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All issues Volume 30 (2023) Parasite, 30 (2023) 24 Full HTML
Open Access
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Parasite
Volume 30, 2023
Article Number 24
Number of page(s) 9
DOI https://doi.org/10.1051/parasite/2023027
Published online 04 July 2023

© F. Binetruy & O. Duron, published by EDP Sciences, 2023

Licence Creative CommonsThis is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Introduction

Ticks are important vectors of viruses, bacteria, and protozoan parasites of medical and veterinary importance [31, 44]. Ticks can also harbor filarial nematodes of the family Onchocercidae Leiper, 1911, commonly known as filariae or filarioids, but their presence is not often the focus of research on tick-borne pathogens [10, 13, 45, 46, 56]. Members of the Onchocercidae family have evolved specialized parasitic life cycles in terrestrial vertebrates and in a variety of blood feeding dipterans (mosquitoes, biting midges, sand flies, horse flies, black flies, etc.) which are the main vectors for most species [29, 46, 64, 67]. In vertebrate hosts, mature filarioids live in the circulatory system or in particular tissues and release microfilariae into the bloodstream. The microfilariae are next taken up by blood-feeding arthropod vectors in which they develop into infective larvae that can be transmitted to a new vertebrate host through further biting. Some filarioids are causative agents of major human (lymphatic filariasis, onchocerciasis, loiasis) and animal diseases (canine heartworm disease) [29, 64, 67].

Early microscopic morphological observations provided evidence for the presence of filarioids in field populations of ticks: Filarioids have been morphologically identified in North America in Ixodes cookei [4] and I. dammini [12], in South America in Amblyomma cajennense [74] and Ornithodoros talaje [37], in Europe in I. ricinus [1, 60, 73] and Rhipicephalus sanguineus sensu lato (s.l.) [7], in North Africa in R. sanguineus s.l. [5], in Australia in I. trichosuri [66] and in Western Asia in O. tartakowskyi [11]. More recently, filarioids were also morphologically identified in Japan in Haemaphysalis flava and H. japonica [71]. Molecular investigations confirmed the presence of filarioids in I. ricinus [60] and R. sanguineus s.l. [13, 45, 56], and further led to their detection in other tick species in North America, I. scapularis [30, 51, 69, 70] and Amblyomma americanum [40, 76]. Field surveys demonstrated that infection rates of filarioids can reach 5% in some populations of I. ricinus [1, 60], and may be as high as 25% in I. scapularis [51] and 50% in some populations of R. sanguineus s.l. [13, 45].

Microscopic observations and molecular typing converge to identify most filarioids of ticks as members of the genera Acanthocheilonema Cobbold, 1870, Monanema Anteson, 1968, Yatesia Bain, Baker & Chabaud, 1982, and Cercopithifilaria Eberhard, 1980 [1, 5, 7, 11, 12, 40, 46, 51, 56, 59, 60, 69, 71]. While early microscopic morphological observations classified some filarioids of ticks as members of the genus Dipetalonema Diesing, 1861, numerous subgenera or species formerly included in this genus were further renamed and elevated to generic rank as Cercopithifilaria [8, 10, 28]. Phylogenetic analyses based on molecular and morphological data further showed that the genera Acanthocheilonema, Monanema, Yatesia and Cercopithifilaria (all associated with ticks), as well as the genera Dipetalonema (associated with biting midges), Litomosoides Chandler, 1931 (associated with parasitic mites), and Cruorifilaria Eberhard, Morales & Orihel, 1976 (not yet associated with a vector), cluster in a monophyletic clade of filarioids, termed as the Dipetalonema lineage or the ONC4 clade, within the family Onchocercidae [8, 10, 28, 46]. Some members of the Dipetalonema lineage could provide immunologically relevant and experimentally tractable laboratory models of human onchocerciasis since they cause skin and ocular immunopathologies in vertebrates analogous to those observed in humans infected with Onchocerca volvulus Leuckart, 1893 [66, 72].

Experimental infection assays have confirmed the vector competence of ticks for filarioids of the genera Acanthocheilonema, Monanema, Yatesia and Cercopithifilaria. Baltazard et al. [11] first experimentally showed that the soft tick O. tartakovskyi can be infected by Acanthocheilonema viteae (Krepkogorskaya, 1933; formerly Dipetalonema viteae) after feeding on infected field vertebrates and that this tick species can further transmitted viable infective larvae to domestic rodents through biting. This tick-borne infection cycle has been conducted for several decades in the laboratory using O. tartakovskyi as an exclusive vector [5, 6, 48], or O. moubata as an alternative vector [63]. Further experimental infection assays showed similar vector competence in pairing of hard ticks (belonging to the genera Ixodes, Rhipicephalus, Amblyomma, Haemaphysalis and Hyalomma) and some filarioids of the Dipetalonema lineage: ticks feeding on infected vertebrates ingest microfilariae, which can develop up to the viable infective third stage in a few weeks and are further excreted with saliva during biting [4, 6, 9, 14–16, 24, 42, 52, 53, 58, 66, 73, 74]. Filarioids of the Dipetalonema lineage also survive transstadially in such ticks since the development from microfilariae to infective third-stage larvae occurs only while the tick is off-host, that is, during ecdysis from tick larva to nymph or from nymph to adult [53, 66]. While all these observations converge to confirm that ticks are effective vectors for these filarioids, members of the Dipetalonema lineage are currently neglected tick-borne parasites [10, 46, 56].

In this study, we conducted a molecular survey of filarioids in ticks in French Guiana. This remote territory is a vast equatorial land located on the north–east coast of South America, mostly covered by dense rainforests and with a low density of human population [68]. We know little on the presence and diversity of tick-borne diseases in French Guiana, but recent surveys led to the description of novel tick-borne pathogens [17, 21, 32, 43]. Here, we surveyed 682 field ticks belonging to 22 tick species of the 33 known from French Guiana [19]. We further used cytochrome c oxidase subunit I (cox1) mitochondrial gene sequences, and phylogenetics for the description of filarioid infections in these ticks. Lastly, we examined and discussed their genetic proximity with already known species of filarioids associated with ticks.

