A review of filarial nematodes parasitizing tick vectors: unraveling global patterns in species diversity, host associations, and interactions with tick-borne pathogens

Abstract

Ticks are well-known vectors of pathogens, including filarial nematodes, which can affect animal and human health. This review synthesizes current knowledge of filarial nematodes that utilize ticks as vectors, focusing on the Onchocercidae. Five genera of onchocercid filarial nematodes (Cercopithifilaria, Cherylia, Cruorifilaria, Monanema, and Yatesia) have demonstrated vector–parasite relationships with ticks, while Acanthocheilonema has only demonstrated molecular detection without confirmed vector competence. Vector competence has been experimentally proven for several species through documented development to infective stages. However, our understanding of tick-borne filarial nematodes remains limited owing to insufficient detection efforts and methodological challenges. Detection methods include traditional dissection, advanced imaging techniques such as microcomputed tomography (micro-CT), and molecular approaches, each with distinct advantages and limitations. To address knowledge gaps, future research should focus on conducting large-scale surveys of tick populations and wildlife to investigate vector competence, health impacts, and coinfection dynamics. Multi-disciplinary collaboration and community involvement in tick surveillance will advance our understanding of tick-borne filarial nematode ecology and epidemiology. Future research priorities include confirming vector competence where only molecular detection exists, investigating natural transmission patterns, and understanding the mechanisms influencing successful development in tick vectors. By addressing these knowledge gaps through systematic surveillance and experimental studies, we can better understand these complex host–parasite–vector relationships and their implications for animal health.

Graphical Abstract

Background

Ticks (Ixodida) are blood-feeding arthropods known to spread (i.e., vector) a range of pathogens of medical and veterinary importance [1, 2]. Ticks transmit a variety of disease-causing microbial taxa, spanning bacteria eliciting Lyme borreliosis, anaplasmosis, rickettsiosis, and ehrlichiosis [3,4,5]; viruses causing Powassan, Heartland, and tick-borne encephalitis infections [3, 6,7,8]; Babesia and Hepatozoon protozoan parasites causing babesiosis and hepatozoonosis, respectively [9,10,11]; and macroparasites, including filarial nematodes of medical and veterinary importance [12,13,14]. Filarial nematodes belonging to the superfamily Filarioidea Weinland, 1858 are further divided into two families: Filariidae Cobbold, 1864 and Onchocercidae Leiper, 1911. These nematodes have indirect life cycles, requiring arthropod intermediate hosts (biological vectors) to complete their development and transmission. Despite the diverse arthropod vectors utilized by filarial nematodes, only a few Onchocercidae species have been shown to use ticks as primary intermediate hosts [15]. Most Filarioidea rely on blood-sucking insects, such as mosquitoes, fleas, black flies, and biting midges, as primary vectors [16, 17]. Tick-transmitted filarioid nematodes show specific host associations; for example, Cercopithifilaria bainae infects domestic dogs and Monanema marmotae parasitizes groundhogs (Marmota monax) [12, 18]. Similarly, other arthropod-transmitted filariae demonstrate host specificity, including some filarioid species that are agents of neglected tropical diseases in humans, such as the black fly-transmitted Onchocerca volvulus and the mosquito-transmitted Wuchereria bancrofti in humans [19, 20].

Morphologically, filarial nematodes are elongated and threadlike, with long cylindrical bodies coated in a rigid cuticle with oral and anal openings, sensory papillae, and reproductive organs [16, 21, 22]. Nematode sexes are separated, with males and females sometimes being morphologically different; males may be shorter with coiled posterior ends, whereas females tend to be more elongated. Microfilariae are the initial larval stages of development of filarial nematodes, which develop in arthropod vectors into first-stage larva (L1), second-stage larvae (L2), and subsequently to third-stage larvae (L3), which is infective to the definitive host [16]. Vertebrate species typically serve as the definitive host for filarial nematodes, wherein adult nematodes attain sexual maturity and reproduce, with females producing fertilized eggs that hatch into microfilariae. Microfilariae can circulate throughout the vertebrate host body and are often ingested by arthropod vectors such as insects (Insecta) and acarines (Acari; ticks and mites) during blood-feeding activities, where the definitive host enables parasite reproduction and continued transmission [22,23,24].

