The use of animals in this study was approved by the animal research ethics committee of Chulalongkorn University Animal Care and Use Protocol (CU-ACUP), Faculty of Medicine, Chulalongkorn University, Bangkok, Thailand (COA No. 023/2564 and COA No. 011/2564).

Collection of biting midges was conducted in two locations, Location 1: Tambon Tha Ruea (TR), an endemic area of leishmaniasis (8°22′42.0”N 99°58′32.0″E) and Location 2: Tambon Khuan Phang (KP), a mixed farm of chickens, goats, and cattle (8°09′35.2”N 99°56′33.8″E), Nakhon Si Thammarat province, southern Thailand. In each location, wild biting midges were collected using five Center for Disease Control and Prevention (CDC) miniature light traps (25 W bulb) with ultraviolet (UV) light for 4 nights in December 2021. Traps were operated from 6.00 pm to 10.00 pm. The midges were kept in plastic boxes covered by two layers of insect nets with moisture papers on the top and transported to the Vector Biology and Vector Borne Disease Research Unit, Department of Parasitology, Faculty of Medicine, Chulalongkorn University.

Live insects were placed on ice to immobilize before investigating under a binocular stereoscopic microscope (SZX10; Olympus, Tokyo, Japan). Males were separated from females by their morphology. Species identification of females was carried out using a taxonomic key according to morphological characters, namely wing spot patterns and head features (palp and antenna) (Dyce et al., 2007; Pramual et al., 2021). In each species, females were grouped by their physiological stage: parous (empty abdomen, with traces of burgundy pigment after blood sucking), engorged (abdomen filled with blood), gravid (abdomen with eggs), and nulliparous (empty abdomen without the presence of blood, they never sucked blood) (Dyce, 1996; Kasičová et al., 2021). Parous (50) and nulliparous (50) females collected from each location were selected to investigate for trypanosomatids. Before dissection, the midges were placed on ice in a small Petri dish containing 0.05% (v/v) Tween 20 in 1× Phosphate Buffered Saline (PBS; 10 mM sodium phosphate, 145 mM sodium chloride, pH 7.2). The whole gut (foregut, midgut, and hindgut) was dissected using sterile fine needles in a drop of 50 μl sterile PBS on a sterile slide under a binocular stereoscopic microscope. Then the gut was transferred into 50 μl sterile PBS on a new sterile slide, covered with a sterile coverslip, and examined for trypanosomatids under a light microscope (Olympus America Inc., USA) at 400 × magnification. Some insects with unidentified trypanosomatids in the whole gut were video recorded and photographed. Each positive sample was divided into three parts in a small volume of PBS for Giemsa’s staining, cultivation, and gDNA extraction. To confirm the species of the infected insects, the female carcass of each positive sample was subjected to gDNA extraction for molecular identification of Culicoides species.

Schneider’s Insect medium (SIM) (Sigma-Aldrich, St Louis, MO, USA), pH 6.8 supplemented with 10% (v/v) heat-inactivated fetal bovine serum (hi-FBS) (Life Technologies-Gibco, Grand Island, NY, USA), 100 μg/mL penicillin–streptomycin and 250 μg/mL gentamicin (Sigma-Aldrich, St. Louis, MO, USA) was used to culture trypanosomatids from insects. Each positive sample in PBS was transferred into a 25 ml flask containing 5 mL SIM, pH 6.8 supplemented with 10% (v/v) hi-FBS and 25 μg/ml gentamicin (SIM complete), and incubated at 26°C. An axenic culture was established by serial dilution and subpassage. Promastigotes were mixed with 7.5% (v/v) glycerol in SIM complete and stored at −80°C.

Trypanosomatid samples isolated directly from insects or cultures in SIM with supplements were smeared on microscope slides and air-dried. The slides were then fixed in absolute methanol and stained with 5% (v/v) Giemsa’s stain solution (Sigma-Aldrich, Darmstadt, Germany). The morphological characteristics and morphometry of the trypanosomatids were examined under a light microscope (Olympus America Inc., USA) at 1,000 × magnification. Light microscopy (LM) images of 50 parasites in each stage were used for morphometry, including: cell body length, cell body width, anterior end to kinetoplast distance, anterior end to nucleus distance, and length of flagellum. The measurement results were presented as mean ± standard deviation.