Materials and methods

Tick collection

Field ticks (n = 682; belonging to 22 tick species) were collected in French Guiana either on vegetation by the drag-flag method (for questing ticks) or directly from their vertebrate hosts or seabird nests (for engorged or questing ticks) (Tables 1 and S1). Samples included five major tick genera belonging to the family Ixodidae (hard ticks): Amblyomma (16 species), Rhipicephalus (2 species), Ixodes (1 species), Dermacentor (1 species) and Haemaphysalis (1 species). Samples include one species of the Ornithodoros genus, belonging to the Argasidae family (soft ticks). All ticks were stored in 75% ethanol until examination. Ticks were further examined under a Leica Z16 APO A macroscope and categorized by species, stage and sex in previous studies [18–20]. Use of genetic resources was approved by the French Ministry of the Environment under reference #TREL19028117S/156, in compliance with the Access and Benefit Sharing procedure implemented by the Loi pour la Reconquête de la Biodiversité.

Table 1

List of tick species examined for infection by filarioids in French Guiana (cf. Table S1 for more details).

Molecular detection and typing of filarioids in ticks

DNA was extracted from individual ticks using a DNeasy Blood & Tissue Kit (QIAGEN, Hilden, Germany), following the manufacturer’s instructions. Each individual extract was then tested by a polymerase chain reaction (PCR) assay for filarioid infection by amplifying a fragment of the cox1 mitochondrial gene (689 bp) using the CO1intF (5′–TGA TTG GTG GTT TTG GTA A–3′) and CO1intR (5′–ATA AGT ACG AGT ATC AAT ATC–3′) primers [26]. PCRs were performed in a total volume of 25 μL containing 10–50 ng of genomic DNA, 8 mM of each dNTP (Thermo Fisher Scientific, Waltham, MA, USA), 10 mM of MgCl2 (Thermo Fisher Scientific), 7.5 μM of each of the internal primers, 2.5 μL of 10× PCR buffer (Thermo Fisher Scientific), and 1.25 U of Taq DNA polymerase (Thermo Fisher Scientific). All PCR amplifications were performed as follows: initial denaturation at 93 °C for 3 min, 35 cycles of denaturation (93 °C, 30 s), annealing (50 °C, 30 s), extension (72 °C, 1 min), and a final extension (72 °C, 5 min). Positive (DNA of the filarioid Loa loa) and negative (water) controls were included in each PCR assay. Following visualization via electrophoresis in 1.5% agarose gel, positive PCR products were sequenced by Eurofins. Sequence chromatograms were cleaned with Chromas Lite (http://www.technelysium.com.au/chromas_lite.html), and alignments were performed using ClustalW, implemented in the MEGA software package (https://www.megasoftware.net/). New sequences obtained in this study were deposited in GenBank under accession numbers OR030075–OR030095.

Molecular phylogenetic analyses

Phylogenetic analyses were based on sequence alignments of the filarioid cox1 gene sequences obtained in this study. Sequences of other filarioids obtained from GenBank, including representative members of the Dipetalonema lineage (Acanthocheilonema, Yatesia, Cercopithifilaria, Cruorifilaria, Litomosoides and Dipetalonema) and of other filarial nematodes were also included in the phylogenetic analyses. The Basic Local Alignment Search Tool (BLAST; https://blast.ncbi.nlm.nih.gov/blast/Blast.cgi) was used to find additional sequences available on GenBank and showing the highest nucleotide similarities with the filarioid gene sequences we characterized in this study. The Gblocks program with default parameters was used to obtain non-ambiguous sequence alignments [27]. Tree-based phylogenetic analyses were performed using maximum-likelihood (ML) analyses using the MEGA software package (https://www.megasoftware.net/). The evolutionary models that best fit the sequence data were determined using the Akaike information criterion. Clade robustness was assessed by bootstrap analysis using 1,000 replicates.

Statistical analyses

All statistical analyses were carried out using the R statistical package (https://www.r-project.org/).

Results

Detection of filarioids in ticks

Among the 682 tick specimens from French Guiana, cox1 PCR assay indicated the presence of filarioids in 21 ticks (3.1%) belonging to five species: A. cajennense (n = 15 infected specimens), A. oblongoguttatum (n = 2), A. romitii (n = 1), I. luciae (n = 2) and R. sanguineus s.l. (n = 1) (Tables 1 and S1). No filarioid was detected in the 17 other tick species. The detection rate of filarioids did not co-vary with the screening effort, i.e., the number of examined specimens per tick species (Spearman’s rank correlation, rs = 0.24, p = 0.28): The tick species observed with filarioids were not always among those for which we examined more specimens. This was shown by A. romitii (n = 2 examined specimens, 1 filarioid-positive specimen: 50%), I. luciae (n = 6 examined specimens, 2 filarioid-positive specimens: 33.3%) and R. sanguineus s.l. (n = 6 examined specimens, 1 filarioid-positive specimens: 16.7%), although we detected filarioid-positive specimens in tick species for which we examined a large number of specimens: A. cajennense (n = 229 examined specimens, 15 filarioid-positive specimens: 6.6%) and A. oblongoguttatum (n = 66 examined specimens, two filarioid-positive specimens: 3%). Filarioids were not detected in some tick species for which we examined a large number of specimens, such as A. longirostre (n = 137 examined specimens but none positive) and Dermacentor nitens (n = 97 examined specimens but none positive) (Tables 1 and S1). Of the 21 filarioid-infected tick specimens, 18 were unfed A. cajennense (n = 15), A. oblongoguttatum (n = 2) and R. sanguineus s.l. (n = 1) ticks while three were engorged A. romitii (n = 1) and I. luciae (n = 2) ticks collected on capybaras and opossums, respectively (Table S1). Of the 21 filarioid-infected tick specimens, 12 were A. cajennense nymphs and nine were adults (A. cajennense, n = 3 including one male and two females; A. oblongoguttatum, n = 2 females; A. romitii, n = 1 female; I. luciae, n = 2 including one male and one female; R. sanguineus s.l., n = 1 male) (Table S1).

Molecular typing of filarioids in ticks

On the basis of cox1 gene sequences, we characterized six genetically distinct filarioids (pairwise nucleotide identity: 84.1-99.8%) in A. cajennense, A. oblongoguttatum, A. romitii, I. luciae and R. sanguineus s.l. Two genetically distinct filarioids were detected in A. cajennense (pairwise nucleotide identity: 87.1%), two in I. luciae (pairwise nucleotide identity: 99.5%), and one in each of the other infected tick species (A. oblongoguttatum, A. romitii and R. sanguineus s.l.). Of the six genetically distinct filarioids, each was specific to its respective tick species, expect one which was shared by A. cajennense and I. luciae.