The potential role of tick vectors in propagating filarial nematodes and affecting animal and human health remains understudied. Several studies have examined and identified filarial nematodes in both hard (Ixodidae) and soft (Argasidae) ticks [14, 25,26,27,28,29,30,31,32,33,34,35], thereby providing evidence that ticks can indeed vector filarial nematodes. Tick feeding strategies can significantly contribute to the biological transmission of filarial nematodes in several ways, including the duration of attachment, feeding preferences, and developmental stages. While slow-feeding ixodid ticks can remain attached to their hosts for several days or weeks, with some species feeding for up to 2 weeks [1], experimental studies by Brianti et al. [36] showed that extended feeding duration alone does not determine successful filarial nematode transmission. The typical development from microfilariae to infective L3 stage often takes longer than a single tick feeding period, usually around 30 days [36]. While this extended development time increases the likelihood of successful transmission to a new vertebrate host during subsequent feeding, reinfection of the same host during a single blood meal is unlikely. In addition, feeding patterns and host associations of ticks can play a crucial role in the life cycle of filarial nematodes [37]. For example, Ramos et al. [38] showed that Cercopithifilaria rugosicauda utilizes Ixodes ricinus nymphs to parasitize roe deer (Capreolus capreolus) in southern Italy. The timing of nymphal activity coincides with host availability, demonstrating how filarial transmission occurs through monotropic tick species—ticks that primarily parasitize specific host species.

While tick-borne filarial nematodes have been documented in various animal hosts, their impact on animal health remains poorly understood. Currently, there are no documented cases of tick-borne filarial nematodes affecting livestock production, and their economic significance, if any, requires further investigation. In companion animals, Cercopithifilaria spp. can infect dogs, with some naturally infected individuals developing dermal nodules, though these infections are rarely associated with clinical signs [23, 32, 36]. While treatments such as topical medications and corticosteroids are available, much remains to be understood about the pathogenicity and epidemiology of these infections [39].

In this review, we aim to synthesize and provide a comprehensive overview of the current knowledge on the global diversity of filarial nematode species that use ticks as vectors. Among the family Onchocercidae, development and transmission require arthropod intermediate hosts, with several genera specifically adapted to using ticks as vectors. Therefore, we focus on genera within Onchocercidae, reporting their known tick vectors and other arthropod vectors. In addition, using North America as a case study, we highlight significant gaps in the distribution and prevalence of tick-borne filarial nematodes and explore potential interactions between filarial nematodes and tick-borne pathogens and discuss their implications for infection dynamics and host immune responses. This review will cover the strengths and weaknesses of existing and current methods to detect filarial nematodes in tick vectors, emphasizing the need for targeted, large-scale research efforts focusing on known tick hotspots and risk interfaces. By synthesizing available information and critically assessing published literature, we aim to understand tick-borne filarial transmission among domestic and wildlife animals in natural ecosystems, with a focus on identifying knowledge gaps regarding these nematodes in North America to guide future research directions.

Roles of vertebrate hosts and tick vectors in the transmission of filarial nematodes

All vertebrate classes except fish serve as definitive hosts for filarial nematodes, enabling their development and transmission [40]. Vertebrate hosts harboring filarial nematodes allow the parasite to complete their life cycle, thus facilitating their transmission to tick vectors (Fig. 1). For instance, the life cycle of C. rugosicauda illustrates the synchronized timing between tick vectors and vertebrate hosts, which is essential for successful transmission. Development in I. ricinus takes 8 weeks for microfilariae to reach the infective L3 stage, followed by 24 weeks in roe deer for L3 to mature into adults [30, 38]. Adult nematodes produce microfilariae that concentrate in dermal tissues, coinciding with tick feeding periods and enabling continued transmission.

