated and aligned against GenBank references using BLASTn for the host taxonomic assignment of each sample as detailed in Table 5. As shown in Fig. 4, six vertebrate host species were identified with the following detection frequencies: Bos indicus (cow, n = 41), Homo sapiens (human, n = 16), Gallus gallus (chicken, n = 5), Cervus sp. (deer, n = 4), and Sus scrofa (pig, n = 1), with high similarity percentages ranging from 94.7–99.3%, and unknown species related to Rupicapra sp. (goat-like species, n = 4) with similarity percentages of 89.9–90.5%.Table 5 The information on sources of blood meals in 46 blood-engorged Culicoides samples identified by host-specific Cytb-PCR and MinION® amplicon sequencing

Species	Sample ID	Leishmania
status	Host-specific Cytb-PCR	Vertebrate COI-PCR and MinION® sequencing	
Cow	Human	Pig	Dog	Bird	BLASTn result	No. assigned
reads	Relative abundance (%)	E value	Percent
Identity	
C. shortti	BF07	Negative	 + 	–	–	–	–	Bos indicus	24,347	100	0	98.9	
C. shortti	BF12	Negative	 + 	–	–	–	–	Bos indicus	14,260	100	0	98.9	
C. shortti	BF18	Negative	 + 	–	–	–	–	Bos indicus	36,482	100	0	98.6	
C. shortti	BF19	Negative	 + 	–	–	–	–	Bos indicus	24,036	100	0	98.6	
C. shortti	BF23	Negative	 + 	–	–	–	–	Bos indicus	17,571	100	0	98.6	
C. shortti	BF29	Negative	 + 	–	–	–	–	Bos indicus	24,789	100	0	98.6	
C. shortti	BF35	Negative	 + 	–	–	–	–	Bos indicus	35,262	100	0	98.4	
C. shortti	BF42	Negative	 + 	–	–	–	–	Bos indicus	25,506	100	0	98.9	
C. fulvus	BF14	Negative	 + 	–	–	–	-	Bos indicus	17,190	100	0	98.6	
C. fulvus	BF26	Negative	 + 	–	–	–	–	Bos indicus	41,004	100	0	98.9	
C. fulvus	BF34	Negative	 + 	–	–	–	–	Bos indicus	128	100	0	99.1	
C. fulvus	BF39	Negative	 + 	–	–	–	–	Bos indicus	390	100	0	98.6	
C. fulvus	BF43	Negative	 + 	–	–	–	–	Bos indicus	774	100	0	98.8	
C. (Trithecoides) sp.	BF16	Negative	 + 	–	–	–	–	Bos indicus	18,375	100	0	98.4	
C. (Trithecoides) sp.	BF25	Negative	 + 	–	–	–	–	Bos indicus	17,822	100	0	98.4	
C. (Trithecoides) sp.	BF27	Negative	 + 	–	–	–	–	Bos indicus	33,602	100	0	98.8	
C. (Trithecoides) sp.	BF38	Negative	 + 	–	–	–	–	Bos indicus	7,642	100	0	99.1	
C. orientalis	BF06	Negative	 + 	–	–	–	–	Bos indicus	20,214	100	0	98.6	
C. orientalis	BF11	Negative	 + 	–	–	–	–	Bos indicus	7,268	100	0	98.6	
C. orientalis	BF13	Negative	 + 	–	–	–	–	Bos indicus	21,969	100	0	98.9	
C. actoni	BF33	Negative	 + 	–	–	–	–	Bos indicus	1,247	100	0	98.6	
C. actoni	BF41	Negative	 + 	–	–	–	–	Bos indicus	10,885	100	0	99.3	
C. actoni	BF44	Negative	 + 	–	–	–	–	Bos indicus	2,111	100	0	99.1	
C. mahasarakhamense	BF45	Negative	–	–	–	–	 + 	Gallus gallus	8,509	100	0	98.9	
C. mahasarakhamense	BF46	Negative	–	–	–	–	 + 	Gallus gallus	2,623	100	0	98.9	
C. guttifer	BF37	Negative	–	–	–	–	 + 	Gallus gallus	8,693	100	0	99.1	
C. insignipennis	BF32	Negative	 + 	–	–	–	–	Bos indicus	18,772	100	0	98.6	
C. shortti	BF01	Negative	 + 	 + 	–	–	–	Bos indicus	814	68.7	0	99.1	
Homo sapiens	371	31.3	0	98.6	
C. shortti	BF05	Negative	 + 	 + 	–	–	–	Bos indicus	10,354	83.4	0	98.6	
Homo sapiens	2,055	16.6	0	98.6	
C. shortti	BF09	Negative	 + 	-	–	–	–	Bos indicus	14,663	97.9	0	98.6	
Homo sapiens	317	2.1	0	98.4	
C. shortti	BF10	L. martiniquensis	 + 	 + 	–	–	–	Bos indicus	17,182	95.4	0	98.6	
Homo sapiens	823	4.6	0	98.6	
C. shortti	BF21	L. martiniquensis	 + 	–	–	–	–	Bos indicus	7,293	96	0	98.4	
Homo sapiens	303	4	0	98.6	
C. fulvus	BF02	L. martiniquensis	 + 	 + 	–	–	–	Bos indicus	23,590	83.8	0	98.4	
Homo sapiens	4,565	16.2	0	98.6	
C. fulvus	BF30	Negative	 + 	–	–	–	–	Bos indicus	21,232	98.6	0	98.6	
Homo sapiens	298	1.4	0	98.6	
C. orientalis	BF20	Negative	 + 	 + 	–	–	–	Bos indicus	16,632	82.9	0	98.6	
Homo sapiens	3,422	17.1	0	98.6	
C. orientalis	BF22	Negative	–	–	 + 	–	–	Sus scrofa	9,465	96.6	0	98.8	
Homo sapiens	336	3.4	0	98.4	
C. actoni	BF03	L. martiniquensis	 + 	 + 	–	–	–	Bos indicus	3796	95.2	0	99.3	
Homo sapiens	193	4.8	0	98.6	
C. actoni	BF40	Negative	 + 	–	–	–	–	Bos indicus	21,723	97.1	0	98.4	
Cervus sp.	648	2.9	7e-177	94.7	
C. (Trithecoides) sp.	BF31	Negative	 + 	–	–	–	–	Bos indicus	3927	87.8	0	98.9	
Homo sapiens	547	12.2	0	98.4	
C. guttifer	BF36	Negative	–	 + 	–	–	 + 	Homo sapiens	11,942	51.5	0	98.6	
Gallus gallus	11,263	48.5	0	98.9	
C. jacobsoni	BF04	Negative	 + 	–	–	–	–	Bos indicus	19,366	95.8	0	98.6	
Homo sapiens	856	4.2	0	98.6	
C. shortti	BF08	Negative	 + 	–	–	–	-	Bos indicus	11,453	95.7	0	98.4	
Rupicapra sp.	328	2.7	5e-93	89.9	
Cervus sp.	192	1.6	2e-177	94.7	
C. shortti	BF15	Negative	 + 	–	–	–	–	Bos indicus	9800	94.4	0	98.6	
Homo sapiens	479	4.6	0	98.6	
Gallus gallus	106	1	0	99.1	
C. shortti	BF24	Negative	 + 	–	–	–	–	Bos indicus	17,122	92.3	0	98.6	
Homo sapiens	993	5.4	0	98.6	
Rupicapra sp.	442	2.4	4e-108	89.9	
C. shortti	BF28	Negative	 + 	–	–	–	–	Bos indicus	26,170	90.8	0	98.4	
Cervus sp.	1362	4.7	2e-177	94.7	
Rupicapra sp.	1299	4.5	1e-124	90.5	
C. shortti	BF17	Negative	 + 	–	–	–	–	Bos indicus	15,967	79.6	0	98.6	
Homo sapiens	2250	11.2	0	98.6	
Cervus sp.	1327	6.6	7e-177	94.7	
Rupicapra sp.	508	2.5	1e-124	90.5	