Genomic DNA of biting midges and trypanosomatids was extracted using a genomic DNA purification kit (Thermo Fisher Scientific Inc., Waltham, MA, USA) according to the manufacturer’s instructions. For molecular identification of Culicoides species, LCO1490 primer (5 ́–GGTCAACAAATCATAAAGATATTGG−3 ́) and HCO2198 primer (5 ́–TAAACTTCAGGGTGACCAAAAAA TCA−3 ́) (Folmer et al., 1994) were used to amplify the approximately 658-bp fragment of the mitochondrial cytochrome c oxidase subunit 1 gene (COI) following the polymerase chain reaction (PCR) method described by Harrup et al. (2016). Host blood from engorged females was identified based on the mitochondrial cytochrome b (cyt b) gene sequence using the primers cyt bb1 (5 ́–CCATCMAACATYTCADC ATGAAA−3 ́) and cyt bb2 (5 ́–GCHCCTCAGAATGAYATTTG KCCTCA−3 ́) as described by Radrova et al. (2013). For molecular identification of Leishmania species in gDNA samples extracted from insects and/or cultures, primers LeF (5 ́–TCCGCCCGAAAGTTCACC GATA−3 ́) and LeR (5 ́–CCAAGTCATCCATCGCGACACG−3 ́) (Spanakos et al., 2008), were used to amplify the internal transcript spacer 1 (ITS1) region (approximately 379 bp) and primers, 70-IR-D (5 ́-CCAAGGTCGAGGAGGTCGACTA-3 ́) and 70-IR-M (5 ́-ACG GGTAGGGGGAGGAAAGA −3 ́) (Requena et al., 2012) were used to amplify the 3 ́untranslated region (3 ́UTR) of Leishmania HSP70-type I (HSP70-I) genes as described by Jariyapan et al. (2021).

For molecular identification of trypanosomatids, primers, TRY927F (5 ́–GAAACAAGAAACACGGGAG −3 ́) and TRY927R (5 ́–CTACTGGGCAGCTTGGA−3 ́), were used to amplify approximately 927 bp of the small subunit ribosomal RNA (SSU rRNA) gene (Noyes et al., 1999) of gDNA extracted from insects and cultures. All amplicons were visualized in 1.5% agarose. The PCR products were purified using a GeneJET PCR Purification kit (Thermo Fisher Scientific, CA, USA). PCR conditions and protocols used in this study are provided in Supplementary Data S1.

For trypanosomatids successfully grown in culture, primers M200 (5 ́–ATGGCTCCVVTCAARGTWGGMAT−3 ́) and M201 (5 ́–TAKCCCCACTCRTTRTCRTACCA −3 ́) for the glycosomal glyceraldehyde-3-phosphate dehydrogenase (gGAPDH) gene (Maslov et al., 1996) were used to confirm species identification. The PCR products of M200/M201 primers were cloned into the pGEM-T Easy Vector system (Promega, Madison, WI, USA), following the manufacturer’s instructions. Recombinant plasmid DNA was extracted using the Invisorb Spin Plasmid Mini Kit (STRATEC Molecular, Berlin, Germany), following the manufacturer’s instructions. The purified PCR products were sent for sequencing at the sequencing service of Macrogen Inc., Seoul, Korea.

The nucleotide sequences of the COI gene of Culicoides biting midges, the cyt b gene of vertebrates, ITS1 and 3’UTR-HSP70-I genes of Leishmania parasites, and SSU rRNA and gGAPDH genes of trypanosomatids were analyzed by comparison with the GenBank database using a BLAST search¹. The sequences of Leishmania parasites and trypanosomatids were aligned and trimmed using MEGA X: Molecular Evolutionary Genetics Analysis across computing platforms version 10.2.6 (Kumar et al., 2018). The best-fitting nucleotide substitution models for both genes were estimated using the Akaike Information Criterion (AIC) in the jModelTest software (Posada, 2008). Phylogenetic trees were constructed using the Maximum Likelihood (ML) method implemented in MEGA (Kumar et al., 2018) with 1,000 bootstrap-replications. Bootstrap values of ≥70% were taken as an indication support (Hillis and Bull, 1993). For phylogenetic analysis of Leishmania parasites, evolutionary models of GTR + I were chosen for ITS1 (329 bp) and 3’UTR-HSP70-I (817 bp). For phylogenetic analysis of Crithidia sp., evolutionary models of GTR + I + G were chosen for SSU rRNA (720 bp) and gGAPDH (654 bp). Genetic distances were calculated using the Kimura 2-parameter (K2P) model implemented in MEGA (Kumar et al., 2018).