Fig. 1

Transmission cycle of filarial nematodes in tick vectors and vertebrate hosts. The cycle begins when a tick vector introduces L3 larvae into the definitive host (e.g., dogs, rodents, raccoons, and opossums) through skin bites. Within the host, L3 larvae develop into adult nematodes that mate and produce microfilariae, which circulate in the skin or blood of the host. When a new tick feeds on an infected host, it ingests the microfilariae, which then develop into infective L1 to L3 larvae within the tick, completing the transmission cycle. Illustration created on Biorender.com

The efficiency of filarial nematode transmission between vertebrate hosts and tick vectors may depend on host–vector compatibility. The availability of compatible hosts and vector feeding preferences may determine the geographic distribution of vector-borne helminths. For instance, domestic dogs serve as efficient reservoirs for filarial nematodes by simultaneously maintaining long-term infections with multiple species and their ability to sustain infections, which makes them consistent sources of microfilariae for brown dog ticks throughout the vector active season [14, 36].

Complexities of tick-borne filarial nematode transmission and development in vertebrate hosts

Early studies by Bain [28], Bain et al. [35], and Beaver and Burgdorfer [29] showed that the hindgut, hemocoel, fat cells, and salivary gland ducts of ticks provide a favorable environment for the development of certain filarial nematodes. In these cases, ticks ingest microfilariae when feeding on an infected vertebrate host [28] (Fig. 1). These microfilariae then migrate to specific tick tissues to find suitable conditions for growth and transformation into infective larvae. For example, many microfilariae migrate from the tick midgut to the hemocoel (i.e., body cavity), where they molt into L1, L2, and subsequently into infective L3, a process that may take 30 days [36]. L3 larvae have been documented in tick salivary glands [13, 36]; however, their transmission to vertebrate hosts via salivary glands is yet to be established. As ticks feed for several days, they regurgitate saliva into the bite site, potentially countering host hemostasis and inflammatory responses [22, 41,42,43]. Tick feeding can potentially deliver L3 larvae into the dermal skin layers of the host, setting off local reactions and enabling nematode transfer [18] (Fig. 1).

The concentration of microfilariae in the skin of infected vertebrates near common tick attachment sites (such as the ears) implies a specific attraction of filarial nematodes to chemical substances in tick saliva, as demonstrated by Ko [18]; a similar mechanism was proposed by Moorhouse [43]. If broadly applicable, this attraction could enhance the probability of microfilariae being ingested by the tick vector during blood feeding, facilitating their transmission to the vector and subsequently to new susceptible vertebrate hosts (Fig. 1). The specific vertebrate tissue harboring these transmitted microfilariae (e.g., skin or blood) corresponds to the feeding strategies of the arthropod vector on hosts [44]. For instance, microfilariae inhabiting subdermal skin layers may be ingested by ticks during their blood-feeding process. Alternatively, some circulating blood-borne microfilariae can be consumed through vessel lacerations by arthropod vectors such as mosquitoes and other flies that feed on blood using anticoagulant strategies. Thus, vertebrate microfilariae niche accessibility aligns with vector hematophagous behavior, which propagates initial establishment within intermediate hosts [45].

The life cycles of filarial nematodes within arthropod vectors are initiated when blood-feeding uptake introduces vertebrate-derived microfilariae into the vector system. The complex process by which nematodes develop from microfilariae to L3 within ticks involves multiple stages that occur in specific organs and tissues, with translocation through the stomach walls [22] (Fig. 1). While the role of saliva in tick-feeding has been well documented in tick-borne pathogens [46, 47], knowledge gaps exist on how filarial nematodes interact with and potentially impact tick organs. As microfilariae and developing larvae migrate through various tick tissues—from gut epithelium to hemocoel and salivary glands—they may affect tissue integrity and function. The efficiency of L3 from tick-to-host transfer could depend on these tissue interactions; aspects of this process remain elusive and require further investigation to comprehend the mechanisms at play.

