Introduction
The nematode genus Kalicephalus Molin, 1861, includes 35 species that parasitise snakes and, rarely, lizards, with a cosmopolitan distribution (Chandel and Dolma Reference Chandel and Dolma2025; Haick et al. Reference Haick, Tavares-Costa, Rebêlo, Neves, Félix, Müller, Trinidade, Costa-Campos, Maschio and Melo2025). Despite the broad diversity and wide distribution of the genus Kalicephalus, only five species and one unidentified species have available genetic sequences. In the Neotropical region, eight species have been reported (Haick et al. Reference Haick, Tavares-Costa, Rebêlo, Neves, Félix, Müller, Trinidade, Costa-Campos, Maschio and Melo2025): Kalicephalus subulatus Molin, 1861; Kalicephalus inermis coronellae Ortlepp, 1923; Kalicephalus costatus costatus (Rudolph, 1819); Kalicephalus appendiculatus Molin, 1861; Kalicephalus rectiphilus neorectiphilus Schad, Reference Schad1962; Kalicephalus inermis inermis Molin, 1861; Kalicephalus inermis macrovulvus (Caballero, 1954), and Kalicephalus atroxi Haick, Tavares-Costa & Melo, Reference Haick, Tavares-Costa, Rebêlo, Neves, Félix, Müller, Trinidade, Costa-Campos, Maschio and Melo2025. Of these species, only K. atroxi and K. c. costatus from the state of Amapá, Brazil have genetic sequences.
The knowledge of the life cycle and transmission routes of Kalicephalus is limited (Anderson Reference Anderson2000). Experimentally, it has been documented that snakes can be infected by oral inoculation with the third-stage larvae (Schad Reference Schad1956). Larval penetration through the skin has been documented; however, it does not appear to be the primary route of infection compared with oral transmission, as evidenced by the low success rate of experimental infections (Schad Reference Schad1956). Another possibility that has been proposed is the involvement of paratenic hosts (e.g., slugs, amphibians) (Anderson Reference Anderson2000); however, this has not yet been demonstrated. Today, the mechanisms by which snakes become infected with Kalicephalus under natural conditions are still unknown.
The Yucatan Neotropical rattlesnake Crotalus tzabcan Klauber, 1952 is a large snake, endemic to the Mexican states of Campeche, Quintana Roo, Yucatan, Chiapas, and Tabasco, as well as northern portions of Belize and Guatemala (Carbajal-Márquez et al. Reference Carbajal-Márquez, Cedeño-Vázquez, González-Solís and Martins2020). This species is listed as Least Concern by the International Union for Conservation of Nature. In Mexico, it is not under any level of protection.
The knowledge of endoparasites of C. tzabcan is limited. The list of helminths and pentastomids associated with C. tzabcan includes Porocephalus crotali Humboldt, 1808 (Pentastomida) (Carbajal-Márquez et al. Reference Carbajal-Márquez, González-Solís and Cedeño-Vázquez2018; González-Solís and Terán-Juárez Reference González-Solís and Terán-Juárez2013), Hastospiculum onchocercum Chitwood, 1932, Hexametra boddaertii (Baird, 1860), and Travassosascaris araujoi Sprent, 1968 (Nematoda) (Carbajal-Márquez et al. Reference Carbajal-Márquez, González-Solís and Cedeño-Vázquez2018). The aim of this study was to provide morphological and molecular data for nematodes recorded from C. tzabcan in Mexico.
Materials and methods
Sample collection and morphological identification
On 15 September 2025, an adult male C. tzabcan was found dead after being run over on a road in Hunucmá, Yucatan, Mexico. In the field, viscera were collected and maintained for approximately 8 h at 4°C until laboratory examination. Viscera were immersed in 0.9% sodium chloride solution and examined under a stereomicroscope. Seventy female nematodes were found in the intestines of the rattlesnake. They were fixed with hot 70% ethanol, counted, and preserved in 70% ethanol for light microscopy, 10% formalin for scanning electron microscopy (SEM), and 100% ethanol for DNA extraction.
For morphological examination, 16 nematodes were cleared and temporarily mounted in lactophenol. Specimens were studied and drawn using a Leica DM750 light microscope equipped with a drawing tube. For SEM micrographs, one specimen was studied at the Laboratorio de Microscopía y Fotografía de la Biodiversidad, Instituto de Biología, Universidad Nacional Autónoma de México (IBUNAM), Mexico City. The specimen was dehydrated using a graded ethanol series, critical-point dried with carbon dioxide, sputter-coated with a gold-palladium mixture, and examined at an accelerating voltage of 10 kV with a Hitachi SU1510 scanning electron microscope. Measurements are presented as mean, followed by standard deviation and the range in parentheses. These measurements are given in micrometres (μm) unless otherwise stated. Voucher specimens were deposited in the Colección Nacional de Helmintos (CNHE), IBUNAM, Mexico City, Mexico.