Scanning electron microscopy (SEM) and transmission electron microscopy (TEM) were used to characterize the ultrastructural morphology of the cultured trypanosomatids. For SEM, live trypanosomatids were pelleted at 1,600 × g for 10 min at room temperature and fixed with 2.5% (v/v) glutaraldehyde in 0.1 cacodylate buffer (pH 7.2) overnight at 4°C. Fixed cells were washed twice in PBS and post-fixed with 1% (w/v) osmium tetroxide in PBS for 1 h. The samples were then dehydrated in a graded series of ethanol. Samples were critical point dried in liquid CO₂ and coated with gold particles in a sputter-coating apparatus. Samples were observed under a scanning electron microscope, JEOL JSM-6610LV (JEOL, Tokyo, Japan), operated at 15 kV. Samples for TEM were prepared as for SEM, except that 2% (w/v) osmium tetroxide were used to post-fix for 2 h at room temperature. After dehydration in a graded series of ethanol, the samples were incubated overnight in an epoxy resin (PolyBed 812)/acetone solution (1:1), and then embedded in pure resin and polymerized for 48 h at 60°C. ultra-thin sections were stained with 4% (w/v) uranyl acetate and 3% (w/v) lead citrate and observed under a transmission electron microscope, JEM-2200FS (JEOL, Tokyo, Japan), operated at 200 kV.

Trypanosomatids were removed from −80°C, grown in SIM complete at 26°C for 4 days, and subpassaged into new SIM complete for growth analysis. The parasites were counted using a Neubauer chamber (BLAUBRAND, Sigma-Aldrich, Saint Louis, MO, USA) from an initial inoculum of 4 × 10⁴ parasites/mL (day 0). The initial inoculum was transferred into a 25 cm³ flask containing 5 ml of the culture medium for 14 flasks. Seven flasks were incubated at 26°C and the rest were incubated at 37°C with 5% CO₂. One flask at each temperature was taken out daily, and a cell scraper (SPL Life Sciences, Gyeonggi, Korea) was applied to remove any adherent cells from the base of the flask. Ten microliters of parasites were collected and mixed at a 1:1 ratio with formaldehyde when all parasites were alive or 0.4% trypan blue solution (Thermo Fisher Scientific, NY, USA) when nonviable forms of parasites were present in the cultures. Live cells were counted using a Neubauer chamber. Also, the cultures were smeared on slides and stained with Giemsa’s staining solution for morphological examination by LM. Experiments were performed on three independent replicates run in duplicate.

To evaluate the ability of trypanosomatids to infect and multiply in mouse macrophages an infection and multiplication assay was performed. Mouse peritoneal exudate macrophages (PEMs) from BALB/c (Mus musculus) (Nomura Siam International Co., Ltd., Bangkok, Thailand) were obtained using the method described previously (Zhang et al., 2008). Round coverslips were placed in 24-well tissue culture plates (ThermoFisher Scientific, Jiangsu, China). PEMs in RPMI 1640 medium (GE Healthcare Life Science-HyClone, UT, United States) supplemented with gentamicin 25 μg/ml and 10% (v/v) hi-FBS were seeded at a density of 2.5 × 10⁵ cells/well and incubated at 37°C with 5% CO₂ for 24 h. After the incubation, non-adherent cells were removed by washing three times with pre-warmed serum-free RPMI 1640 medium. Test trypanosomatids (from Day 3 with the greatest number of motile forms) and L. martiniquensis promastigotes (from Day 5) were used to infect PEMs at a ratio of 10:1 cells to macrophages. After incubation for 3 h at 37°C and 5% CO₂, the coverslips were washed with pre-warmed serum-free RPMI 1640 medium for three times, replaced with 10% hi-FBS RPMI-1640 medium, and incubated at the same conditions. Coverslips were removed from the culture plates and stained with Giemsa’s staining solution every 24 h until 96 h post-infection. Two hundred macrophages were counted to determine the infection rate and average number of intracellular parasites per macrophage. The infection index, and the intracellular parasite multiplication ratio were determined (Chanmol et al., 2019). Results were expressed as mean ± standard deviation and based on three independent infection experiments each performed in duplicate.

All statistical analyses were performed using GraphPad Prism version 9.1 software. Statistical differences between L. martiniquensis and Crithidia sp. infections at the different time points within one group were determined using two-way ANOVA with Bonferroni’s post hoc multiple comparisons for growth, infection index, and intracellular multiplication ratio. Tests were considered statistically significant if p < 0.05.