Fig. 4 (A) Circos plot illustrating the association between blood meal origins and each Culicoides species. (B) Bar chart showing the relative abundance of sequenced reads in 46 blood-engorged Culicoides samples analyzed by MinION® amplicon sequencing in this study

Performance comparison between host-specific Cytb-PCR and MinION® amplicon sequencing

Although there were differences in the frequency of hosts detected between host-specific Cytb-PCR and nanopore amplicon sequencing, the most common host was cow, identified in 41 of 46 samples (89.1%) by both techniques. The second most common host was human, 15.2% by host-specific Cytb-PCR and 34.8% by amplicon deep sequencing. The superiority of amplicon deep sequencing in generating the high-throughput sequenced read dataset allowed the identification of additional hosts not detected by host-specific Cytb-PCR and showed evidence of multiple blood meal origins within certain individuals of the engorged collection as shown in Table 5. Cytb-PCR showed evidence of two host species in only seven samples (15.2%). In contrast, amplicon deep sequencing identified 19 samples (41.3%) with multiple hosts, including 14 with two hosts, four with three hosts, and one with four hosts. Peridomestic species (cow, chicken, and pig), humans, and wildlife (deer and unknown species related to Rupicapra sp.) were identified as host species in Culicoides samples with evidence of multiple blood meals. Of the 11 samples with cow and human as a blood source, four samples, consisting of two C. shortti, one C. actoni, and one C. fulvus, were found positive for L. martiniquensis.

Discussion

The increasing prevalence of autochthonous human leishmaniasis, particularly in transmission areas of Thailand, has highlighted the importance of identifying vector and animal reservoir species and developing vector management strategies for effective transmission interruption. Recent experimental evidence has demonstrated successful promastigote development of several Leishmania (Mundinia) strains in laboratory-colonized C. sonorensis, suggesting that Culicoides biting midges are the imperative vectors of Leishmania (Mundinia) species rather than phlebotomine sand flies, which are traditionally known to be vectors of leishmaniasis [12–14]. However, biological information on the species diversity, population dynamics, infection prevalence, and host feeding patterns of Culicoides vectors remains poorly understood.

Temporal variability in species composition and abundance was observed in this study, suggesting that seasonality may influence the availability of favorable habitats for larvae of each Culicoides species. All Culicoides species were caught with the lowest abundance in summer with C. innoxius as the dominant species, whereas C. oxystoma and C. (Trithecoides) spp. were found to be the dominant species during the rainy season and winter, respectively. This could be because the summer could have contributed to the dryness of the soil habitats, strongly affecting immature Culicoides which require a moist environment and optimal environmental factors for their oviposition, larval development, and adult emergence [45–47], thus reducing the number of Culicoides populations during this period. In addition to seasonality, host availability and wind speed may also have influenced the species richness and abundance of Culicoides during these three collection periods [48–51].

This study demonstrated the concordance between SSU rRNA-qPCR and conventional ITS1-PCR results, showing Leishmania prevalence in at least 11 midge species, namely C. oxystoma, C. guttifer, C. orientalis, C. mahasarakhamense, C (Trithecoides) spp., C. innoxius, C. shortti, C. arakawae, C. sumatrae, C. actoni, and C. fulvus, with an overall prevalence of infection of 5.7%. Culicoides mahasarakhamense was first recorded as a vector of L. martiniquensis in Lamphun Province, Northern Thailand, in 2021 [26]. More recently, C. mahasarakhamense, C. guttifer, C. (Trithecoides) spp., C. jacobsoni, C. oxystoma, and C. orientalis have been implicated in the transmission of leishmaniasis in Chiang Rai Province, the most endemic area of leishmaniasis in Northern Thailand [27]. Therefore, the detection of Leishmania in the latter six species in our study represents the new records in Northern Thailand. Of note, L. martiniquensis was ubiquitously detected in all positive species over three seasonal periods with a high prevalence in the rainy season, followed by winter, whereas L. orientalis was only positive in three samples of C. oxystoma in the rainy season. Importantly, our finding indicates the sympatric occurrence of these two Leishmania (Mundinia) species circulating in the major livestock-associated biting midges in the emerging focus of leishmaniasis in Northern Thailand, suggesting that they may share Culicoides vector species for transmission to humans and animal reservoirs.

It could also be observed that Leishmania was detected with the highest frequency in the predominant species at a given collection time. It can be assumed that Leishmania parasites are transmitted to mammalian hosts through the bites of infected vectors; therefore, the maintenance of parasite transmission in nature appears to be closely associated with vector abundance [28, 47, 52, 53]. On the other hand, dominant vector species are more likely to increase the opportunities for pathogen transmission than less abundant species. Our findings are also consistent with the previous literature, which reported that the most abundant Culicoides species identified on cattle farms in Southern Ireland were associated with the transmission of arboviral diseases, including bluetongue and Schmallenberg viruses [54].

The intraspecific genetic diversity of L. martiniquensis and L. orientalis circulating in Culicoides biting midges, humans, and animal reservoirs in Thailand and other countries was revealed by the haplotype network of the ITS1 region in this study. Intriguingly, the haplotype network of each parasite species exhibited a star-like distribution with a dominant central haplotype and its multiple descendant lineages with few base differences, suggesting demographic expansion following the genetic bottleneck effect [55]. Negative neutrality statistics also signified that populations of these two parasites, particularly at regional and country levels, were likely to have undergone recent selective sweeps or population expansions [56, 57]. Essentially, the genetic divergence of the dominant core haplotype into novel descendent haplotypes appears to represent an evolutionary process by which the parasites have successfully adapted to a wide range of insect and mammalian host species in different geographical locations [58, 59].

For L. martiniquensis, the dominant haplotype H01 was the most ancestral and widespread globally, being shared by several individuals from different geographical origins. The existence of allopatric L. martiniquensis isolates with the H01 haplotype in different geographical locations strongly supports the hypothesis of a supercontinent origin, suggesting that the species divergence of the subgenus Mundinia occurred before the break-up of Gondwana [60, 61]. More importantly, haplotype H01 contained the sequences amplified from Culicoides samples in our collection and all leishmaniasis patients in Northern Thailand recorded to date, including the index case in this study. This genetic similarity confirms that natural Culicoides individuals are infected with the same Leishmania parasites circulating in humans in the same transmission area. Contrarily, L. orientalis was identified exclusively in Thailand [5], with populations of this species in the Northern region and throughout the country exhibiting significantly higher haplotype diversity than those of L. martiniquensis. Our results are consistent with previous literature showing considerable genetic differentiation of L. orientalis populations circulating in Culicoides and asymptomatic seropositive individuals in Chiang Rai Province [27, 62].