The filarioid cox1 gene sequence we detected in R. sanguineus s.l. was identical to those of Cercopithifilaria bainae Almeida & Vicente, 1984, previously characterized from dogs in Italy and the USA (GenBank accession numbers: KF270686, KP760175, MH390716; 100% pairwise nucleotide identities) [23, 46, 65]. However, none of the filarioid cox1 gene sequences we identified in A. cajennense, A. oblongoguttatum, A. romitii and I. luciae was 100% identical to other filarioid sequences available on GenBank, although showing high levels of nucleotide identity with some members of the Dipetalonema lineage. The filarioid cox1 gene sequence of A. oblongoguttatum showed highest pairwise nucleotide identities with members of the genus Cercopithifilaria, including Cercopithifilaria bainae infecting dogs (GenBank accession numbers: KF270686, KP760175, MH390716; 87.1% pairwise nucleotide identities) [23, 46, 65], Cercopithifilaria grassii (Noe, 1907) infecting dogs (JQ837810; 88.7%) [57], Cercopithifilaria sp. II sensu Otranto et al., 2013 [55] infecting dogs (KX898457; 87.9%) [49] and Cercopithifilaria multicauda Uni & Bain, 2001, infecting Japanese serow (AB178848; 87.6%) [2]. The filarioid cox1 gene sequence of A. romitii and one of A. cajennense showed highest pairwise nucleotide identities with Cruorifilaria tuberocauda Eberhard, Morales & Orihel,1976, infecting capybara (KP760176; 98.7-90.1%) [46]. The two filarial cox1 gene sequences of I. luciae and one of A. cajennense showed highest pairwise nucleotide identities with members of the genus Dipetalonema infecting monkeys, including Dipetalonema yatesi Notarnicola, Jiménez & Gardner, 2007 (MW199183; 88%) [75], Dipetalonema gracile (Rudolphi, 1809) (KP760181; 88.9%) [46] and Dipetalonema caudispina (Molin, 1858) (MT502200; 88.9%) [62].

Phylogeny of filarioids in ticks

A ML analysis based on cox1 nucleotide sequences was further conducted to examine the phylogenetic proximity between the filarioids of A. cajennense, A. oblongoguttatum, A. romitii, I. luciae and R. sanguineus s.l. and with other filarioids. Phylogenetic reconstruction revealed a clustering of the genera Cercopithifilaria, Cruorifilaria, Litomosoides, Yatesia, Acanthocheilonema and Dipetalonema in a unique monophyletic clade, the Dipetalonema lineage, distinct to other members of the family Onchocercidae (Fig. 1). Phylogenetic reconstruction further showed within the Dipetalonema lineage that:

  1. The filarioids of A. oblongoguttatum and R. sanguineus s.l. formed a robust subclade, supported by a 91% bootstrap value, together with all species of the genus Cercopithifilaria (Fig. 1). This suggested that the filarioids of A. oblongoguttatum and R. sanguineus s.l. belong to the genus Cercopithifilaria. While the filarioid of R. sanguineus s.l. was probably Cercopithifilaria bainae (as suggested by their 100% pairwise nucleotide identity), the filarioid of A. oblongoguttatum was distinct from already known Cercopithifilaria species.

  2. The filarioid of A. romitii and one of A. cajennense delineated a robust subclade (95% bootstrap value) together with Cruorifilaria tuberocauda (Fig. 1), suggesting that all belong to the genus Cruorifilaria. The filarioid of A. romitii and one of A. cajennense were distinct to already known Cruorifilaria species.

  3. The other filarioid of A. cajennense and those of I. luciae clustered in a unique robust subclade (provisionally named Dipetalonema-like hereafter; 99% bootstrap value) substantially divergent from all other genera of the Dipetalonema lineage (Fig. 1). The phylogenetic proximity of this filarioid of A. cajennense to those of I. luciae, and their 99.8–100% pairwise nucleotide identities, suggested that they probably belong to the same species. The closest relative of Dipetalonema-like is the genus Dipetalonema, while other genera are more distantly related. This suggests that these Dipetalonema-like filarioids may belong to a potential novel genus which could be a sister genus of Dipetalonema.

thumbnail Figure 1

Phylogeny of Onchocercid filarioids constructed using maximum-likelihood (ML) estimations based on cox1 nucleotide sequences with a total of 649 unambiguously aligned bp (best-fit approximation for the evolutionary model: GTR+G+I). Major genera of the Dipetalonema lineage (Acanthocheilonema, Yatesia, Cercopithifilaria, Cruorifilaria, Litomosoides and Dipetalonema), including representative species with indication of vector range (ticks, biting midges, parasitic mites), are indicated. Filarioid sequences obtained in this study from the ticks A. cajennense (n = 15), A. oblongoguttatum (n = 2), A. romitii (n = 2), I. luciae (n = 2) and R. sanguineus s.l. (n = 1) are in bold. GenBank accession numbers of sequences used in analyses are shown on the phylogenetic trees. Numbers at nodes indicate percentage support of 1,000 bootstrap replicates. Only bootstrap values >70% are shown. The scale bar is in units of substitution/site.

Discussion

We show here that substantial diversity of filarioids of the Dipetalonema lineage is present in ticks of French Guiana. We conducted extensive molecular screening of 22 tick species, and identified filarioids in five tick species, A. cajennense, A. oblongoguttatum, A. romitii, I. luciae and R. sanguineus s.l. No filarioid was detected in the 17 other tick species. While pairwise nucleotide identities and phylogenetics assigned the filarioid of R. sanguineus s.l. to an already described species, Cercopithifilaria bainae, all other filarioids detected in this study are distinct to already known species of Cercopithifilaria, Cruorifilaria and Dipetalonema on the basis of their cox1 gene sequences. To our knowledge, this is also the first observation of Cruorifilaria in ticks (and, more broadly, in blood-feeding arthropods), suggesting that A. cajennense and A. romitii are vectors of this filarioid genus.