A total of 1,094 biting midges of the genus Culicoides were captured in 2 locations, 349 females (parous = 276, engorged = 2, nulliparous = 71) and 6 males from Tha Ruea and 724 females (parous = 527, engorged = 12, nulliparous =185) and 15 males from Khuan Phang (Supplementary Table S1). No gravid females were found in either location. Three species of biting midges were identified in Tha Ruea, C. peregrinus (n = 343, 96.62%), C. mahasarakhamense (n = 9, 2.53%), and C. oxystoma (n = 3, 0.85%). In Khuan Phang seven species were identified, C. peregrinus (n = 623, 84.3%), C. imicola (n = 31, 4.19%), C. shortti (n = 27, 3.65%), C. huffi (n = 25, 3.38%), C. mahasarakhamense (n = 20, 2.7%), C. palpifer (n = 11, 1.5%), and C. oxystoma (n = 2, 0.27%). Thus, in both locations C. peregrinus was the most abundant species. For engorged females (n = 14), all blood host identifications were of Bos indicus cattle (GenBank accession no. OR088254–OR088267).

Fifty parous and 50 nulliparous females collected from each location were dissected for trypanosomatid infection. Trypanosomatids were observed under light microscopy (LM) in 26 out of the 100 samples of parous females from the two locations. However, no trypanosomatids were found in any nulliparous flies. All infected flies were identified as C. peregrinus by wing spot patterns (Figure 1A) and molecular methods (GenBank accession no. OR077408–OR077433). Light microscopic examination showed trypanosomatids present in the digestive tract of the midges (Figure 1 and Supplementary Videos S1, S2). None of the midges dissected had remains of bloodmeals in their midguts. Although the midges were dissected carefully, due to their small size and delicate nature, the digestive tracts of some midges were torn and trypanosomatids released. However, in most samples with an intact digestive tract trypanosomatids were found in the midgut and hindgut, with choanomastigote and/or promastigote morphologies observed (Figure 1B). Interestingly, trypanosomatids were observed in the foregut, midgut, and hindgut of one C. peregrinus, KP10 (Figure 1C). In this insect, video records revealed typical movement of Leishmania promastigotes in the foregut (Supplementary Video S1) and Crithidia in the hindgut (Supplementary Video S2). Molecular identification (see below) of the KP10 sample revealed that this midge was co-infected with L. martiniquensis and a possible novel Crithidia species (Table 1). Trypanosomatids were examined in slides prepared from infected midges, and various forms of promastigotes and chroanomastigotes were observed (Figure 1D). In the insect KP19 many promastigote forms of Leishmania were observed including procyclic promastigotes (Figures 1E,F), nectomonad promastigotes (Figure 1G), and leptomonad promastigotes (Figure 1H), however, no metacyclic promastigotes were observed. The parasites in KP19 were identified as L. martiniquensis by molecular methods (Table 1). The presence of Leishmania promastigotes in the foregut of midges TR17, KP10, KP17 and KP19 in the absence of bloodmeals are indicative of established infections.

Four isolates of trypanosomatids were successfully isolated into culture, two from C. peregrinus collected in Tha Ruea (TR2 and TR3) and two from C. peregrinus collected in Khuan Phang (KP1 and KP4) (Table 1). Unfortunately, other cultures including all those from midges with L. martiniquensis were too contaminated with bacteria and fungi and had to be discarded.

Molecular methods were used to identify the trypanosomatids. For molecular identification of Leishmania species, 100 gDNA samples extracted from the dissected insects were subjected to the PCR amplification of the ITS1 and 3’UTR-HSP70-I regions. Four samples of the dissected insects, from TR17, KP10, KP17, and KP19, were positive for Leishmania and their ITS1 amplicons were successfully sequenced. Moreover, a second molecular marker, 3’UTR-HSP70-I, was used to confirm the identification of the Leishmania species of the samples. BLAST analysis of the ITS1 and 3’UTR-HSP70-I sequences of the samples revealed that the ITS1 sequences were closest to that of L. martiniquensis, GenBank accession number MK603827.1, with 98.94% identity, whereas the 3’UTR-HSP70-I sequences were very similar (98.64 and 99.77% identity) or identical (100% identity) to that of L. martiniquensis, GenBank accession number MK607435.1, and another one was identical to L. martiniquensis, GenBank accession number MK607437.1 (Table 1). The ITS1 and 3’UTR-HSP70-I sequences were subjected to phylogenetic analyses together with other human Leishmania strains (Figure 2). The results from Maximum Likelihood tree based on amplified section of the ITS1 and 3’UTR-HSP70-I sequences showed that the new sequences fell in one clade with L. martiniquensis with 99% bootstrap values (K2P 0–0.78%) for ITS1 (Figure 2A) and 100% bootstrap values (K2P 0–1.63%) for 3’UTR-HSP70-I (Figure 2B). These results indicate that the trypanosomatid DNAs detected in the C. peregrinus samples, TR17, KP10, KP17, and KP19, were all of L. martiniquensis.