Once in the definitive host, the infective L3 typically molts and migrates. Some L3 may move from the skin via lymphatic vessels to other vertebrate host tissues to continue development. Others may remain in the skin to mature into adults that mate and produce microfilariae, enabling acquisition by new ticks to perpetuate further transmission cycles [32] (Fig. 1). The immunogenic nature of incoming L3 larvae may influence the establishment of infection in the vertebrate host and host immune defenses at bite sites or downstream migration routes can impede nematode survival. Deciphering the fundamental processes governing these multifaceted vertebrate–tick–parasite associations may necessitate a deeper understanding of the complex interaction and impact on the circulation of filarial nematodes, as this affects host health.

Detection methods for tick-borne filarial nematodes

Detecting and screening tick vectors for filarial nematode infections is essential for assessing the prevalence of these nematodes in tick populations, identifying potential risk areas for transmission, and developing targeted control strategies [14, 36]. Various techniques, including dissection [36, 38], histopathological analyses [14], and molecular approaches, such as polymerase chain reaction (PCR) and DNA sequencing [12, 33] (Table 1), can be used to assess the status of tick infection with filarial nematodes. These methods differ in sensitivity, specificity, and practicality, making it essential to consider the strengths and weaknesses of each approach when designing studies or surveillance programs.

Table 1 Comprehensive assessment of methodologies employed in the detection of filarial nematodes in tick vectors

Classical approaches to detecting tick-borne filarial nematodes

Dissection and microscopic examination of ticks represent a classical means of investigating filarial infection and developmental progression within different tick tissues (Fig. 2A). Using these methodologies, filarial nematodes have been observed in hard tick species, including Amblyomma cajennense [31], Haemaphysalis flava, H. japonica [48], Ixodes cookei [18, 49], I. scapularis [29], and I. ricinus [30, 35, 38, 50]. Soft ticks, such as Ornithodoros tartakovskyi [51] and O. talaje [52], have also been found to carry filarial nematodes. Initial investigations into filarial nematode infections in ticks relied upon microscopic visualization methods for identification and morphological diagnosis. While microscopy and dissection techniques provided early verification that ticks harbor filarial nematodes, these methods have limitations in species identification. Morphological identification of filarial larvae is challenging owing to similar anatomical features among species and cryptic species complexes [53], particularly in early developmental stages. Therefore, while genus-level identification is possible through microscopy and dissection, molecular methods are typically needed for species-level determination.

Fig. 2

Complementary approaches for detecting and characterizing filarial nematodes in tick vectors. (A) Classical methods involve invasive techniques such as dissecting and microscopically examining excised tick tissues to visualize and quantify filarial nematodes; (B) noninvasive imaging techniques, such as X-ray micro-computed tomography (micro-CT), allow for the visualization and localization of filarial nematodes within tick samples without dissection, generating high-resolution two-dimensional (2D) and three-dimensional (3D) images; (C) molecular techniques, such as DNA/RNA extraction (often requiring the destruction of the sample), followed by PCR, sequencing, or next-generation sequencing and computational analysis, enable the identification and characterization of filarial nematodes at the nucleotide level. Illustration created on Biorender.com