As previously published, the characterization of blood meal sources of Culicoides collected from different geographical locations in Thailand was based on conventional PCR targeting the mitochondrial Cytb and COI genes [28, 31, 48, 63] or the nuclear prepronociceptin (PNOC) gene [64], followed by Sanger sequencing. To our knowledge, domestic and peridomestic animals, including cattle, dogs, goats, and chickens, were identified as hosts of Culicoides species collected from areas of leishmaniasis endemicity [28, 31]. Notably, only three species, Culicoides brevitarsis, C. imicola, and C. oxystoma, collected from non-endemic areas, were recorded as opportunistic human blood-feeders [48, 64]. Thus, evidence of human blood feeding in leishmaniasis-endemic areas has never been demonstrated. As mentioned in the Background section, Sanger-based blood meal identification appears to be more prone to bias because this traditional sequencing method could only generate a single sequence for an individual sample. Multiplex Cytb-PCR is another good option for the identification of common vertebrate host species due to its high sensitivity and accuracy. However, this assay cannot identify wildlife species because of the limited number of primer pairs used. We therefore circumvent this by using a nanopore-based metabarcoding approach, which can differentiate long-read amplicon populations in samples of multiple origins to identify host species with higher taxonomic resolution without bias.

The vertebrate COI metabarcoding dataset revealed that all nine Culicoides species in our study are predominantly maintained by cows, followed by humans, chickens, deer, goat-like species, and pigs. Seven Culicoides species, namely C. shortti, C. fulvus, C. orientalis, C. actoni, C. (Trithecoides) sp., C. guttifer, and C. jacobsoni, represent the new records of feeding on human blood with percentages of human sequence reads ranging from 1.4 to 31.3% and exceptionally 51.5% in one sample of C. guttifer. However, most human feeding samples have a low percentage of human sequence reads, reflecting that human feeding is less frequent or opportunistic. This may be because most Culicoides species tend to be crepuscular in activity [29, 65]. They forage most actively at dawn and dusk and also bite at night when humans are in settlements and unlikely to be exposed to their bites. Our metabarcoding data are consistent with previous studies [31, 48, 63], showing that two C. guttifer and two C. mahasarakhamense samples in our engorged collection are ornithophilic and feed mainly on chickens.

Previously, L. martiniquensis has been reported to cause cutaneous leishmaniasis in cattle in Switzerland [66] and horses in Germany [67] and the USA [68]. In addition to cattle and horses, L. martiniquensis was molecularly detected in the visceral tissues of black rats captured in the endemic area of Songkhla Province, Southern Thailand [69], and in the buffy coat of a black rat from the patient’s residence in Chiang Rai Province [11]. Furthermore, Leishmania antibodies were also found in water buffaloes, dogs, and black rats from Chiang Rai Province [11]. These molecular and serological findings implicated domestic and peridomestic animals as reservoirs of autochthonous Leishmania parasites. In this study, blood-engorged Culicoides samples were also analyzed for Leishmania to shed light on the role of vertebrate hosts in the transmission cycle. Interestingly, four samples in our engorged collection, consisting of two C. shortti, one C. fulvus, one C. actoni, contained cow and human blood and all tested positive for L. martiniquensis. This strongly suggests that cattle may serve as an animal reservoir for L. martiniquensis, posing the risk of zoonotic transmission in this transmission area.

Our nanopore-based blood meal analysis provided promising evidence that Culicoides in leishmaniasis-endemic areas feed on humans and animal reservoirs, supporting the first criterion of Killick-Kendrick [30]. In addition, Leishmania parasites identified in Culicoides in this study were found to be genetically identical to those from patients in the same geographical area. This finding confirms Killick-Kendrick’s third criterion, which states that parasites from vectors and vertebrate hosts are indistinguishable [30]. Accordingly, our molecular evidence supports the role of positive Culicoides species as putative vectors of leishmaniasis. However, it remains insufficient to incriminate Culicoides as true natural vectors of Leishmania (Mundinia) parasites in Thailand, as parasite DNA may be detectable after host feeding, but parasite development and transmission may not occur [24]. To prove the major involvement of Thai Culicoides species in leishmaniasis transmission cycles, the full development of Leishmania (Mundinia) parasites in incriminated Culicoides species, at least by dissection of field-caught specimens and more ideally in controlled laboratory experiments, and the capability to transmit to the vertebrate hosts need to be demonstrated, which will also fulfill the second and fourth criteria of Killick-Kendrick [12–14, 24, 25, 30]. Therefore, colonization, experimental infection, and transmission of suspected vector species will be the next steps to confirm Culicoides as proven vectors responsible for the autochthonous transmission of Leishmania (Mundinia) parasites in Thailand.

Finally, our results revealed the seasonal population dynamics and host preference of Culicoides midge populations, as well as the infection prevalence, and haplotype diversity of two Leishmania (Mundinia) species sympatrically circulating in different Culicoides vector species in the newly emerging focus of leishmaniasis in Northern Thailand. The novel data from this study are an important step towards a more complete understanding of Culicoides midges and their medical importance in the transmission of leishmaniasis in areas of endemicity in Thailand as well as effective prevention and control of this neglected parasitic disease.

Conclusions

In this research, we investigated the species diversity, seasonal dynamics of Culicoides biting midges, and infection prevalence and genetic diversity of L. martiniquensis and L. orientalis circulating in different Culicoides species in the emerging focus of visceral leishmaniasis in Lampang Province, Northern Thailand. Our molecular findings revealed that L. martiniquensis was ubiquitously identified in at least 11 Culicoides species, whereas L. orientalis was only detected in C. oxystoma. Haplotype diversity analysis indicated recent population divergence of these two parasite species. The L. martiniquensis isolate identified in our index case was genetically identical to those circulating in Culicoides in the same area. Evidence of human blood feeding has been demonstrated, supporting the putative role of Culicoides in the autochthonous transmission of human leishmaniasis in Thailand. Blood meal analysis also suggested that peridomestic and wild animals, including cattle, deer, goat-like species, and pigs, may serve as parasite reservoirs with a risk of zoonotic transmission. Confirmation of metacyclic development by dissection of field-caught specimens, as well as colonization of Thai Culicoides species and experimental infection with Leishmania (Mundinia) parasites under laboratory conditions, will be performed in the next study. Essentially, the novel information from this study will facilitate a better understanding of the complexity of leishmaniasis transmission in endemic areas of Thailand and be a step towards developing vector control strategies to mitigate the endemicity of this neglected tropical disease effectively.

Supplementary Information

Additional file 1: Table S1. The information on 167 ITS1 sequences of Leishmania martiniquensis and L. orientalis isolates in humans, Culicoides biting midges, and animal hosts from Thailand and other geographical regions.

Abbreviations

UV Ultraviolet

RNA Ribonucleic acid

DNA Deoxyribonucleic acid

qPCR Quantitative polymerase chain reaction

PCR Polymerase chain reaction

Ct Cycle threshold

IPTG Isopropyl-beta-D-1-thiogalactopyranoside

X-gal 5-Bromo-4-chloro-3-indolyl-beta-D-galactoside

HIV Human immunodeficiency virus

Acknowledgements

We sincerely thank the staff members of the Center of Excellence in Vector Biology and Vector-Borne Disease and the Department of Parasitology, Faculty of Medicine, Chulalongkorn University, Bangkok, Thailand, for facilitating us with instrumentation and laboratory assistance.

Author contributions

KP conceptualized the idea of the study and designed the experiments. KP, CP, SSu, RB, PP and PS collected specimens. CP, SSu, CSa, TP, CSri, RA, MM, JS, PS and KP performed laboratory investigations. CP, SSri and KP analyzed the data. KP and CP wrote the paper. KP revised the manuscript. All authors have read and approved the final version of the manuscript.

Funding

This work has been supported by the Ratchadapiseksompote Fund, Graduate Affairs, Faculty of Medicine, Chulalongkorn University, Grant number GA68/001), the Faculty of Medicine, Chulalongkorn University, Bangkok, Thailand.