The SSU rRNA of 24 samples were successfully amplified and sequenced allowing the identification of other trypanosomatids. BLAST analysis showed that among these, 19 sequences were similar to each other and to a Crithidia sp. reported from Thailand by Songumpai et al. (2022) (GenBank accession number OP698037–OP698038) with 99.35–100% identity and to Cr. thermophila (GenBank accession number KY264937.1) with 97.00–97.74% identity. The other five sequences were also very similar to another Crithidia sp. reported from Thailand by Songumpai et al. (2022) (GenBank accession number OP217136) and Sunantaraporn et al. (2022) (GenBank accession number MW694347.1) with 99.89–100% and 98.92–99.14% identity, respectively (Table 1). These data showed that two midges, TR17 and KP10, were co-infected with both L. martinquensis and Crithidia, as suspected from dissection for KP10.

Phylogenetic analysis of the SSU rRNA sequences demonstrated that representative sequences from the group of 19, namely from insects TR2, TR3, KP1, TR17, KP4, and KP10 clustered together into one distinct clade with 99% bootstrap values (K2P 0.19–0.77%), designated here as Crithidia sp. Clade A (Figure 3A). The corresponding isolates were thus given strain designations Crithidia sp. CLA-TR2, Crithidia sp. CLA-TR3, and so on. Four of these, CLA-TR2, CLA-TR3, CLA-KP1 and CLA-KP4, were the four isolates successfully isolated into culture. Representatives of the group of 5 from insects TR21, TR29, and KP47 also clustered together with 99% bootstrap values (K2P 0–1.35%), and were designated as Crithidia sp. Clade B. The availability of cultures for CLA-TR2, CLA-TR3, CLA-KP1, and CLA-KP4 enabled additional DNA extractions and analysis of their gGAPDH sequences. These gGAPDH sequences also clustered together with 98% bootstrap values (K2P 0.31–0.77%) (Figure 3B). Amongst available sequences they were most similar to those of Cr. fasciculata (GenBank accession number AF047493.1) and Cr. brachyflagelli (GenBank accession number JF717835.1) with 94.39–94.61% identity (Table 1). However, they were almost as close to a clade of Cr. thermophila sequences (K2P 3.77–4.25%), like what was observed in the SSU rRNA analysis (Figure 3A).

In summary, from Tha Ruea seven female C. peregrinus were found infected with Crithidia sp. Clade A organisms and four females with Crithidia sp. Clade B organisms. Successful cultures were obtained only from insects TR2 and TR3, both Clade A organisms (Table 1). In this area the infection rate of L. martiniquensis in the 50 parous C. peregrinus was 2%, represented by one insect (TR17) also co-infected with Crithidia sp. Clade A organisms. In Khuan Phang, twelve females were infected with Crithidia sp. Clade A organisms and one female with Crithidia sp. Clade B organisms. In this area the infection rate of the C. peregrinus with L. martiniquensis was 6%, with one insect (TR10) again co-infected with Crithidia sp. Clade A organisms, but also in two other insects (KP17 and KP19) that only contained Leishmania. We also successfully cultured two Crithidia sp. Clade A from two C. peregrinus (KP1 and KP4) (Table 1).