Patton et al. [54] established comprehensive protocols for dissecting tick organs and tissues, and for the collection of biofluids (i.e., hemolymph, saliva), enabling examination of tick internal structures for the presence of pathogens. These methods, later applied by Santos et al. [55] to study filarial nematodes, can be adapted to target specific tissues where these parasites develop, facilitating visual identification of larvae within ticks. Studies have demonstrated the presence of filarial nematodes in tick organs, such as the hindgut, hemocoel, fat cells, and salivary ducts [18, 29, 35, 36]. Controlled infection experiments have directly validated the ability of specific tick genera, including Amblyomma, Ixodes, and Rhipicephalus, to effectively acquire and transmit filarial nematodes [18, 31, 36, 49, 56, 57]. Despite being constrained by morphological assessment limitations and operator expertise, adopting standardized dissection and preparation protocols would result in more consistent documentation of tissue distributions and quantification of tick-borne filarial burden. Thus far, no definitive reports have explicitly validated the precise anatomical locations occupied by developing L1–L3 filarial nematodes inside tick vectors. Expanding confirmation of occupied tick niches through other methods, such as targeted imaging and molecular approaches, may enable tracking of filarial occupation across tick tissues, thereby elucidating vector–parasite interface dynamics and transmission.

Imaging techniques for detecting tick-borne filarial nematodes

The emergence of advanced biological imaging techniques, such as 3D X-ray microcomputed tomography (micro-CT), has ushered in exciting new potential for visualizing filarial nematode stages within arthropod vectors. Micro-CT instruments use X-ray beams to irradiate targeted samples; detectors capture unabsorbed photon signals to create complex 3D renderings that specialized software programs can analyze [58, 59] (Fig. 2B). This approach facilitates the examination of filarial nematodes inside arthropod vectors with minimal physical disruption of tissues. While some sample preparation involves fixation or contrast enhancement through staining, these processes are generally less invasive than traditional dissection methods.

The utilization of 3D X-ray micro-CT, supported by histopathological analysis, has facilitated the observation of Acanthocheilonema spirocauda larvae within the fat bodies (L1) and hemocoels (L3) of seal lice (Anoplura: Echinophthirius horridus) [58, 60]. Acanthocheilonema spirocauda is a filarial nematode that primarily infects seals and sea lions as definitive hosts. Hall et al. [61] used micro-CT imaging to visualize Onchocerca L1 larvae embedded within blackfly thoracic musculature. Previous studies demonstrated the potential of micro-CT imaging to reveal the precise spatiotemporal details of infection progression within arthropod vectors [58, 60, 61]. Application of micro-CT could help reveal favorable habitats, migratory routes, and life cycle transitions of larval stages of filarial nematodes in tick vectors.

Molecular techniques for detecting tick-borne filarial nematodes

Cutting-edge molecular techniques have enabled more precise identification of nematode species and isolates transmitted by tick vectors globally (Table 1) (Fig. 2C). For example, genetic analyses have confirmed filarial nematodes in ticks such as I. ricinus [38] and R. sanguineus sensu lato (s.l.) [62,63,64]. These investigations employed molecular techniques, including DNA extraction, PCR, sequencing, and in some cases, alternative genetic assays such as metabarcoding, to identify filarial nematodes in ticks (Table 1). Molecular identification of filarial nematodes in tick vectors often relies on genetic markers such as 12S mitochondrial ribosomal RNA and cytochrome c oxidase subunit I [12, 24, 33, 34, 65]. When proper reference material is available, these markers allow accurate identification at both genus and species levels, particularly for distinguishing closely related species.

Employing PCR-based assays targeting specific filarial genes consistently facilitated the identification of prominent Onchocercidae species, such as those in the genera Cercopithifilaria, Monanema, and Acanthocheilonema, across numerous hard tick species (Table 2) [13, 62, 65,66,67]. These genetic screening techniques have enabled extensive detection of tick-borne filarial nematodes across tick vector populations, revealing the presence of pathogenic filariae spanning multiple tick genera and species. In North America, genetic analyses have enabled the discovery of filarial nematode DNA in several tick species. However, it is important to note that these molecular techniques cannot detect viable parasites or their developmental stages. There is a risk of detecting remnant DNA or recently ingested microfilariae from blood meals rather than active infections. Therefore, to confirm the biological relevance of these findings and avoid spurious detections, molecular assays should be combined with other methods, such as microscopy, 3D X-ray micro-CT, or experimental transmission studies. For example, molecular screening and high throughput sequencing (HTS) have facilitated filarial detection and species identification in I. scapularis, substantiated through multiple reports [3, 4, 13, 66]. Similar findings confirming the presence of filarial nematodes using comparable molecular techniques were also obtained in A. americanum ticks through studies by Zhang et al. [25] and Henning et al. [67] and in R. sanguineus s.l. by Lineberry et al. [33, 34].