Availability of data and materials

All data supporting the findings of this study are available within the paper and its Supplementary Information. The Leishmania ITS1 sequences obtained in this study have been uploaded into GenBank with assigned accession numbers OR917763-OR917790 and PQ014661-PQ014664.

Declarations

Ethics approval and consent to participate

The procedures and research methodology for clinical specimen collection, entomological investigation and molecular analysis were approved by the Institutional Review Board of the Faculty of Medicine, Chulalongkorn University, Bangkok, Thailand (IRB No. 0613/67, COE No. 048/2024) and the animal research ethics committee of Chulalongkorn University Animal Care and Use Protocol (CU-ACUP), Faculty of Medicine, Chulalongkorn University, Bangkok, Thailand (COA no. 004/2564).

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.

Publisher's Note

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

1. Leelayoova S Siripattanapipong S Manomat J Piyaraj P Tan-Ariya P Bualert L Leishmaniasis in Thailand: a review of causative agents and situations Am J Trop Med Hyg 2017 96 534 542 10.4269/ajtmh.16-0604 28093539
Leelayoova S, Siripattanapipong S, Manomat J, Piyaraj P, Tan-Ariya P, Bualert L, et al. Leishmaniasis in Thailand: a review of causative agents and situations. Am J Trop Med Hyg. 2017;96:534–42.28093539
2. Pothirat T Tantiworawit A Chaiwarith R Jariyapan N Wannasan A Siriyasatien P First isolation of Leishmania from Northern Thailand: case report, identification as Leishmania martiniquensis and phylogenetic position within the Leishmania enriettii complex PLoS Negl Trop Dis 2014 8 e3339 10.1371/journal.pntd.0003339 25474647
Pothirat T, Tantiworawit A, Chaiwarith R, Jariyapan N, Wannasan A, Siriyasatien P, et al. First isolation of Leishmania from Northern Thailand: case report, identification as Leishmania martiniquensis and phylogenetic position within the Leishmania enriettii complex. PLoS Negl Trop Dis. 2014;8:e3339.25474647
3. Jariyapan N Daroontum T Jaiwong K Chanmol W Intakhan N Sor-Suwan S Leishmania (Mundinia) orientalis n sp (Trypanosomatidae), a parasite from Thailand responsible for localised cutaneous leishmaniasis Parasit Vectors 2018 11 351 10.1186/s13071-018-2908-3 29914526
Jariyapan N, Daroontum T, Jaiwong K, Chanmol W, Intakhan N, Sor-Suwan S, et al. Leishmania (Mundinia) orientalis n. sp. (Trypanosomatidae), a parasite from Thailand responsible for localised cutaneous leishmaniasis. Parasit Vectors. 2018;11:351.29914526
4. Srivarasat S Brownell N Siriyasatien P Noppakun N Asawanonda P Rattanakorn K Case report: autochthonous disseminated cutaneous, mucocutaneous, and visceral leishmaniasis caused by Leishmania martiniquensis in a patient with HIV/AIDS from northern Thailand and Literature Review Am J Trop Med Hyg 2022 107 1196 1202 10.4269/ajtmh.22-0108 36375453
Srivarasat S, Brownell N, Siriyasatien P, Noppakun N, Asawanonda P, Rattanakorn K, et al. Case report: autochthonous disseminated cutaneous, mucocutaneous, and visceral leishmaniasis caused by Leishmania martiniquensis in a patient with HIV/AIDS from Northern Thailand and literature review. Am J Trop Med Hyg. 2022;107:1196–202.36375453
5. Anugulruengkitt S Songtaweesin WN Thepnarong N Tangthanapalakul A Sitthisan M Chatproedprai S Case report: simple nodular cutaneous leishmaniasis caused by Autochthonous Leishmania (Mundinia) orientalis in an 18-Month-old girl: the first pediatric case in Thailand and Literature Review Am J Trop Med Hyg 2023 108 44 50 10.4269/ajtmh.22-0385 36410322
Anugulruengkitt S, Songtaweesin WN, Thepnarong N, Tangthanapalakul A, Sitthisan M, Chatproedprai S, et al. Case report: simple nodular cutaneous leishmaniasis caused by autochthonous Leishmania (Mundinia) orientalis in an 18-month-old girl: the first pediatric case in Thailand and literature review. Am J Trop Med Hyg. 2023;108:44–50.36410322
6. Burza S Croft SL Boelaert M Leishmaniasis Lancet 2018 392 951 970 10.1016/S0140-6736(18)31204-2 30126638
Burza S, Croft SL, Boelaert M. Leishmaniasis. Lancet. 2018;392:951–70.30126638
7. Srisuton P Phumee A Sunantaraporn S Boonserm R Sor-Suwan S Brownell N Detection of Leishmania and Trypanosoma dna in field-caught sand flies from endemic and non-endemic areas of leishmaniasis in Southern Thailand Insects 2019 10 238 10.3390/insects10080238 31382501
Srisuton P, Phumee A, Sunantaraporn S, Boonserm R, Sor-Suwan S, Brownell N, et al. Detection of Leishmania and Trypanosoma DNA in field-caught sand flies from endemic and non-endemic areas of leishmaniasis in Southern Thailand. Insects. 2019;10:238.31382501
8. Chusri S Thammapalo S Chusri S Thammapalo S Silpapojakul K Siriyasatien P Animal reservoirs and potential vectors of Leishmania siamensis in southern Thailand Southeast Asian J Trop Med Public Health 2014 45 13 19 24964648
Chusri S, Thammapalo S, Chusri S, Thammapalo S, Silpapojakul K, Siriyasatien P. Animal reservoirs and potential vectors of Leishmania siamensis in Southern Thailand. Southeast Asian J Trop Med Public Health. 2014;45:13–9.24964648
9. Siripattanapipong S Leelayoova S Ninsaeng U Mungthin M Detection of DNA of Leishmania siamensis in Sergentomyia (Neophlebotomus) iyengari (Diptera: Psychodidae) and molecular identification of blood meals of sand flies in an affected area Southern Thailand J Med Entomol 2018 55 1277 1283 29688539
Siripattanapipong S, Leelayoova S, Ninsaeng U, Mungthin M. Detection of DNA of Leishmania siamensis in Sergentomyia (Neophlebotomus) iyengari (Diptera: Psychodidae) and molecular identification of blood meals of sand flies in an affected area, Southern Thailand. J Med Entomol. 2018;55:1277–83.29688539
10. Preativatanyou K Chinwirunsirisup K Phumee A Khositharattanakool P Sunantaraporn S Depaquit J Species diversity of phlebotomine sand flies and sympatric occurrence of Leishmania (Mundinia) martiniquensis, Leishmania (Leishmania) donovani complex, and Trypanosoma spp in the visceral leishmaniasis focus of southern Thailand Acta Trop 2023 244 106949 10.1016/j.actatropica.2023.106949 37211153
Preativatanyou K, Chinwirunsirisup K, Phumee A, Khositharattanakool P, Sunantaraporn S, Depaquit J, et al. Species diversity of phlebotomine sand flies and sympatric occurrence of Leishmania (Mundinia) martiniquensis, Leishmania (Leishmania) donovani complex, and Trypanosoma spp. in the visceral leishmaniasis focus of Southern Thailand. Acta Trop. 2023;244:106949.37211153