The presence of filarioid DNA in A. cajennense, A. oblongoguttatum, A. romitii, I. luciae and R. sanguineus s.l. suggests that these tick species may serve as vectors for filarial nematodes. Obviously, these observations are not conclusive evidence of vector competence: ticks feeding on parasitemic vertebrates can be positive for filarioid DNA due to the presence of infected blood in the tick digestive tract, but may not be competent to transmit infectious filarioids during their next blood meal. However, experimental infection assays have directly confirmed the vector competence of Ixodes, Amblyomma and Rhipicephalus ticks for filarioids of the Dipetalonema lineage, including the brown dog tick R. sanguineus s.l. for Cercopithifilaria bainae [4, 6, 24, 42, 52, 53, 58, 66, 73, 74]. In the field, the detection of viable microfilariae and infective filarial larvae in the hindgut, haemocoele, fat cells or ducts of salivary glands of ticks also support these observations [5, 7, 12]. In addition, the accumulation of microfilariae in the dermis of infected vertebrates at the sites of attachment of ticks suggest a specific chemotactic response of filarioids to tick saliva [42, 50]. Furthermore, our detection of filarioids from questing (unfed) A. cajennense, A. oblongoguttatum, and R. sanguineus s.l. ticks that have already digested their previous blood meals, have further moulted, and are seeking vertebrates for their next blood meal suggests that these tick species can acquire and maintain filarioids of the Dipetalonema lineage, a pattern of transstadial transmission also observed for other tick species [53, 66]. Finally, the observation that all tick-associated filarioids cluster in a monophyletic clade, the Dipetalonema lineage, distinct from clades of filarioids specifically associated with transmission by blood-feeding dipterans, supports the conclusion that ticks are vectors of these filarioids.

Tick-borne filarioids have been detected from most continents, suggesting worldwide distribution of the Dipetalonema lineage [10, 46]. Some species may have global distribution such as Cercopithifilaria bainae, specifically associated with dogs and R. sanguineus s.l., which was first formally identified in Brazil [3], and further in Europe [41, 56, 65], Asia [13, 45], Iran [61], North America [23, 47], and now in French Guiana (this study). The other filarioids we characterized in French Guiana have not been detected anywhere and their transmission cycles remain unknown. In fact, the vertebrate hosts of the Cercopithifilaria sp. of A. oblongoguttatum remain unidentified since this tick is a generalist species feeding on a wide range of mammals [19].

For the Cruorifilaria spp. of A. romitii and A. cajennense, two biological features suggest that capybara could be their vertebrate host. First, the single described Cruorifilaria species, Cruorifilaria tuberocauda, has been reported as a parasite of capybara in other South American regions [25, 33], and, second, A. romitii is a tick species highly specific to capybara, while A. cajennense is a generalist tick species often found on these animals [19]. For the Dipetalonema-like filarioids, opossums could be vertebrate hosts since one of the infected tick species, I. luciae, is a specialized tick species that feeds primarily on opossums [19]. Past studies have morphologically identified four filarial species, all of the Dipetalonema lineage, in South American opossums: Acanthocheilonema pricei (Vaz et Pereira, 1934), Cercopithifilaria didelphis (Esslinger et Smith, 1979), Skrjabinofilaria skrjabini Travassos, 1925, and Cherylia guyanensis Bain et al., 1985 [9, 35]. No molecular data are currently available for these four species, which prevents any definitive conclusions.

The detection of members of the Dipetalonema lineage in ticks of significant medical or veterinary interest is of concern, but the risk of acquiring a tick-borne filarial infection is unknown. While A. cajennense and A. oblongoguttatum are the tick species most commonly biting humans in South America [36, 39], in the present study, we found that 3–6.6% of field specimens were infected with filarioids of the Dipetalonema lineage. Humans are therefore exposed to bites of filarioid-infected ticks without the risk of infection being documented to date. In North America, a survey of humans with a tick-bite history for the presence of antibodies against the I. scapularis-associated filarioid found, however, no evidence of infection [69]. In animals, filarioids of the Dipetalonema lineage are often considered innocuous, but some studies and evidence suggest that they are not. In dogs, recent surveys showed that Cercopithifilaria bainae can induce erythematous and pruritic dermatitis, subcutaneous nodules, ulcerative skin lesions and chronic polyarthritis [23, 38, 54]. Infections of dogs by other members of the Dipetalonema lineage, Acanthocheilonema spp., were also associated with clinical symptoms, such as anemia and high levels of plasma proteins [22, 34]. In capybaras, adults of Cruorifilaria tuberocauda form tangled masses in the kidneys, and less frequently in the pulmonary arteries, where they cause extensive tissue damage with severe endarteritis, leading to obstruction of vessels and infarction of adjacent tissues [33].

In conclusion, we have identified a wide range of Cercopithifilaria, Cruorifilaria and Dipetalonema-like filarial nematodes in French Guiana. The repeated detection of the Dipetalonema lineage in major tick genera on most continents confirms that these are widespread but neglected tick-borne filarioids of largely undocumented veterinary (and perhaps medical) concern. Their pathogenicity, epidemiology, developmental cycles and transmission mechanisms by ticks must be further investigated.

Acknowledgments

We are grateful to Roxane Barosi and Sébastien Pion for technical support. We wish to thank all members of LabEx CEBA who provided tick specimens, in particular Benoit de Thoisy, François Catzeflis, Frederic Delsuc, Denis Blanchet and Philippe Gaucher. We are grateful to Rolland Ruffine, to the Groupe d’Étude et de Protection des Oiseaux en Guyane (GEPOG) and to members of Institut Pasteur de Guyane, in particular to Isabelle Dufour, Stanislas Talaga and Agathe Chavy, for support in the field. We also thank Xavier Baudrimont from the Direction de l’Alimentation, de l’Agriculture et de la Forêt de Guyane (DAAF) and to breeders for having allowed us to collect ticks on cattle and horses.