Table 2 Global report of filarioid species recovered from tick vectors and their potential hosts

In Europe, C. rugosicauda was initially detected in the hard tick I. ricinus through dissection and morphological studies across Austria, Germany, and France [30, 35, 50]. Later, Ramos et al. [38] used both morphological and molecular techniques, specifically dissection, microscopy, PCR, and sequencing, to detect C. rugosicauda L3 in I. ricinus nymphs from southern Italy, confirming the distribution of this nematode across Europe. Similarly, Binetruy and Duron [24] used molecular methods to detect filarial nematodes in ticks collected from French Guiana, South America. They discovered filarial species, including representatives from the genera Cercopithifilaria, Cruorifilaria, and putative Dipetalonema-related nematodes, inhabiting Amblyomma, Ixodes, and Rhipicephalus ticks in the region. Further characterization through multi-locus sequence analysis by Bruley and Duron [68] provided additional insights into tick-borne filarial diversity in this region.

Filarial nematodes and tick microbiome interactions

There is growing interest in understanding how the tick microbiome, which refers to the diverse community of microorganisms residing within the tick, contributes to modifying tick life cycles as well as pathogen and parasite acquisition and transmission. Ticks harbor a diverse microbiome comprising endogenous and exogenous bacteria, fungi, protozoa, viruses, and filarial nematodes that can colonize tick organs [3, 55, 69]. Owing to their hematophagous lifestyle, host tissue salivation, and frequent host switching, ticks provide suitable habitats for various microbes [70]. Ticks are vectors of numerous medical and veterinary pathogens, including bacteria (such as Rickettsiales, Anaplasmataceae, and spirochetes), protozoa, and viruses, and individual ticks may be concurrently infected (coinfected) with multiple microbial pathogens and filarial nematode larvae [3, 69]. Despite the frequent occurrence of multiple infections, most investigations typically examined tick-borne microbes [71, 72] and filarial nematodes in ticks individually rather than as an integrated multi-kingdom community [14, 24, 33, 66].

Coinfection adds complexity to the tick microbiome, creating interactions that may be mutualistic, antagonistic, or manipulative between coexisting organisms [73] (Fig. 3). While studies in other arthropod vectors show that such interactions can affect transmission dynamics and host health [74, 75], relationships between tick-borne pathogens and filarial nematodes remain largely unexplored. One documented example is the association between filarial nematodes and endosymbiotic Wolbachia bacteria in tick vectors [3, 25]. Using high-throughput sequencing, Cross et al. [3] demonstrated a significant positive correlation between onchocercid filarial nematodes and Wolbachia in I. scapularis ticks from Wisconsin. Understanding these and similar relationships could provide insights into disease transmission patterns and tick physiology.

Fig. 3

Potential interactions between tick microbiome and pathogen transmission across tick tissues. The tick microbiome, consisting of a diverse array of microorganisms acquired from the vertebrate host during blood feeding, colonizes different regions of the tick gut. These microbes can potentially influence pathogen acquisition, maintenance, and transmission through diverse biological interactions, ultimately shaping pathogen transmission dynamics from the tick vector to the vertebrate host. Understanding the complex interplay between the tick microbiome and pathogen transmission is essential for developing effective tick-borne disease control strategies. SG, salivary glands; HM, hemocoel. Illustration created on Biorender.com and adapted from [92]

Studies have documented co-occurrence of filarial nematodes with other pathogens in tick vectors. For instance, Zhang et al. [26] found filarial nematodes alongside other pathogens in A. americanum ticks, while Ramos et al. [69] and Santos et al. [55] observed C. bainae co-occurring with Hepatozoon canis in R. sanguineus s.l. However, while these studies document co-occurrence, the potential interactions between these organisms and their influence on transmission dynamics and vector competence remain to be determined.