11. Sriwongpan P Nedsuwan S Manomat J Charoensakulchai S Lacharojana K Sankwan J Prevalence and associated risk factors of Leishmania infection among immunocompetent hosts, a community-based study in Chiang Rai Thailand PLoS Negl Trop Dis 2021 15 e0009545 10.1371/journal.pntd.0009545 34252099
Sriwongpan P, Nedsuwan S, Manomat J, Charoensakulchai S, Lacharojana K, Sankwan J, et al. Prevalence and associated risk factors of Leishmania infection among immunocompetent hosts, a community-based study in Chiang Rai, Thailand. PLoS Negl Trop Dis. 2021;15:e0009545.34252099
12. Becvar T Vojtkova B Siriyasatien P Votypka J Modry D Jahn P Experimental transmission of Leishmania (Mundinia) parasites by biting midges (Diptera: Ceratopogonidae) PLoS Pathog 2021 17 e1009654 10.1371/journal.ppat.1009654 34115806
Becvar T, Vojtkova B, Siriyasatien P, Votypka J, Modry D, Jahn P, et al. Experimental transmission of Leishmania (Mundinia) parasites by biting midges (Diptera: Ceratopogonidae). PLoS Pathog. 2021;17:e1009654.34115806
13. Seblova V Sadlova J Vojtkova B Votypka J Carpenter S Bates PA The biting midge Culicoides sonorensis (Diptera: Ceratopogonidae) is capable of developing late stage infections of Leishmania enriettii PLoS Negl Trop Dis 2015 9 e0004060 10.1371/journal.pntd.0004060 26367424
Seblova V, Sadlova J, Vojtkova B, Votypka J, Carpenter S, Bates PA, et al. The biting midge Culicoides sonorensis (Diptera: Ceratopogonidae) is capable of developing late stage infections of Leishmania enriettii. PLoS Negl Trop Dis. 2015;9:e0004060.26367424
14. Chanmol W Jariyapan N Somboon P Bates MD Bates PA Development of Leishmania orientalis in the sand fly Lutzomyia longipalpis (Diptera: Psychodidae) and the biting midge Culicoides soronensis (Diptera: Ceratopogonidae) Acta Trop 2019 199 105157 10.1016/j.actatropica.2019.105157 31491400
Chanmol W, Jariyapan N, Somboon P, Bates MD, Bates PA. Development of Leishmania orientalis in the sand fly Lutzomyia longipalpis (Diptera: Psychodidae) and the biting midge Culicoides soronensis (Diptera: Ceratopogonidae). Acta Trop. 2019;199:105157.31491400
15. Phumee A Tawatsin A Thavara U Pengsakul T Thammapalo S Depaquit J Detection of an Unknown Trypanosoma DNA in a Phlebotomus stantoni (Diptera: Psychodidae) collected from southern thailand and records of new sand flies with reinstatement of Sergentomyia hivernus raynal & gaschen, 1935 (Diptera: Psychodidae) J Med Entomol 2017 54 429 434 27744363
Phumee A, Tawatsin A, Thavara U, Pengsakul T, Thammapalo S, Depaquit J, et al. Detection of an unknown Trypanosoma DNA in a Phlebotomus stantoni (Diptera: Psychodidae) collected from Southern thailand and records of new sand flies with reinstatement of Sergentomyia hivernus Raynal & Gaschen, 1935 (Diptera: Psychodidae). J Med Entomol. 2017;54:429–34.27744363
16. Sick F Beer M Kampen H Wernike K Culicoides biting midges-underestimated vectors for arboviruses of public health and veterinary importance Viruses 2019 11 376 10.3390/v11040376 31022868
Sick F, Beer M, Kampen H, Wernike K. Culicoides biting midges-underestimated vectors for arboviruses of public health and veterinary importance. Viruses. 2019;11:376.31022868
17. Mullens GR Murphree CS Mullen GR Durden LA Biting midges (Ceratopogonidae) Medical and veterinary entomology 2019 3 San Diego Academic Press 213 236
Mullens GR, Murphree CS. Biting midges (Ceratopogonidae). In: Mullen GR, Durden LA, editors. Medical and veterinary entomology. 3rd ed. San Diego: Academic Press; 2019. p. 213–36.
18. Prasad G, Bhatnagar P. 2000. Culicoides biology and transmission of viruses. V National Training programme Compendium of Acarines and Insects of Veterinary and Medical Importance On Centre of Advanced Studies Department of Parasitology Veterinary College Bangalore. p. 34–8
19. Svobodová M Dolnik OV Čepička I Rádrová J Biting midges (Ceratopogonidae) as vectors of avian trypanosomes Parasit Vectors 2017 10 224 10.1186/s13071-017-2158-9 28482865
Svobodová M, Dolnik OV, Čepička I, Rádrová J. Biting midges (Ceratopogonidae) as vectors of avian trypanosomes. Parasit Vectors. 2017;10:224.28482865
20. Bernotienė R Iezhova TA Bukauskaitė D Chagas CRF Kazak M Valkiūnas G Development of Trypanosoma everetti in Culicoides biting midges Acta Trop 2020 210 105555 10.1016/j.actatropica.2020.105555 32473117
Bernotienė R, Iezhova TA, Bukauskaitė D, Chagas CRF, Kazak M, Valkiūnas G. Development of Trypanosoma everetti in Culicoides biting midges. Acta Trop. 2020;210:105555.32473117
21. Rebêlo JM Rodrigues BL Bandeira MD Moraes JL Fonteles RS Pereira SR Detection of Leishmania amazonensis and Leishmania braziliensis in Culicoides (Diptera, Ceratopogonidae) in an endemic area of cutaneous leishmaniasis in the Brazilian Amazonia J Vector Ecol 2016 41 303 308 10.1111/jvec.12227 27860021
Rebêlo JM, Rodrigues BL, Bandeira MD, Moraes JL, Fonteles RS, Pereira SR. Detection of Leishmania amazonensis and Leishmania braziliensis in Culicoides (Diptera, Ceratopogonidae) in an endemic area of cutaneous leishmaniasis in the Brazilian Amazonia. J Vector Ecol. 2016;41:303–8.27860021
22. Slama D Haouas N Remadi L Mezhoud H Babba H Chaker E First detection of Leishmania infantum (Kinetoplastida: Trypanosomatidae) in culicoides spp. (Diptera: Ceratopogonidae) Parasit Vectors. 2014 7 51 10.1186/1756-3305-7-51 24460752
Slama D, Haouas N, Remadi L, Mezhoud H, Babba H, Chaker E. First detection of Leishmania infantum (Kinetoplastida: Trypanosomatidae) in Culicoides spp. (Diptera: Ceratopogonidae). Parasit Vectors. 2014;7:51.24460752
23. Ríos-Tostado JJ Castillo-Ureta H Torres-Montoya EH Torres-Avendaño JI Olimón-Andalón V Romero-Higareda CE Molecular detection of leishmania (L) mexicana (Kinetoplastida: Trypanostomatidae) DNA in Culicoides furens (Diptera: Ceratopogonidae) from an area with autochthonous canine leishmaniasis in northwestern mexico Acta Parasitol 2021 66 1055 8 10.1007/s11686-021-00335-1 33554301
Ríos-Tostado JJ, Castillo-Ureta H, Torres-Montoya EH, Torres-Avendaño JI, Olimón-Andalón V, Romero-Higareda CE, et al. Molecular detection of Leishmania (L.) mexicana (Kinetoplastida: Trypanostomatidae) DNA in Culicoides furens (Diptera: Ceratopogonidae) from an area with autochthonous canine leishmaniasis in Northwestern Mexico. Acta Parasitol. 2021;66:1055–8.33554301
24. Dostálová A Volf P Leishmania development in sand flies: parasite-vector interactions overview Parasit Vectors 2012 5 276 10.1186/1756-3305-5-276 23206339
Dostálová A, Volf P. Leishmania development in sand flies: parasite-vector interactions overview. Parasit Vectors. 2012;5:276.23206339