Supplementary Materials

Table S1: Details of tick species and specimens examined for infection by filarioids in French Guiana. Access here

References

  1. Aeschlimann A, Burgdorfer W, Matile H, Peter O, Wyler R. 1979. Aspects nouveaux du rôle de vecteur joué par Ixodes ricinus L. en Suisse. Note préliminaire. Acta Tropica, 36, 181–191. [PubMed] [Google Scholar]
  2. Agatsuma T, Iwagami M, Uni S, Takaoka H, Katsumi A, Kimura E, Bain O. 2005. Molecular phylogenetic relationships among seven Japanese species of Cercopithifilaria. Parasitology International, 54, 195–199. [CrossRef] [PubMed] [Google Scholar]
  3. Almeida GLG, Vicente JJ. 1984. Cercopithifilaria bainae sp. n. parasita de Canis familiaris (L.) (Nematoda, Filarioidea). Atas Soc Biol Rio de Janeiro, 24, 18. [Google Scholar]
  4. Anteson RK. 1968. Biological studies of Monanema marmotae (Webster 1967) a filarioid parasite of the woodchuck Marmota monax canadensis. University of Connecticut: Storrs, Connecticut. [Google Scholar]
  5. Bain O. 1972. Recherches sur la morphogénèse des Filaires chez l’hôte intermédiaire. Annales de Parasitologie Humaine et Comparée, 47, 251–303. [CrossRef] [EDP Sciences] [Google Scholar]
  6. Bain O. 1967. Biologie larvaire et mécanisme de transmission de la Filaire Dipetalonema viteae. Annales de Parasitologie Humaine et Comparée, 42, 211–267. [CrossRef] [EDP Sciences] [Google Scholar]
  7. Bain O, Aeschlimann A, Chatelanat P. 1982. Présence, chez des tiques de la région de Genève, de larves infestantes qui pourraient se rapporter à la filaire de chien Dipetalonema grassii. Annales de Parasitologie Humaine et Comparée, 57, 643–646. [CrossRef] [EDP Sciences] [Google Scholar]
  8. Bain O, Baker M, Chabaud AG. 1982. Nouvelles données sur la lignée Dipetalonema (Filarioidea, Nematoda). Annales de Parasitologie Humaine et Comparée, 57, 593–620. [CrossRef] [EDP Sciences] [Google Scholar]
  9. Bain O, Petit G, Jacquet-viallet P, Houin R. 1985. Cherylla guyanensis n. gen., n. sp., – Filaire d’un Marsupial sud-américain, transmise par tique. Annales de Parasitologie Humaine et Comparée, 60, 727–737. [CrossRef] [EDP Sciences] [Google Scholar]
  10. Bain O, Casiraghi M, Martin C, Uni S. 2008. The nematoda filarioidea: Critical analysis linking molecular and traditional approaches. Parasite, 15, 342–348. [CrossRef] [EDP Sciences] [PubMed] [Google Scholar]
  11. Baltazard M, Chabaud AG, Minou A. 1952. Cycle evolutif d’une filaire parasite de merion. Comptes Rendus Hebdomadaires des Séances de l’Academie des Sciences, 234, 2115–2117. [Google Scholar]
  12. Beaver PC, Burgdorfer W. 1984. A microfilaria of exceptional size from the ixodid tick, Ixodes dammini, from Shelter Island, New York. The Journal of Parasitology, 70, 963–966. [CrossRef] [PubMed] [Google Scholar]
  13. Bezerra-Santos MA, de Macedo LO, Nguyen VL, Manoj RR, Laidoudi Y, Latrofa MS, Beugnet F, Otranto D. 2022. Cercopithifilaria spp. in ticks of companion animals from Asia: new putative hosts and vectors. Ticks and Tick-Borne Diseases, 13, 101957. [CrossRef] [PubMed] [Google Scholar]
  14. Bianco AE. 1984. Laboratory maintenance of Monanema globulosa, a rodent filaria with skin-dwelling microfilariae. Zeitschrift Für Parasitenkunde, 70, 255–264. [CrossRef] [Google Scholar]
  15. Bianco AE, Muller RL. 1977. A hard tick as vector of a new rodent filaria. Transactions of the Royal Society of Tropical Medicine and Hygiene, 71, 383. [Google Scholar]
  16. Bianco AE, Nelson GS, Muller R. 1983. Biology of Monanema globulosa, a rodent filaria with skin-dwelling microfilariae. Journal of Helminthology, 57, 259–278. [CrossRef] [Google Scholar]
  17. Binetruy F, Buysse M, Barosi R, Duron O. 2020. Novel Rickettsia genotypes in ticks in French Guiana, South America. Scientific Reports, 10, 2537. [CrossRef] [PubMed] [Google Scholar]
  18. Binetruy F, Buysse M, Lejarre Q, Barosi R, Villa M, Rahola N, Paupy C, Ayala D, Duron O. 2020. Microbial community structure reveals instability of nutritional symbiosis during the evolutionary radiation of Amblyomma ticks. Molecular Ecology, 29, 1016–1029. [CrossRef] [PubMed] [Google Scholar]
  19. Binetruy F, Chevillon C, de Thoisy B, Garnier S, Duron O. 2019. Survey of ticks in French Guiana. Ticks and Tick-Borne Diseases, 10, 77–85. [CrossRef] [PubMed] [Google Scholar]
  20. Binetruy F, Dupraz M, Buysse M, Duron O. 2019. Surface sterilization methods impact measures of internal microbial diversity in ticks. Parasites and Vectors, 12, 268. [CrossRef] [Google Scholar]
  21. Binetruy F, Garnier S, Boulanger N, Talagrand-Reboul É, Loire E, Faivre B, Noël V, Buysse M, Duron O. 2020. A novel Borrelia species, intermediate between Lyme disease and relapsing fever groups, in neotropical passerine-associated ticks. Scientific Reports, 10, 10596. [CrossRef] [PubMed] [Google Scholar]
  22. Bolio ME, Montes AM, Gutierrez C, Alonso FD, Bernal LJ, Sauri C, RodríguezVivas RI. 2002. Hallazgos clínicos en perros parasitados por Dipetalonema dracunculoides. Archivos de Medicina Veterinaria: XXXIV. p. 283–286. [Google Scholar]
  23. Boyd M, Santoro D, Craft WF, Ginn PE, Childress AL, Wellehan JFX, Walden HS. 2019. Dermatitis caused by autochthonous Cercopithifilaria bainae from a dog in Florida, USA: clinical, histological and parasitological diagnosis and treatment. Veterinary Dermatology, 30, 68–e20. [CrossRef] [PubMed] [Google Scholar]
  24. Brianti E, Otranto D, Dantas-Torres F, Weigl S, Latrofa MS, Gaglio G, Napoli E, Brucato G, Cauquil L, Giannetto S, Bain O. 2012. Rhipicephalus sanguineus (Ixodida, Ixodidae) as intermediate host of a canine neglected filarial species with dermal microfilariae. Veterinary Parasitology, 183, 330–337. [CrossRef] [PubMed] [Google Scholar]
  25. Campo-Aesen I, Planas-Girón G. 1986. La Cruorifilaria tuberocauda (Nematoda: Filarioidea) en capibaras de Venezuela. Medicina Cutánea Ibero-Latino-Americana, 14, 109–113. [Google Scholar]
  26. Casiraghi M, Anderson TJC, Bandi C, Bazzocchi C, Genchi C. 2001. A phylogenetic analysis of filarial nematodes: comparison with the phylogeny of Wolbachia endosymbionts. Parasitology, 122(Pt 1), 93–103. [CrossRef] [PubMed] [Google Scholar]
  27. Castresana J. 2000. Selection of conserved blocks from multiple alignments for their use in phylogenetic analysis. Molecular Biology and Evolution, 17, 540–552. [PubMed] [Google Scholar]
  28. Chabaud AG, Bain O. 1976. La lignée Dipetalonema. Nouvel essai de classification. Annales de Parasitologie Humaine et Comparée, 51, 365–397. [CrossRef] [EDP Sciences] [Google Scholar]