Potential interactions between filarial nematodes and other tick-borne pathogens

The potential impact of filarial nematodes on tick-borne disease transmission remains poorly understood, despite the well-established role of ticks as vectors of numerous pathogens in humans and animals. This interplay between filarial nematodes and established tick-borne pathogen transmission systems presents an important area for further research. Ticks transmit a wide range of disease-causing agents, including Borrelia burgdorferi, which causes Lyme disease, Rickettsia rickettsii, which is responsible for Rocky Mountain spotted fever, and Anaplasma species, which leads to anaplasmosis [76, 77]. Key tick anatomical structures (e.g., guts, hemocoel, and salivary glands) serve as critical interfaces between the tick, its pathogens, and the vertebrate host (Fig. 3). These tissues provide distinct microenvironments potentially affecting pathogen and parasite establishment and development. While interactions between tick-borne pathogens and tick tissues have been well studied [78, 79], how filarial nematodes interact with these pathogens and tick tissues [22] requires further investigation to know the mechanism at play. These interactions likely vary with vertebrate host species, tick species, tick developmental stages and sex, and blood-feeding status (i.e., engorged versus nonengorged ticks).

As filarial nematodes migrate through tick tissues, they encounter various pathogens within the tick microbiome [80,81,82]. These interactions occur in different tick compartments, particularly in the hindgut, where many pathogens undergo critical developmental processes and transovarial transmission (i.e., the passage of pathogens from an adult female tick to her eggs). The presence of filarial nematodes during migration through these tissues may influence pathogen distribution and the local microbiome composition in ways that could affect transmission dynamics (Fig. 3). However, the specific mechanisms and consequences of these interactions remain largely unexplored.

Tick vectors of filarial nematodes: global perspective, current evidence, and research needs

Members of the superfamily Filarioidea (families Filariidae and Onchocercidae) parasitize primarily terrestrial vertebrate species, including mammals, birds, reptiles, and amphibians. The Onchocercidae show broader vector associations than Filariidae [45], with several genera specifically adapted to tick transmission [16]. Our current understanding of tick-borne filarial diversity and distribution remains incomplete [13, 33, 67], with five genera (Cercopithifilaria, Cherylia, Cruorifilaria, Monanema, and Yatesia) documented as using ticks as vectors (Table 3). Some of these genera may also utilize other arthropod vectors, reflecting their adaptability to different transmission routes (Table 4).

Table 3 List of known tick-borne filarial nematode genera (family Onchocercidae), their vertebrate hosts, and geographic regions

Table 4 Filarial nematode genera with unknown or non-tick vectors listed by family and genus, vertebrate and intermediate hosts, and their geographic regions

Vector competence has been experimentally shown for several filarial species through demonstrated development to infective stages (Table 5). Cercopithifilaria bainae and C. grassii develop successfully in R. sanguineus s.l., which serves as their primary vector worldwide [32, 35, 83], while C. rugosicauda shows specific adaptation to I. ricinus [30, 38]. Among Monanema species, confirmed vector relationships include M. marmotae in I. cookei, M. globulosa in Haemaphysalis leachi, and M. martini in multiple tick species (R. sanguineus s.l., R. turanicus, and Hyalomma truncatum) [18, 57, 84]. Acanthocheilonema viteae represents the only species in its genus with confirmed development in tick vectors, specifically the Ornithodoros species [51, 85, 86], although Acanthocheilonema DNA has been detected in other tick species without confirmed vector competence. Experimental studies have also demonstrated Yatesia hydrochoerus development in A. cajennense complex ticks, and C. guyanensis shows laboratory development in I. ricinus, although the latter represents a non-natural vector relationship [31, 87].