Molecular analysis
Total genomic DNA of two specimens was extracted using the DNeasy Blood and Tissue Kit (Qiagen, Hilden, Germany) following the manufacturer’s protocol. Three molecular genetic markers were targeted: the cytochrome c oxidase subunit 1 (COI) gene, the internal transcribed spacer (ITS) region, and the 28S rRNA gene (28S). A fragment of the COI gene was amplified using primers LCO1490 and HC02198 (Folmer et al. Reference Folmer, Black, Hoeh, Lutz and Vrijenhoek1994). Partial sequences of the ITS region were amplified using the primers NC5 and NC2 (Zhu et al. Reference Zhu, Gasser, Podolska and Chilton1998). A fragment of the 28S rRNA gene was amplified using the primers 391 (Nadler et al. Reference Nadler, Carreno, Adams, Kinde, Baldwin and Mundo-Ocampo2003) and 536 (García-Varela and Nadler Reference García-Varela and Nadler2005). Thermal conditions of the polymerase chain reaction (PCR) amplifications were those described by Folmer et al. (Reference Folmer, Black, Hoeh, Lutz and Vrijenhoek1994), Zhu et al. (Reference Zhu, Gasser, Podolska and Chilton1998), and Hernández-Mena et al. (Reference Hernández-Mena, García-Varela and Pérez-Ponce de León2017) for COI, ITS, and 28S, respectively. The PCR primers, along with additional internal primers 503 (Nadler et al. Reference Nadler, Carreno, Adams, Kinde, Baldwin and Mundo-Ocampo2003) and 504 (Hernández-Mena et al. Reference Hernández-Mena, García-Varela and Pérez-Ponce de León2017) for 28S, were used for Sanger sequencing at Macrogen (Seoul, Korea). Contiguous sequences were assembled and edited using Geneious Pro 4.8.4 (Kearse et al. Reference Kearse, Moir, Wilson, Stones-Havas, Cheung, Sturrock, Buxton, Cooper, Markowitz, Duran, Thierer, Ashton, Meintjes and Drummond2012).
Phylogenetic analyses were inferred using newly generated sequences of Kalicephalus sp. together with Ancylostomatina sequences retrieved from GenBank. For the ITS region, trimmed sequences of the ITS1 were used. Oswaldocruzia filiformis (Goeze, 1782) was selected as the outgroup based on its phylogenetic distance from members of the families Ancylostomatidae and Diaphanocephalidae (Haick et al. Reference Haick, Tavares-Costa, Rebêlo, Neves, Félix, Müller, Trinidade, Costa-Campos, Maschio and Melo2025). Newly obtained and previously published sequences were aligned using ClustalW implemented in MEGA 11. The alignment was trimmed at both ends to the length of the shortest sequence, and positions that could not be aligned unambiguously were excluded from the analysis. Phylogenetic relationships were inferred using the maximum-likelihood (ML) method implemented in MEGA 11. The nucleotide substitution model with the best fit was determined using the ‘Find Best DNA/Protein Models’ function under the Bayesian Information Criterion (BIC), which selected the Tamura 3-parameter model with a proportion of invariant sites (T93+I) as the optimal model. Node support was assessed using 1,000 bootstrap replicates. Bayesian phylogenetic inference was conducted in MrBayes v3.2.6 under the general time-reversible (GTR)+I+G substitution model (nst = 6; rates = invgamma). Two independent runs with four Markov chains each were performed for 5,000,000 generations, sampling every 1,000 generations. Convergence was evaluated by examining the average standard deviation of split frequencies, with values below 0.01 considered indicative of adequate mixing. The first 25% of sampled trees were discarded as burn-in, and a majority-rule consensus tree was generated from the remaining trees. Posterior probabilities were used to estimate clade support.
Pairwise genetic distances among verified Kalicephalus sequences available in GenBank were calculated separately for each dataset in MEGA 11 using the uncorrected p-distance model, with pairwise deletion applied to positions containing gaps or missing data.
Results and discussion
Kalicephalus sp.
Taxonomic summary
Host: Crotalus tzabcan Klauber, 1952 (Squamata: Viperidae).
Material deposited: Three female specimens (CNHE-12563) deposited at CNHE.
Locality: Municipality of Hunucmá, state of Yucatan, Mexico (20°59′57.03″N, 89°49′57.53″W).