  29. Cháves-González LE, Morales-Calvo F, Mora J, Solano-Barquero A, Verocai GG, Rojas A. 2022. What lies behind the curtain: Cryptic diversity in helminth parasites of human and veterinary importance. Current Research in Parasitology & Vector-Borne Diseases, 2, 100094. [CrossRef] [PubMed] [Google Scholar]
  30. Cross ST, Kapuscinski ML, Perino J, Maertens BL, Weger-Lucarelli J, Ebel GD, Stenglein MD. 2018. Co-Infection Patterns in Individual Ixodes scapularis Ticks Reveal Associations between Viral, Eukaryotic and Bacterial Microorganisms. Viruses, 10, 388. [CrossRef] [PubMed] [Google Scholar]
  31. Dantas-Torres F, Chomel BB, Otranto D. 2012. Ticks and tick-borne diseases: A One Health perspective. Trends in Parasitology, 28, 437–446. [CrossRef] [PubMed] [Google Scholar]
  32. Duron O, Koual R, Musset L, Buysse M, Lambert Y, Jaulhac B, Blanchet D, Alsibai KD, Lazrek Y, Epelboin L, Deshuillers P, Michaud C, Douine M. 2022. Novel chronic anaplasmosis in splenectomized patient, Amazon rainforest. Emerging Infectious Diseases, 28, 1673–1676. [CrossRef] [PubMed] [Google Scholar]
  33. Eberhard ML, Morales GA, Orihel TC. 1976. Cruorifilaria tuberocauda gen. et sp. n. (Nematoda: Filarioidea) from the Capybara, Hydrochoerus hydrochaeris in Colombia. Journal of Parasitology, 62, 604–607. [CrossRef] [Google Scholar]
  34. Espinosa N, Rosero A, Villegas CL, Garcia IC, Gaviria-Cantin T, Nieto AP, Ferro BE, Nieto Ramirez LM. 2022. First Report of Acanthocheilonema reconditum Outbreak in Canines with Clinical Signs of Anemia from Southwestern Colombia. Pathogens, 11, 1434. [CrossRef] [PubMed] [Google Scholar]
  35. Esslinger JH, Smith JL. 1979. Dipetalonema (Acanthocheilonema) didelphis sp. n. (Nematoda: Filarioidea) from opossums, with a redescription of D. (A.) pricei (Vaz and Pereira 1934). The Journal of Parasitology, 65, 928–933. [CrossRef] [PubMed] [Google Scholar]
  36. Estrada-Peña A, Jongejan F. 1999. Ticks feeding on humans: A review of records on human-biting Ixodoidea with special reference to pathogen transmission. Experimental and Applied Acarology, 23, 685–715. [CrossRef] [Google Scholar]
  37. Frank W. 1962. Biologie von Macdonaldius oschei (Filarioidea, Onchocercinae), zugleich ein Beitrag über die Wirtsspezifität von Ornithodoros talaje (Ixodoidea) Argasidae). Zeitschrift Fur Parasitenkunde, 22, 107–108. [CrossRef] [Google Scholar]
  38. Gabrielli S, Giannelli A, Brianti E, Dantas-Torres F, Bufalini M, Fraulo M, La Torre F, Ramos RAN, Cantacessi C, Latrofa MS, Cancrini G, Otranto D. 2014. Chronic polyarthritis associated to Cercopithifilaria bainae infection in a dog. Veterinary Parasitology, 205, 401–404. [CrossRef] [PubMed] [Google Scholar]
  39. Guglielmone AA, Beati L, Barros-Battesti DM, Labruna MB, Nava S, Venzal JM, Mangold AJ, Szabó MPJ, Martins JR, González-Acuña D, Estrada-Peña A. 2006. Ticks (Ixodidae) on humans in South America. Experimental and Applied Acarology, 40, 83–100. [CrossRef] [PubMed] [Google Scholar]
  40. Henning TC, Orr JM, Smith JD, Arias JR, Rasgon JL, Norris DE. 2016. Discovery of filarial nematode DNA in Amblyomma americanum in Northern Virginia. Ticks and Tick-Borne Diseases, 7, 315. [Google Scholar]
  41. Ionică AM, D’Amico G, Mitková B, Kalmár Z, Annoscia G, Otranto D, Modrý D, Mihalca AD. 2014. First report of Cercopithifilaria spp. in dogs from Eastern Europe with an overview of their geographic distribution in Europe. Parasitology Research, 113, 2761–2764. [CrossRef] [PubMed] [Google Scholar]
  42. Ko RC. 1972. The transmission of Ackertia marmotae Webster, 1967 (Nematoda: Onchocercidae) of groundhogs (Marmota monax) by Ixodes cookei. Canadian Journal of Zoology, 50, 437–450. [CrossRef] [PubMed] [Google Scholar]
  43. Koual R, Buysse M, Grillet J, Binetruy F, Ouass S, Sprong H, Duhayon M, Boulanger N, Jourdain F, Alafaci A, Verdon J, Verheyden H, Rispe C, Plantard O, Duron O. 2023. Phylogenetic evidence for a clade of tick-associated trypanosomes. Parasites and Vectors, 16, 1–13. [CrossRef] [Google Scholar]
  44. De La Fuente J, Estrada-Pena A, Venzal JM, Kocan KM, Sonenshine DE. 2008. Overview: Ticks as vectors of pathogens that cause disease in humans and animals. Frontiers in Bioscience: A Journal and Virtual Library, 13, 6938–6946. [CrossRef] [Google Scholar]
  45. Latrofa MS, Dantas-Torres F, Giannelli A, Otranto D. 2014. Molecular detection of tick-borne pathogens in Rhipicephalus sanguineus group ticks. Ticks and Tick-Borne Diseases, 5, 943–946. [Google Scholar]
  46. Lefoulon E, Bain O, Bourret J, Junker K, Guerrero R, Cañizales I, Kuzmin Y, Satoto TBT, Cardenas-Callirgos JM, de Souza Lima S, Raccurt C, Mutafchiev Y, Gavotte L, Martin C. 2015. Shaking the Tree: multi-locus sequence typing usurps current Onchocercid (Filarial Nematode) Phylogeny. PLoS Neglected Tropical Diseases, 9, e0004233. [CrossRef] [PubMed] [Google Scholar]
  47. Lineberry MW, Sundstrom KD, Little SE, Stayton EM, Allen KE. 2020. Detection of Cercopithifilaria bainae infection in shelter dogs and ticks in Oklahoma, USA. Parasites and Vectors, 13, 216. [CrossRef] [Google Scholar]
  48. Londono I. 1976. Transmission of microfilariae and infective larvae of Dipetalonema viteae (Filarioidea) among vector ticks, Ornithodoros tartakowskyi (Argasidae), and loss of microfilariae in coxal fluid. Journal of Parasitology, 62, 786–788. [CrossRef] [Google Scholar]
  49. Maia C, Casero M, Annoscia G, Latrofa MS, Colella V, Pereira A, Azevedo F, Otranto D. 2017. Cercopithifilaria sp. II in Vulpes vulpes: new host affiliation for an enigmatic canine filarioid. Parasitology Research, 116, 441–443. [CrossRef] [PubMed] [Google Scholar]
  50. Moorhouse DE. 1969. The accumulation of microfilariae beneath the attachment of Ixodes tasmani. Transactions of the Royal Society of Tropical Medicine and Hygiene, 63, 22. [PubMed] [Google Scholar]
  51. Namrata P, Miller JM, Shilpa M, Reddy PR, Bandoski C, Rossi MJ, Sapi E. 2014. Filarial nematode infection in Ixodes scapularis ticks collected from Southern Connecticut. Veterinary Sciences, 1, 5–15. [CrossRef] [Google Scholar]
  52. Olmeda-Garcia AS, Rodriguez-Rodriguez JA. 1994. Stage-specific development of a filarial nematode (Dipetalonema dracunculoides) in vector ticks. Journal of Helminthology, 68, 231–235. [CrossRef] [PubMed] [Google Scholar]
  53. Olmeda-García AS, Rodríguez-Rodríguez JA, Rojo-Vázquez FA. 1993. Experimental transmission of Dipetalonema dracunculoides (Cobbold 1870) by Rhipicephalus sanguineus (Latreille 1806). Veterinary Parasitology, 47, 339–342. [CrossRef] [PubMed] [Google Scholar]