25. Vaselek S Volf P Experimental infection of Phlebotomus perniciosus and Phlebotomus tobbi with different Leishmania tropica strains Int J Parasitol 2019 49 831 835 10.1016/j.ijpara.2019.05.009 31401063
Vaselek S, Volf P. Experimental infection of Phlebotomus perniciosus and Phlebotomus tobbi with different Leishmania tropica strains. Int J Parasitol. 2019;49:831–5.31401063
26. Sunantaraporn S Thepparat A Phumee A Sor-Suwan S Boonserm R Bellis G Culicoides Latreille (Diptera: Ceratopogonidae) as potential vectors for Leishmania martiniquensis and Trypanosoma sp in northern Thailand PLoS Negl Trop Dis. 2021 15 e0010014 10.1371/journal.pntd.0010014 34910720
Sunantaraporn S, Thepparat A, Phumee A, Sor-Suwan S, Boonserm R, Bellis G, et al. Culicoides Latreille (Diptera: Ceratopogonidae) as potential vectors for Leishmania martiniquensis and Trypanosoma sp. in Northern Thailand. PLoS Negl Trop Dis. 2021;15:e0010014.34910720
27. Ampol R Somwang P Khositharattanakool P Promrangsee C Pataradool T Tepboonreung P Nanopore-based surveillance of Leishmania Parasites in Culicoides Latrielle (Diptera: Ceratopogonidae) caught from the affected community and tham phra cave in chiang rai province, the endemic area of leishmaniasis in Northern Thailand Insects 2024 15 327 10.3390/insects15050327 38786883
Ampol R, Somwang P, Khositharattanakool P, Promrangsee C, Pataradool T, Tepboonreung P, et al. Nanopore-based surveillance of Leishmania parasites in Culicoides Latrielle (Diptera: Ceratopogonidae) caught from the affected community and Tham Phra cave in Chiang Rai Province, the endemic area of leishmaniasis in Northern Thailand. Insects. 2024;15:327.38786883
28. Songumpai N Promrangsee C Noopetch P Siriyasatien P Preativatanyou K First evidence of co-circulation of emerging leishmania martiniquensis, leishmania orientalis, and crithidia sp in culicoides biting midges (diptera: ceratopogonidae), the putative vectors for autochthonous transmission in Southern Thailand Trop Med Infect Dis 2022 7 379 10.3390/tropicalmed7110379 36422930
Songumpai N, Promrangsee C, Noopetch P, Siriyasatien P, Preativatanyou K. First evidence of co-circulation of emerging Leishmania martiniquensis, Leishmania orientalis, and Crithidia sp. in Culicoides biting midges (diptera: ceratopogonidae), the putative vectors for autochthonous transmission in Southern Thailand. Trop Med Infect Dis. 2022;7:379.36422930
29. Wirth WW Hubert AA The Culicoides of Southeast Asia (Diptera Ceratopogonidae) 44 1989 FL Memoirs of the American Entomological Institute
Wirth WW, Hubert AA. The Culicoides of Southeast Asia (Diptera Ceratopogonidae) 44. FL: Memoirs of the American Entomological Institute; 1989.
30. Killick-Kendrick R The biology and control of phlebotomine sand flies Clin Dermatol 1999 17 279 289 10.1016/S0738-081X(99)00046-2 10384867
Killick-Kendrick R. The biology and control of phlebotomine sand flies. Clin Dermatol. 1999;17:279–89.10384867
31. Sunantaraporn S Hortiwakul T Kraivichian K Siriyasatien P Brownell N Molecular identification of host blood meals and detection of blood parasites in Culicoides Latreille (Diptera: Ceratopogonidae) collected from phatthalung province Southern Thailand Insects 2022 13 912 10.3390/insects13100912 36292860
Sunantaraporn S, Hortiwakul T, Kraivichian K, Siriyasatien P, Brownell N. Molecular identification of host blood meals and detection of blood parasites in Culicoides Latreille (Diptera: Ceratopogonidae) collected from Phatthalung Province, Southern Thailand. Insects. 2022;13:912.36292860
32. Pramual P Jomkumsing P Piraonapicha K Jumpato W Integrative taxonomy uncovers a new Culicoides (Diptera: Ceratopogonidae) biting midge species from Thailand Acta Trop 2021 220 105941 10.1016/j.actatropica.2021.105941 33951420
Pramual P, Jomkumsing P, Piraonapicha K, Jumpato W. Integrative taxonomy uncovers a new Culicoides (Diptera: Ceratopogonidae) biting midge species from Thailand. Acta Trop. 2021;220:105941.33951420
33. Santos D Ribeiro GC Cabral AD Sperança MA A non-destructive enzymatic method to extract DNA from arthropod specimens: Implications for morphological and molecular studies PLoS ONE 2018 13 e0192200 10.1371/journal.pone.0192200 29390036
Santos D, Ribeiro GC, Cabral AD, Sperança MA. A non-destructive enzymatic method to extract DNA from arthropod specimens: Implications for morphological and molecular studies. PLoS ONE. 2018;13:e0192200.29390036
34. van der Meide W Guerra J Schoone G Farenhorst M Coelho L Faber W Comparison between quantitative nucleic acid sequence-based amplification, real-time reverse transcriptase PCR, and real-time PCR for quantification of Leishmania parasites J Clin Microbiol 2008 46 73 78 10.1128/JCM.01416-07 17959763
van der Meide W, Guerra J, Schoone G, Farenhorst M, Coelho L, Faber W, et al. Comparison between quantitative nucleic acid sequence-based amplification, real-time reverse transcriptase PCR, and real-time PCR for quantification of Leishmania parasites. J Clin Microbiol. 2008;46:73–8.17959763
35. Manomat J Leelayoova S Bualert L Tan-Ariya P Siripattanapipong S Mungthin M Prevalence and risk factors associated with Leishmania infection in Trang Province, southern Thailand PLoS Negl Trop Dis 2017 11 e0006095 10.1371/journal.pntd.0006095 29155831
Manomat J, Leelayoova S, Bualert L, Tan-Ariya P, Siripattanapipong S, Mungthin M, et al. Prevalence and risk factors associated with Leishmania infection in Trang Province, Southern Thailand. PLoS Negl Trop Dis. 2017;11:e0006095.29155831
36. RStudio Team. RStudio: integrated development environment for R. RStudio. 2021. Available online: http://www.rstudio.com/ Accessed 17 July 2024
37. Jombart T adegenet: a R package for the multivariate analysis of genetic markers Bioinformatics 2008 24 1403 1405 10.1093/bioinformatics/btn129 18397895
Jombart T. adegenet: a R package for the multivariate analysis of genetic markers. Bioinformatics. 2008;24:1403–5.18397895
38. Paradis E pegas: an R package for population genetics with an integrated-modular approach Bioinformatics 2010 26 419 420 10.1093/bioinformatics/btp696 20080509
Paradis E. pegas: an R package for population genetics with an integrated-modular approach. Bioinformatics. 2010;26:419–20.20080509
39. Pfeifer B Wittelsbürger U Ramos-Onsins SE Lercher MJ PopGenome: an efficient Swiss army knife for population genomic analyses in R Mol Biol Evol 2014 31 1929 1936 10.1093/molbev/msu136 24739305