The results of the phylogenetic analysis revealed Crithidia sp. Clade A organisms to be related to Cr. thermophila, an unusual thermotolerant trypanosomatid. To examine whether Clade A organisms might have similar properties further experiments were performed on strain CLA-KP1. Morphological and ultrastructural analyses of cultures in SIM complete at 26°C revealed two distinct morphotypes, choanomastigotes with rounded posterior ends and promastigotes with elongated posterior ends (Figures 4, 5). A wide collar-shaped reservoir (collar-like extension) surrounding the anterior end through which a single flagellum emerges was noted in both morphotypes (Figures 4A,B, 5B). This single flagellum arose from a basal body (kinetosome) located next to a kinetoplast (Figures 5A–C) and was surrounded by the flagellar pocket (Figure 5). The flagellum exhibited nine pairs of peripheral microtubules surrounding two central microtubules, the typical 9 × 2 + 2 axonemal pattern (Figure 5D). The kinetoplast was located just beyond the basal body, anterior or parallel to the nucleus. The nucleus was oval located in the middle of the cell body with visible accumulations of chromatin. Some other structures, such as acidocalcisomes, glycosomes and ribosomes, were observed (Figures 5A–C). Different forms of the choanomastigotes were observed, including those without a free flagellum (haptomonad forms) (Figures 4A–F, 5A) and others with a free flagellum protruding from the flagellar pocket at the anterior end of the cells (Figures 4B–F). A form of choanomastigote comprising attached clusters of cells (Rosette form) could be observed at the bottom of culture flasks (Figures 4A–E). Promastigotes were slender in shape with a free flagellum at the anterior end of the cells. In addition, morphological differences in some details, such as body width, body length, and flagellum length were observed (Figures 4G–J). Comparative morphological measurements of the choanomastigotes and promastigotes determined under LM are shown in Table 2. Promastigotes with the longest body and flagellum length known as a nectomonad form were noted (Figures 4D–J and Table 2). A single layer of subpellicular microtubules was observed around the body except for the membrane inside the area of the flagellar pocket (Figures 5A–C).

CLA-KP1 organisms were cultured at 26 and 37°C with an initial cell density of 4 × 10⁴ cells/ml and observed daily for their growth for seven successive days. On the first 4 days, at both temperatures, similar growth patterns were observed. The parasites entered the exponential growth phase after 1 day of the culture and showed rapid multiplication (day 1 to day 4) with a doubling time of approximately 13.5 and 15.6 h for 26°C and 37°C, respectively. After 4 days of culture, at 26°C, the cultures entered the stationary phase and reached their peak on day 6 with a density of 1.9 × 10⁸ cells/ml. The cultures remained in this phase to the last day of the experiments. At 37°C, CLA-KP1 reached maximum growth on day four with a density of 9.8 × 10⁷ cells/mL, remained in stationary phase for 2 days, then the population dropped dramatically after 6 days of culture until the end of the experiment with a density of 1.0 × 10⁶ cells/ml. The average parasite density at 26°C was significantly greater than 37°C after 5 days of culture (Figure 6 and Supplementary Table S2). Both morphotypes were found in the cultures at both temperatures. However, the predominant morphotype at 26°C was promastigotes, whereas at 37°C choanomastigotes were mainly observed.

The ability of CLA-KP1 to infect and multiply in PEMs of BALB/c mice was determined and compared with L. martiniquensis (Figure 7). Crithidia sp. CLA-KP1 could infect the PEMs and was found at 24 and 48 h but no parasites were observed at 72 and 96 h. At both time points, the infection rate, the average number of intracellular parasites per cell, and the infection index of CLA-KP1 were statistically significantly lower than that of L. martiniquensis (Figures 7H–J). At 24 h post-infection, the percentage of PEMs infected with CLA-KP1 was 22.27 ± 0.55 and dramatically decreased to 6.68 ± 1.85 at 48 h, which differed from the stable infection rate in Leishmania parasites at the same time points. The average number of intracellular CLA-KP1 parasites within infected PEMs was 2.35 ± 0.09 and 1.56 ± 0.17 parasites/cell at 24 and 48 h, respectively. At both time points, the average number of intracellular parasites/macrophage was approximately 2.6 times lower than that of L. martiniquensis. The infection index of CLA-KP1 was 51.97 ± 0.45 and 10.25 ± 2.10 at 24 h and 48 h, respectively, which was lower than that of the Leishmania parasites approximately 3.5 and 12.5 times, respectively. The intracellular parasite multiplication ratio of CLA-KP1 was only calculated at 48 h post-infection (0.21) because after that the parasites disappeared. A decrease in the intracellular parasite multiplication ratio was also observed in L. martiniquensis after 24 h post-infection, however, the Leishmania parasites were still be found in the PEMs at the end of the experiment (96 h) (Figure 7 and Supplementary Table S3). In summary, these results demonstrated that Crithidia sp. CLA-KP1 could infect and persist in the PEMs up to 48 h but could not multiply within them.