  54. Otranto D. 2015. Diagnostic challenges and the unwritten stories of dog and cat parasites. Veterinary Parasitology, 212, 54–61. [CrossRef] [PubMed] [Google Scholar]
  55. Otranto D, Brianti E, Dantas-Torres F, Miró G, Latrofa MS, Mutafchiev Y, Bain O. 2013. Species diversity of dermal microfilariae of the genus Cercopithifilaria infesting dogs in the Mediterranean region. Parasitology, 140, 99–108. [CrossRef] [PubMed] [Google Scholar]
  56. Otranto D, Brianti E, Latrofa M, Annoscia G, Weigl S, Lia R, Gaglio G, Napoli E, Giannetto S, Papadopoulos E, Mir G, Dantas-Torres F, Bain O. 2012. On a Cercopithifilaria sp. transmitted by Rhipicephalus sanguineus: A neglected, but widespread filarioid of dogs. Parasites and Vectors, 5, 1. [CrossRef] [Google Scholar]
  57. Otranto D, Varcasia A, Solinas C, Scala A, Brianti E, Dantas-Torres F, Annoscia G, Martin C, Mutafchiev Y, Bain O. 2013. Redescription of Cercopithifilaria bainae Almeida & Vicente, 1984 (Spirurida, Onchocercidae) from a dog in Sardinia, Italy. Parasites and Vectors, 6, 132. [CrossRef] [Google Scholar]
  58. Petit G, Bain O, Carrat C, De Marval F. 1988. Development of the filaria Monanema martini in the epidermis of ixodid ticks. Annales de Parasitologie Humaine et Comparée, 63, 54–63. [CrossRef] [EDP Sciences] [Google Scholar]
  59. Petit G, Bain O, Cassone J, Seureau C. 1988. The filaria Cercopithifilaria roussilhoni in the tick vector. Annales de Parasitologie Humaine et Comparée, 63, 296–302. [CrossRef] [EDP Sciences] [Google Scholar]
  60. Ramos RAN, Giannelli A, Dantas-Torres F, Mallia E, Passantino G, Lia RP, Latrofa MS, Mutafchiev Y, Otranto D. 2013. Cercopithifilaria rugosicauda (Spirurida, Onchocercidae) in a roe deer and ticks from southern Italy. International Journal for Parasitology: Parasites and Wildlife, 2, 292–296. [CrossRef] [Google Scholar]
  61. Sazmand A, Bahiraei Z, Nemati F, Annoscia G, Bezerra-Santos MA, Nayebzadeh H, Salemi AM, Mousavi SM, Sadjjadi SM, Otranto D. 2022. Dermal microfilariae of dogs, jackals and cats in different regions of Iran. Parasites and Vectors, 15, 28. [CrossRef] [Google Scholar]
  62. Shaffer CA, Milstein MS, Lindsey LL, Wolf TM, Suse P, Marawanaru E, Kipp EJ, Garwood T, Travis DA, Terio KA, Larsen PA. 2022. “Spider Monkey Cotton”: Bridging Waiwai and scientific ontologies to characterize spider monkey (Ateles paniscus) filariasis in the Konashen community owned conservation area, Guyana. International Journal of Primatology, 43, 253–272. [CrossRef] [Google Scholar]
  63. Singh DP, Chatterjee RK, Sen AB. 1988. Studies on susceptibility of Ornithodoros moubata as vector of Dipetalonema viteae (Filariodea). Journal of Communicable Diseases, 20, 220–225. [Google Scholar]
  64. Small ST, Tisch DJ, Zimmerman PA. 2014. Molecular epidemiology, phylogeny and evolution of the filarial nematode Wuchereria bancrofti. Infection, Genetics and Evolution, 28, 33–43. [CrossRef] [PubMed] [Google Scholar]
  65. Solinas C, Varcasia A, Brianti E, Giannetto S, Pipia AP, Columbano N, Tosciri G, Dantas-Torres F, Garippa G, Otranto D, Scala A. 2014. Cercopithifilaria spp. in dogs in Sardinia Island (Italy). Parasitology Research, 113, 675–679. [CrossRef] [PubMed] [Google Scholar]
  66. Spratt DM, Haycock P. 1988. Aspects of the life history of Cercopithifliraia johnstoni (Nematoda:Filarioidea). International Journal for Parasitology, 18, 1087–1092. [CrossRef] [PubMed] [Google Scholar]
  67. Taylor MJ, Hoerauf A, Bockarie M. 2010. Lymphatic filariasis and onchocerciasis. Lancet, 376, 1175–1185. [CrossRef] [PubMed] [Google Scholar]
  68. de Thoisy B, Duron O, Epelboin L, Musset L, Quénel P, Roche B, Binetruy F, Briolant S, Carvalho L, Chavy A, Couppié P, Demar M, Douine M, Dusfour I, Epelboin Y, Flamand C, Franc A, Ginouvès M, Gourbière S, Houël E, Kocher A, Lavergne A, Le Turnier P, Mathieu L, Murienne J, Nacher M, Pelleau S, Prévot G, Rousset D, Roux E, Schaub R, Talaga S, Thill P, Tirera S, Guégan JF. 2021. Ecology, evolution, and epidemiology of zoonotic and vector-borne infectious diseases in French Guiana: Transdisciplinarity does matter to tackle new emerging threats. Infection, Genetics and Evolution, 93, 104916. [CrossRef] [PubMed] [Google Scholar]
  69. Tokarz R, Tagliafierro T, Ian Lipkin W, Marques AR. 2020. Characterization of a Monanema nematode in Ixodes scapularis. Parasites and Vectors, 13, 371. [CrossRef] [Google Scholar]
  70. Tokarz R, Tagliafierro T, Sameroff S, Cucura DM, Oleynik A, Che X, Jain K, Lipkin WI. 2019. Microbiome analysis of Ixodes scapularis ticks from New York and Connecticut. Ticks and Tick-Borne Diseases, 10, 894–900. [CrossRef] [PubMed] [Google Scholar]
  71. Uni S, Bain O, Fujita H, Matsubayashi M, Fukuda M, Takaoka H. 2013. Infective larvae of Cercopithifilaria spp. (Nematoda: Onchocercidae) from hard ticks (Ixodidae) recovered from the Japanese serow (Bovidae). Parasite, 20, 1. [CrossRef] [EDP Sciences] [PubMed] [Google Scholar]
  72. Vuong PN, Spratt D, Wanji S, Aimard L, Bain O. 1993. Onchocerca-like lesions induced by the filarioid nematode Cercopithifilaria johnstoni, in its natural hosts and in the laboratory rat. Annales de Parasitologie Humaine et Comparee, 68, 176–181. [CrossRef] [EDP Sciences] [Google Scholar]
  73. Winkhardt HJ. 1980. Untersuchungen über den Entwicklungszyklus von Dipetalonema rugosicauda (syn. Wehrdikmansia rugosicauda) (Nematoda: Filarioidea) II. Die Entwicklung von Dipetalonema rugosicauda im Zwischenwirt Ixodes ricinus und Untersuchungen über das Vorkommen der Mikrofilarien im Reh (Capreolus capreolus). Tropenmedizin und Parasitologie, 31, 21–30. [PubMed] [Google Scholar]
  74. Yates JA, Lowrie RC. 1984. Development of Yatesia hydrochoerus (Nematoda: Filarioidea) to the infective stage in Ixodid ticks. Proceedings of the Helminthological Society of Washington, 51, 187–190. [Google Scholar]
  75. Zárate-Rendón DA, Salazar-Espinoza MN, Catalano S, Sobotyk C, Mendoza AP, Rosenbaum M, Verocai G. 2022. Molecular characterization of Dipetalonema yatesi from the black-faced spider monkey (Ateles chamek) with phylogenetic inference of relationships among Dipetalonema of Neotropical primates. International Journal for Parasitology: Parasites and Wildlife, 17, 152. [CrossRef] [Google Scholar]
  76. Zhang X, Norris DE, Rasgon JL. 2011. Distribution and molecular characterization of Wolbachia endosymbionts and filarial nematodes in Maryland populations of the lone star tick (Amblyomma americanum). FEMS Microbiology Ecology, 77, 50–56. [CrossRef] [PubMed] [Google Scholar]