Site of infection: Intestines.
GenBank accession numbers: 28S (PX972675, PX972676), COI (PX975252), ITS (PX972677, PX972678).
General. Anterior and middle part of body almost uniform in width. Anterior end slightly narrower than the rest of the anterior part of body. Posterior part of body tapering. Anterior edge slightly tilted ventrally. Body cuticle finely transversely striated. Cephalic inflation present. Corona radiata absent. Buccal capsule with two valves formed by chitinous pieces, each valve with an anterior transverse ridge and three parenchymatous extensions (Figure 1a), and without membranous cuticular processes. Anterior plates of buccal capsule valves thin. Oral opening with two pairs of lateral papillae and one amphid on each side, positioned at the median portion of the oral margin (Figure 1b). Oesophagus claviform, muscular, wider in bulbar region. Nerve ring anterior to the mid-region of oesophagus. Excretory pore at the level of the posterior part of the oesophageal bulb (Figure 1a). Deirids minute, situated slightly posterior to the excretory pore. Vulva postequatorial, with slightly raised lips (Figure 1c, d). Amphidelphic uterus (Figure 1c). Eggs in ovejector and vagina in morula stage (Figure 1c). Second- and third-stage larvae present in the ovejector and uteri of some specimens (Figure 1c). Tail long, gradually tapering to a conoid tip (Figure 1e).
Line drawings (a, c, e, f) and SEM micrographs (b, d) of Kalicephalus sp. from Crotalus tzabcan. (a) anterior part, right lateral view. (b) head, apical view (arrowhead = cephalic papillae; asterisks = amphids). (c) vulvar region showing ovejector with eggs and a larva, right lateral view. (d) vulvar region, ventral view (arrow = vulva). (e) posterior part, right lateral view. (f) third-stage larva, left lateral view.
Figure 1. Long description
A six-panel anatomical study of a nematode.
* Panel a: A line drawing of the anterior part in right lateral view. It shows the buccal capsule with internal teeth-like structures and a muscular esophagus. A vertical scale bar indicates 200 micrometers.
* Panel b: An S E M micrograph of the head in apical view. Four white arrowheads point to cephalic papillae arranged around the central mouth opening. Two asterisks mark the lateral amphids. A vertical scale bar indicates 50 micrometers.
* Panel c: A line drawing of the vulvar region in right lateral view. The internal ovejector contains several oval eggs and one coiled larva. A vertical scale bar indicates 200 micrometers.
* Panel d: An S E M micrograph of the vulvar region in ventral view. A white arrow points to the transverse slit of the vulva on the textured body surface. A vertical scale bar indicates 200 micrometers.
* Panel e: A line drawing of the posterior part in right lateral view. It shows the tapering tail and internal intestinal structures. A vertical scale bar indicates 200 micrometers.
* Panel f: A line drawing of a third-stage larva in left lateral view. The entire body is shown in a curved posture, revealing the internal digestive tract. A vertical scale bar indicates 200 micrometers.
Females. Body 9.3 ± 1.1 (7.7–11.6) mm long and 313.1 ± 54.2 (230–430) in maximum width. Buccal capsule 121.6 ± 17.2 (100–150) deep; head diameter 129.5 ± 16.7 (100–160). Oesophagus 325.2 ± 31.8 (290–385) long and 142.2 ± 11.1 (125–162) in maximum width at the bulb region. Nerve ring, excretory pore, and deirids were located at 236.2 ± 52.8 (195–400), 381.9 ± 36.3 (340–460), and 420.4 ± 38.9 (357–490) from apex, respectively. Vulva situated at 3.6 ± 0.6 (2.6–4.5) mm from caudal extremity, with a vulvar ratio of 1.6 ± 0.2 (1.2–1.9). Vagina vera 42.5 ± 8.9 (30–50) long, vestibule 156.7 ± 22.7 (120–200) long, sphincter 41.2 ± 10.4 (20–50) long, infundibulum 168 ± 29.2 (130–220) long. Tail 408.6 ± 39.4 (340–472). Eggs 62.8 ± 9.1 (51–70) long and 42 ± 2.2 (40–45) wide.
Larvae. Third-stage larvae (n = 2) 1,110 and 1,090 long and 38 and 40 wide (Figure 1f). Oesophagus 180 and 190 long. Genital primordium situated at 655 and 725 from anterior extremity. Tail 70 and 90 long. Second-stage larvae (n = 2) 720 long and 22 and 27 wide. Oesophagus 140 and 150 long. Genital primordium situated at 420 and 440 from apex. Tail 60 long.