Pfeifer B, Wittelsbürger U, Ramos-Onsins SE, Lercher MJ. PopGenome: an efficient Swiss army knife for population genomic analyses in R. Mol Biol Evol. 2014;31:1929–36.24739305
40. Kent RJ Norris DE Identification of mammalian blood meals in mosquitoes by a multiplexed polymerase chain reaction targeting cytochrome B Am J Trop Med Hyg 2005 73 336 342 10.4269/ajtmh.2005.73.336 16103600
Kent RJ, Norris DE. Identification of mammalian blood meals in mosquitoes by a multiplexed polymerase chain reaction targeting cytochrome B. Am J Trop Med Hyg. 2005;73:336–42.16103600
41. Cicero C Johnson NK Speciation in sapsuckers (Sphyrapicus): III mitochondrial-DNA sequence divergence at the cytochrome-b locus Auk 1995 112 547 63
Cicero C, Johnson NK. Speciation in sapsuckers (Sphyrapicus): III mitochondrial-DNA sequence divergence at the cytochrome-b locus. Auk. 1995;112:547–63.
42. Reeves LE Gillett-Kaufman JL Kawahara AY Kaufman PE Barcoding blood meals: New vertebrate-specific primer sets for assigning taxonomic identities to host DNA from mosquito blood meals PLoS Negl Trop Dis 2018 12 e0006767 10.1371/journal.pntd.0006767 30161128
Reeves LE, Gillett-Kaufman JL, Kawahara AY, Kaufman PE. Barcoding blood meals: New vertebrate-specific primer sets for assigning taxonomic identities to host DNA from mosquito blood meals. PLoS Negl Trop Dis. 2018;12:e0006767.30161128
43. De Coster W Rademakers R NanoPack 2: population-scale evaluation of long-read sequencing data Bioinformatics 2023 39 5 10.1093/bioinformatics/btad311
De Coster W, Rademakers R. NanoPack 2: population-scale evaluation of long-read sequencing data. Bioinformatics. 2023;39:5.
44. Vierstraete AR Braeckman BP Amplicon_sorter: A tool for reference-free amplicon sorting based on sequence similarity and for building consensus sequences Ecol Evol 2022 12 e8603 10.1002/ece3.8603 35261737
Vierstraete AR, Braeckman BP. Amplicon_sorter: A tool for reference-free amplicon sorting based on sequence similarity and for building consensus sequences. Ecol Evol. 2022;12:e8603.35261737
45. Zimmer JY Brostaux Y Haubruge E Francis F Larval development sites of the main Culicoides species (Diptera: Ceratopogonidae) in northern Europe and distribution of coprophilic species larvae in Belgian pastures Vet Parasitol 2014 205 676 686 10.1016/j.vetpar.2014.08.029 25241330
Zimmer JY, Brostaux Y, Haubruge E, Francis F. Larval development sites of the main Culicoides species (Diptera: Ceratopogonidae) in Northern Europe and distribution of coprophilic species larvae in Belgian pastures. Vet Parasitol. 2014;205:676–86.25241330
46. Zimmer JY Saegerman C Losson B Beckers Y Haubruge E Francis F Chemical composition of silage residues sustaining the larval development of the Culicoides obsoletus/Culicoides scoticus species (Diptera: Ceratopogonidae) Vet Parasitol 2013 191 197 201 10.1016/j.vetpar.2012.08.014 22963713
Zimmer JY, Saegerman C, Losson B, Beckers Y, Haubruge E, Francis F. Chemical composition of silage residues sustaining the larval development of the Culicoides obsoletus/Culicoides scoticus species (Diptera: Ceratopogonidae). Vet Parasitol. 2013;191:197–201.22963713
47. Duan YL Bellis G Liu BG Li L Diversity and seasonal abundance of Culicoides (Diptera, Ceratopogonidae) in Shizong County, Yunnan Province China Parasite 2022 29 26 10.1051/parasite/2022027 35543529
Duan YL, Bellis G, Liu BG, Li L. Diversity and seasonal abundance of Culicoides (Diptera, Ceratopogonidae) in Shizong County, Yunnan Province, China. Parasite. 2022;29:26.35543529
48. Gomontean B Vaisusuk K Chatan W Wongpakam K Sankul P Lachanthuek L Diversity, abundance and host blood meal analysis of Culicoides Latreille (Diptera: Ceratopogonidae) from cattle pens in different land use types from Thailand Insects 2023 14 574 10.3390/insects14070574 37504581
Gomontean B, Vaisusuk K, Chatan W, Wongpakam K, Sankul P, Lachanthuek L, et al. Diversity, abundance and host blood meal analysis of Culicoides Latreille (Diptera: Ceratopogonidae) from cattle pens in different land use types from Thailand. Insects. 2023;14:574.37504581
49. Murray M Local dispersal of the biting midge Culicoides brevitarsis kieffer (diptera, ceratopogonidae) in southeastern Australia Aust J Zool 1987 35 559 573 10.1071/ZO9870559
Murray M. Local dispersal of the biting midge Culicoides brevitarsis Kieffer (Diptera, Ceratopogonidae) in Southeastern Australia. Aust J Zool. 1987;35:559–73.
50. Ducheyne E De Deken R Bécu S Codina B Nomikou K Mangana-Vougiaki O Quantifying the wind dispersal of Culicoides species in Greece and Bulgaria Geospat Health 2007 1 177 189 10.4081/gh.2007.266 18686243
Ducheyne E, De Deken R, Bécu S, Codina B, Nomikou K, Mangana-Vougiaki O, et al. Quantifying the wind dispersal of Culicoides species in Greece and Bulgaria. Geospat Health. 2007;1:177–89.18686243
51. Mignotte A Garros C Dellicour S Jacquot M Gilbert M Gardès L High dispersal capacity of Culicoides obsoletus (Diptera: Ceratopogonidae), vector of bluetongue and Schmallenberg viruses, revealed by landscape genetic analyses Parasit Vectors 2021 14 93 10.1186/s13071-020-04522-3 33536057
Mignotte A, Garros C, Dellicour S, Jacquot M, Gilbert M, Gardès L, et al. High dispersal capacity of Culicoides obsoletus (Diptera: Ceratopogonidae), vector of bluetongue and Schmallenberg viruses, revealed by landscape genetic analyses. Parasit Vectors. 2021;14:93.33536057
52. Purse BV Carpenter S Venter GJ Bellis G Mullens BA Bionomics of temperate and tropical Culicoides midges: knowledge gaps and consequences for transmission of Culicoides-borne viruses Annu Rev Entomol 2015 60 373 392 10.1146/annurev-ento-010814-020614 25386725
Purse BV, Carpenter S, Venter GJ, Bellis G, Mullens BA. Bionomics of temperate and tropical Culicoides midges: knowledge gaps and consequences for transmission of Culicoides-borne viruses. Annu Rev Entomol. 2015;60:373–92.25386725
53. Wittmann EJ Baylis M Climate change: effects on culicoides–transmitted viruses and implications for the UK Vet J 2000 160 107 117 10.1016/S1090-0233(00)90470-2 10985802
Wittmann EJ, Baylis M. Climate change: effects on Culicoides–transmitted viruses and implications for the UK. Vet J. 2000;160:107–17.10985802
54. Collins ÁB Mee JF Doherty ML Barrett DJ England ME Culicoides species composition and abundance on Irish cattle farms: implications for arboviral disease transmission Parasit Vectors 2018 11 472 10.1186/s13071-018-3010-6 30119685
Collins ÁB, Mee JF, Doherty ML, Barrett DJ, England ME. Culicoides species composition and abundance on Irish cattle farms: implications for arboviral disease transmission. Parasit Vectors. 2018;11:472.30119685