Cite this article as: Binetruy F & Duron O. 2023. Molecular detection of Cercopithifilaria, Cruorifilaria and Dipetalonema-like filarial nematodes in ticks of French Guiana. Parasite 30, 24.

All Tables

Table 1

List of tick species examined for infection by filarioids in French Guiana (cf. Table S1 for more details).

In the text

All Figures

thumbnail Figure 1

Phylogeny of Onchocercid filarioids constructed using maximum-likelihood (ML) estimations based on cox1 nucleotide sequences with a total of 649 unambiguously aligned bp (best-fit approximation for the evolutionary model: GTR+G+I). Major genera of the Dipetalonema lineage (Acanthocheilonema, Yatesia, Cercopithifilaria, Cruorifilaria, Litomosoides and Dipetalonema), including representative species with indication of vector range (ticks, biting midges, parasitic mites), are indicated. Filarioid sequences obtained in this study from the ticks A. cajennense (n = 15), A. oblongoguttatum (n = 2), A. romitii (n = 2), I. luciae (n = 2) and R. sanguineus s.l. (n = 1) are in bold. GenBank accession numbers of sequences used in analyses are shown on the phylogenetic trees. Numbers at nodes indicate percentage support of 1,000 bootstrap replicates. Only bootstrap values >70% are shown. The scale bar is in units of substitution/site.
In the text

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