Five nucleotide sequences (two ITS, two 28S rRNA, and one COI) of Kalicephalus sp. from C. tzabcan were generated and used to calculate pairwise genetic distances. Phylogenetic analyses were performed only using the ITS dataset because of the limited number of sequences available for the other genetic markers (Supplementary Table S1). For the molecular analyses, only GenBank sequences with verified status were included. Sequences of Kalicephalus brachycephalus Maplestone, 1931 and Kalicephalus indicus Ortlepp, 1923 were excluded because they were classified as ‘unverified genome assembly’.
The amplified ITS from C. tzabcan yielded two sequences of 810 and 774 bp (PX972677, PX972678), with no nucleotide variation in the aligned region (ITS1). The ITS1-based phylogeny recovered well-supported clades corresponding to Ancylostomatidae and Diaphanocephalidae. The newly generated sequences from C. tzabcan clustered within the genus Kalicephalus, forming a distinct lineage clearly separated from Diaphanocephalus galeatus (Rudolphi, 1819) (Diaphanocephalidae) (Figure 2). The C. tzabcan-derived sequences were grouped with Kalicephalus viperae (Rudolphi, 1819) from Iran, with high support (bootstrap = 100, posterior probability = 0.99) (Figure 2). The genetic distances between Kalicephalus sp. and K. viperae were 2%. This subclade was recovered as sister to the group formed by K. atroxi and K. c. costatus, both from Brazil.
Phylogeny of Ancylostomatina inferred from ITS1 sequences. The positions of Kalicephalus sp. from Crotalus tzabcan are indicated in bold. GenBank accession numbers are shown after each taxon. Nodal support values are given as maximum-likelihood bootstrap percentages followed by Bayesian posterior probabilities; values >70% and >0.90, respectively, are shown.
Figure 2. Long description
A horizontal rectangular cladogram with a scale bar of 0.05 at the bottom left.
* The top section is highlighted in yellow and labeled D I A P H A N O C E P H A L I D A E. It contains two main branches. The upper branch leads to Diaphanocephalus galeatus (M N 5 9 2 6 7 5). The lower branch, supported by a value of 100/0.99, splits into two sub-clades. The first sub-clade contains Kalicephalus sp. (P X 9 7 2 6 7 7) and Kalicephalus sp. (P X 9 7 2 6 7 8) in bold, and Kalicephalus viperae (M T 4 6 5 4 5 0). The second sub-clade, supported by 87/0.99, contains Kalicephalus atroxi (P V 7 8 8 9 6 5), Kalicephalus atroxi (P V 7 8 8 9 6 7), and Kalicephalus costatus costatus (P V 7 8 8 9 6 6).
* The middle section is highlighted in pink and labeled A N C Y L O S T O M A T I D A E. Supported by a nodal value of 100/0.98, it contains five taxa: Ancylostoma braziliense (J Q 8 1 2 6 9 3), Ancylostoma ceylanicum (L C 0 3 6 5 6 7), Ancylostoma duodenale (E U 3 4 4 7 9 7), Ancylostoma caninum (A M 8 5 0 1 0 6), and Ancylostoma tubaeforme (J Q 8 1 2 6 9 1).
* The bottom section is labeled Outgroup and consists of a single lineage: Oswaldocruzia filiformis (M G 5 8 6 0 8 8).
Sequencing of the 28S rRNA gene from C. tzabcan yielded two sequences (PX972675, PX972676) of 1,105 and 1,067 bp, respectively. The aligned region showed no nucleotide variation between the two isolates. These sequences of Kalicephalus sp. showed a 12% genetic distance from another sequence named ‘Kalicephalus cristatus’ from Australia (AM039741), the only available sequence from GenBank for this gene. However, the latter sequence probably corresponds to Kalicephalus costatus indicus Ortlepp, 1923, as discussed by Pereira et al. (Reference Pereira, Ferreira, Tomas, Elisei, Paiva and Tavares2018).
The COI sequence obtained from C. tzabcan yielded a fragment of 655 bp (PX975252). Pairwise comparison revealed a genetic distance of 12% from two available sequences (KC130688, KC130691) of Kalicephalus sp. from Mexico.