55. Ferreri M Qu W Han B Phylogenetic networks: a tool to display character conflict and demographic history Afr J Biotech 2011 10 12799 12803 10.5897/AJB11.010
Ferreri M, Qu W, Han B. Phylogenetic networks: a tool to display character conflict and demographic history. Afr J Biotech. 2011;10:12799–803.
56. Ramos-Onsins SE Rozas J Statistical properties of new neutrality tests against population growth Mol Biol Evol 2002 19 2092 2100 10.1093/oxfordjournals.molbev.a004034 12446801
Ramos-Onsins SE, Rozas J. Statistical properties of new neutrality tests against population growth. Mol Biol Evol. 2002;19:2092–100.12446801
57. Nielsen R Williamson S Kim Y Hubisz MJ Clark AG Bustamante C Genomic scans for selective sweeps using SNP data Genome Res 2005 15 1566 1575 10.1101/gr.4252305 16251466
Nielsen R, Williamson S, Kim Y, Hubisz MJ, Clark AG, Bustamante C. Genomic scans for selective sweeps using SNP data. Genome Res. 2005;15:1566–75.16251466
58. Savigamin C Khositharattanakool P Somwang P Wacharapluesadee S Siriyasatien P Preativatanyou K Identification of common human infectious and potentially zoonotic novel genotypes of Enterocytozoon bieneusi in cavernicolous bats in Thailand Parasitol Res 2024 123 233 10.1007/s00436-024-08253-7 38850488
Savigamin C, Khositharattanakool P, Somwang P, Wacharapluesadee S, Siriyasatien P, Preativatanyou K. Identification of common human infectious and potentially zoonotic novel genotypes of Enterocytozoon bieneusi in cavernicolous bats in Thailand. Parasitol Res. 2024;123:233.38850488
59. Li W Xiao L Ecological and public health significance of Enterocytozoon bieneusi One Health 2021 12 100209 10.1016/j.onehlt.2020.100209 33426263
Li W, Xiao L. Ecological and public health significance of Enterocytozoon bieneusi. One Health. 2021;12:100209.33426263
60. Harkins KM Schwartz RS Cartwright RA Stone AC Phylogenomic reconstruction supports supercontinent origins for Leishmania Infect Genet Evol 2016 38 101 109 10.1016/j.meegid.2015.11.030 26708057
Harkins KM, Schwartz RS, Cartwright RA, Stone AC. Phylogenomic reconstruction supports supercontinent origins for Leishmania. Infect Genet Evol. 2016;38:101–9.26708057
61. Barratt J Kaufer A Peters B Craig D Lawrence A Roberts T Isolation of novel trypanosomatid, zelonia australiensis sp nov (kinetoplastida: trypanosomatidae) provides support for a gondwanan origin of dixenous parasitism in the leishmaniinae PLoS Negl Trop Dis. 2017 11 e0005215 10.1371/journal.pntd.0005215 28081121
Barratt J, Kaufer A, Peters B, Craig D, Lawrence A, Roberts T, et al. Isolation of novel trypanosomatid, Zelonia australiensis sp. nov. (Kinetoplastida: Trypanosomatidae) provides support for a gondwanan origin of dixenous parasitism in the Leishmaniinae. PLoS Negl Trop Dis. 2017;11:e0005215.28081121
62. Ruang-Areerate T Ruang-Areerate P Manomat J Naaglor T Piyaraj P Mungthin M Genetic variation and geographic distribution of Leishmania orientalis and Leishmania martiniquensis among Leishmania/HIV co-infection in Thailand Sci Rep 2023 13 23094 10.1038/s41598-023-50604-4 38155252
Ruang-Areerate T, Ruang-Areerate P, Manomat J, Naaglor T, Piyaraj P, Mungthin M, et al. Genetic variation and geographic distribution of Leishmania orientalis and Leishmania martiniquensis among Leishmania/HIV co-infection in Thailand. Sci Rep. 2023;13:23094.38155252
63. Jomkumsing P Surapinit A Saengpara T Pramual P Genetic variation, DNA barcoding and blood meal identification of Culicoides Latreille biting midges (Diptera: Ceratopogonidae) in Thailand Acta Trop 2021 217 105866 10.1016/j.actatropica.2021.105866 33607064
Jomkumsing P, Surapinit A, Saengpara T, Pramual P. Genetic variation, DNA barcoding and blood meal identification of Culicoides Latreille biting midges (Diptera: Ceratopogonidae) in Thailand. Acta Trop. 2021;217:105866.33607064
64. Kamyingkird K Choocherd S Chimnoi W Klinkaew N Kengradomkij C Phoosangwalthong P Molecular Identification of Culicoides species and host preference blood meal in the african horse sickness outbreak-affected area in hua hin district, prachuap Khiri Khan province Thailand Insects 2023 14 369 10.3390/insects14040369 37103184
Kamyingkird K, Choocherd S, Chimnoi W, Klinkaew N, Kengradomkij C, Phoosangwalthong P, et al. Molecular identification of Culicoides species and host preference blood meal in the african horse sickness outbreak-affected area in Hua Hin District, Prachuap Khiri Khan Province, Thailand. Insects. 2023;14:369.37103184
65. Mellor PS Carpenter S White DM Mellor P Baylis M Mertens P Bluetongue virus in the insect host Bluetongue 2009 London Elsevier 295 320
Mellor PS, Carpenter S, White DM. Bluetongue virus in the insect host. In: Mellor P, Baylis M, Mertens P, editors. Bluetongue. London: Elsevier; 2009. p. 295–320.
66. Lobsiger L Müller N Schweizer T Frey CF Wiederkehr D Zumkehr B An autochthonous case of cutaneous bovine leishmaniasis in Switzerland Vet Parasitol 2010 169 408 414 10.1016/j.vetpar.2010.01.022 20153118
Lobsiger L, Müller N, Schweizer T, Frey CF, Wiederkehr D, Zumkehr B, et al. An autochthonous case of cutaneous bovine leishmaniasis in Switzerland. Vet Parasitol. 2010;169:408–14.20153118
67. Müller N Welle M Lobsiger L Stoffel MH Boghenbor KK Hilbe M Occurrence of Leishmania sp in cutaneous lesions of horses in Central Europe Vet Parasitol 2009 166 346 351 10.1016/j.vetpar.2009.09.001 19800739
Müller N, Welle M, Lobsiger L, Stoffel MH, Boghenbor KK, Hilbe M, et al. Occurrence of Leishmania sp. in cutaneous lesions of horses in Central Europe. Vet Parasitol. 2009;166:346–51.19800739
68. Reuss SM Dunbar MD Calderwood Mays MB Owen JL Mallicote MF Archer LL Autochthonous Leishmania siamensis in horse, Florida, USA Emerg Infect Dis 2012 18 1545 1547 10.3201/eid1809.120184 22932732
Reuss SM, Dunbar MD, Calderwood Mays MB, Owen JL, Mallicote MF, Archer LL, et al. Autochthonous Leishmania siamensis in horse, Florida, USA. Emerg Infect Dis. 2012;18:1545–7.22932732
69. Chusri S Hortiwakul T Silpapojakul K Siriyasatien P Consecutive cutaneous and visceral leishmaniasis manifestations involving a novel Leishmania species in two HIV patients in Thailand Am J Trop Med Hyg 2012 87 76 80 10.4269/ajtmh.2012.11-0749 22764295
Chusri S, Hortiwakul T, Silpapojakul K, Siriyasatien P. Consecutive cutaneous and visceral leishmaniasis manifestations involving a novel Leishmania species in two HIV patients in Thailand. Am J Trop Med Hyg. 2012;87:76–80.22764295