Table 5 Filarial nematodes and their tick–vector associations: host relationships, evidence for vector competence, and relevant references

These relationships likely reflect ecological adaptations, where tick feeding behaviors—including wide host range and prolonged attachment—facilitate filarial transmission. For instance, the widespread distribution of Cercopithifilaria species appears to be facilitated by the association with cosmopolitan tick vectors such as R. sanguineus s.l. [14]. Molecular detection methods have identified additional potential tick-filarial associations awaiting confirmation of vector competence, including Cruorifilaria DNA in Amblyomma species and Monanema-related sequences in various tick species [13, 24, 25, 66, 67]. Research priorities include confirming vector competence where only molecular detection exists and investigating natural transmission patterns in endemic areas. Studies should focus on documenting complete developmental cycles in tick vectors, identifying factors that influence successful transmission, and understanding the specificity of tick-filarial relationships. Standardized methods are needed to differentiate between transient presence of filarial DNA from blood meals and established infections in tick tissues.

Potential factors limiting tick-borne transmission of major filarial nematodes

The transmission of filarial nematodes, particularly the well-studied genera Brugia, Dirofilaria, and Onchocerca, has been predominantly associated with insect vectors (Table 4). For example, Onchocerca species develop in black flies and biting midges [16], while Dirofilaria and Brugia species use mosquitoes as their primary vectors [88]. The apparent inability of ticks to transmit these nematodes raises intriguing questions about vector–parasite specificity and the evolutionary processes shaping these relationships. Some filarial nematodes may depend on their arthropod vectors extensively, leading to highly specialized interactions. While this dependency may provide plausible explanations for the absence of tick-borne transmission in some filarial nematodes, it is important to consider anatomical and physiological limitations. Dipterans possess distinct circulatory systems and specific tissues, such as Malpighian tubules, which are crucial for filarial larval development in some species [89]. In contrast, ticks have an open circulatory system and different organ arrangements [1]. The internal structure of ticks, lacking specific organs found in insects, may not provide the necessary anatomical environment for the development of some filarial nematodes. The feeding behavior of vectors is another critical factor. While mosquitoes and other blood-sucking insects typically feed quickly, aligning with the transmission strategy of many filarial nematodes, hard ticks feed slowly over several days [1]. This prolonged feeding pattern might be incompatible with the life cycle and transmission dynamics of some filarial species that have evolved to be transmitted during brief feeding events. The feeding duration mismatch could disrupt the delicate timing for successful filarial nematode transmission and establishment. Ticks possess distinct innate immune responses that differ from insects [90, 91]. However, the specific role of tick immunity in filarial nematode establishment and development remains to be investigated through experimental studies.

Knowledge gaps of tick-borne filarial nematodes in North America

North America lacks thorough investigations of tick-borne filarial associations and transmission dynamics among endemic tick vectors and vertebrate hosts. The only known filarial nematodes documented in North America belong to the genera Acanthocheilonema, Cercopithifilaria, and Monanema (Table 2), with ixodid tick vectors implicated in transmitting these filarial nematodes to their definitive hosts. In contrast to findings from South America, Europe, Africa, Australia, and Asia, this limited evidence highlights substantial knowledge gaps in the biodiversity, host associations (both definitive and intermediate), and geographic distribution of tick-borne filarial nematodes across North America. Expanded surveillance of filarial nematodes in North American ticks is warranted to characterize the distribution and diversity of unknown tick-borne parasites and their potential tick vectors (Table 4).

Conclusions

This review highlights our current understanding of tick-borne filarial nematodes and identifies critical knowledge gaps. Key priorities include determining the diversity and distribution of these parasites through systematic surveillance, investigating vector competence and transmission dynamics, and understanding how coinfections with other tick-borne pathogens affect disease ecology. Research initiatives should focus on comprehensive surveys using modern detection methods, conducting experimental transmission studies, and investigating host–parasite–vector relationships. Through collaborative research efforts across disciplines, we can better understand these complex relationships and develop effective control strategies.