The morphology of the vulva and tail of the female specimens distinguishes Kalicephalus sp. collected from C. tzabcan from the majority of Kalicephalus species reported from the Neotropics. Kalicephalus costatus costatus, K. subulatus, K. appendiculatus, K. i. inermis, K. i. macrovulvus, and K. atroxi have a pedunculate vulva on a prominent hump or vulvar lips slightly raised in a shallow depression or prominent vulvar lips, while Kalicephalus sp. has vulvar lips slightly raised. In addition, K. c. costatus, K. appendiculatus, K. r. neoctiphilus, K. i. inermis, and K. atroxi feature a short tail with a terminal spike, expansion, or knob before the tip, or a long tail that abruptly rounds at the tip or ends in a spine, whereas Kalicephalus sp. has a long tail, gradually tapering to a conoid tip. The vulva and tail morphology of our specimens resembles K. i. coronellae. The morphometric measurements of females of this species vary considerably depending on the host (Schad Reference Schad1962). Therefore, to confirm the identity of this taxon, male specimens should be collected to characterise the spicules and bursal rays.
This is the first record of the genus Kalicephalus in C. tzabcan. In Mexico, this nematode genus has been reported in two rattlesnake species. Comroe (Reference Comroe1948) described Kalicephalus conoides Comroe, Reference Comroe1948, a synonym of K. i. coronellae, from Crotalus triseriatus (Wagler, 1830) in Michoacán. del Prado Vera (Reference del Prado Vera1971) reported Kalicephalus inermis macrovulvus Caballero, 1954 from Crotalus molossus Baird & Girard, 1853 in Mexico City.
In this study, second- and third-stage larvae of Kalicephalus were found in the ovejector and uteri of adult worms. A similar finding was reported in an in vitro study on the survival of Kalicephalus willeyi Linstow, 1904. Ahmed and Saleh (Reference Ahmed and Saleh1967) reported that, in culture, first-stage larvae of K. willeyi invaded female worms through the vulva and survived and completed their developmental stages in 3–5 days, thriving on the uterine contents and adjacent organs. This indicates that uterine contents may provide highly nutritive elements for the propagation of Kalicephalus larvae (Ahmed and Saleh Reference Ahmed and Saleh1967). This trait may be linked to specific environmental conditions (e.g., limited nutrient availability) or to an alternative life cycle in the host, in which third-stage larvae are released to facilitate autoinfection or transmission to new hosts. However, further field and laboratory studies are required to test this hypothesis.
The phylogenetic analyses showed that Kalicephalus sp. from C. tzabcan was grouped with K. viperae from Iran, with high support (bootstrap = 100, posterior probability = 0.99), whereas sequences from Neotropical species (K. atroxi and K. c. costatus) were grouped in another subclade. The genetic distance between the species of these subclades ranged from 10 to 13%. In a previous study, Haick et al. (Reference Haick, Tavares-Costa, Rebêlo, Neves, Félix, Müller, Trinidade, Costa-Campos, Maschio and Melo2025) reported a divergence between Kalicephalus species from Neotropical snakes (K. atroxi and K. c. costatus) and species from Oriental snakes (K. indicus and K. brachycephalus). However, it is important to note that the node separating these groups had low support (bootstrap < 70, posterior probability < 0.90), and that the sequences of the Oriental species are designated as ‘unverified assembly’. Consequently, these sequences were not included in the present study. The clustering of our sequences with K. viperae from Iran should be interpreted only as preliminary evidence of molecular affinity within the genus, rather than as sufficient evidence to support biogeographic hypotheses, such as a historically widespread ancestral distribution or the relative age of different lineages. As additional sequences of Kalicephalus species from a broad range of geographic regions become available, together with multilocus analyses incorporating mitochondrial markers (e.g., COX1/COI or complete mitochondrial genomes), the phylogenetic relationships within the genus may be better resolved. Such data will provide a more robust framework for evaluating interspecific relationships and reconstructing the evolutionary and biogeographic history of Kalicephalus.
In conclusion, we report the first record of the genus Kalicephalus in C. tzabcan by morphological and molecular identification. Additionally, we report for the first time the presence of larvae in the reproductive tract of Kalicephalus females naturally parasitising a rattlesnake. Further studies are required to identify this nematode taxon to species level and elucidate the mechanisms underlying the intrauterine larval development in Kalicephalus species.
Supplementary material
The supplementary material for this article can be found at http://doi.org/10.1017/S0022149X26102259.
Acknowledgements
The authors thank Wilson Moguel Chin for his assistance with DNA extraction and amplification, and Berenit Mendoza Garfias and Gisela Martínez Flores for SEM micrographs.
Financial support
This study received no specific grant from any funding agency, commercial or not-for-profit sectors.
Competing interests
The authors declare none.
Ethical standards
All applicable institutional, national, and international guidelines for the care and use of animals were followed. The rattlesnake was collected under a licence from the Mexican Ministry of Environment (license number SPARN/DGVS/01639/25).
