The Role of Hematophagous Arthropods, Other Than Mosquitoes and Ticks, in Arbovirus Transmission

Blitvich, Bradley J.

doi:10.3390/v17070932

Open AccessReview

The Role of Hematophagous Arthropods, Other Than Mosquitoes and Ticks, in Arbovirus Transmission

by

Bradley J. Blitvich

Department of Veterinary Microbiology and Preventive Medicine, College of Veterinary Medicine, Iowa State University, 2116 Veterinary Medicine, Ames, IA 50011, USA

Viruses 2025, 17(7), 932; https://doi.org/10.3390/v17070932

Submission received: 15 May 2025 / Revised: 16 June 2025 / Accepted: 26 June 2025 / Published: 30 June 2025

(This article belongs to the Section Invertebrate Viruses)

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Abstract

Arthropod-borne viruses (arboviruses) significantly impact human, domestic animal, and wildlife health. While most arboviruses are transmitted to vertebrate hosts by blood-feeding mosquitoes and ticks, a growing body of evidence highlights the importance of other hematophagous arthropods in arboviral transmission. These lesser-known vectors, while often overlooked, can play crucial roles in the maintenance, amplification, and spread of arboviruses. This review summarizes our understanding of hematophagous arthropods, other than mosquitoes and ticks, in arboviral transmission, as well as their associations with non-arboviral viruses. Thirteen arthropod groups are discussed: bat flies, blackflies, cimicids (bat bugs, bed bugs, and bird bugs), Culicoides midges, fleas, hippoboscid flies, lice, mites, muscid flies (including horn flies and stable flies), phlebotomine sandflies, tabanids (including deer flies and horse flies), triatomines, and tsetse flies. Some of these arthropods are regarded as known or likely arboviral vectors, while others have no known role in arbovirus transmission. Particular attention is given to species associated with arboviruses of medical and veterinary significance. As the burden of arboviruses continues to grow, it is critical not to overlook the potential contribution of these lesser-known vectors.

Keywords:

arbovirus; arthropod; hematophagous; transmission; vector competence

1. Arboviruses

The term arthropod-borne virus (arbovirus) is a non-taxonomic classification that encompasses a diverse group of viruses that are transmitted by bite to susceptible vertebrate hosts by hematophagous (blood-feeding) arthropods. According to the recent comprehensive arbovirus catalog compiled by Huang and colleagues, there are 460 species (615 species/subtypes) of arboviruses [1]. These viruses belong to multiple families, but the majority are classified within the Flaviviridae, Nairoviridae, Peribunyaviridae, Phenuiviridae, Rhabdoviridae, Sedoreoviridae, and Togaviridae [1,2,3]. A defining characteristic of arboviruses is their ability to replicate in both arthropod vectors and vertebrate hosts. Many arboviruses are zoonotic and have caused major disease outbreaks among humans, domestic animals, and wildlife [2,3,4,5,6,7,8].

The most prevalent arboviral diseases are transmitted by mosquitoes and ticks. Mosquito-borne arboviruses of global public health importance include all four serotypes of dengue virus (Flaviviridae), chikungunya virus (Togaviridae), Japanese encephalitis virus (Flaviviridae), West Nile virus (WNV; Flaviviridae), yellow fever virus (Flaviviridae), and Zika virus (Flaviviridae) [9,10,11,12,13,14]. Tick-borne arboviruses of regional concern include Crimean–Congo hemorrhagic fever virus (CCHFV; Nairoviridae) in Africa, Asia, and Europe; Kyasanur Forest disease virus (KFDV; Flaviviridae) in Asia; Powassan virus (Flaviviridae) in North America and Russia; Rift Valley fever virus (RVFV; Phenuiviridae) in Africa; severe fever with thrombocytopenia syndrome virus (SFTSV; Peribunyaviridae) in Asia; and tick-borne encephalitis virus (TBEV; Flaviviridae) in Europe and Asia [15,16,17,18,19]. All known arboviruses have RNA genomes, except for African swine fever virus (ASFV; Asfarviridae) [20]. Viruses that cannot replicate in arthropods but are mechanically transmitted by them (i.e., via contaminated mouthparts) are not considered to be arboviruses [21].

2. Hematophagous Arthropods and Vector Competence

Hematophagous arthropods are taxonomically diverse, belonging to multiple families and orders (Table 1). While some hematophagous arthropods transmit viruses to vertebrates, many have no confirmed role in virus transmission. Some hematophagous arthropods also transmit bacteria, protozoa, and helminths [22,23,24]. To be regarded as an epidemiologically important arboviral vector, an arthropod must meet several key criteria: (i) the virus must replicate to sufficient levels within the arthropod’s midgut, disseminate to the salivary glands, and be transmitted via saliva during blood-feeding; (ii) the arthropod and virus must co-occur temporally and spatially; (iii) the arthropod must feed on the relevant vertebrate host; and (iv) a significant proportion of the arthropod population must be found infected with the virus in natural settings [25,26,27].

Table 1. A list of all known hematophagous arthropods and their taxonomic classification.

Arthropod	Family	Order
Bat flies	Nycteribiidae and Streblidae	Diptera
Blackflies	Simuliidae	Diptera
¹ Blowflies	Calliphoridae	Diptera
Cimicids	Cimicidae	Hemiptera
Culicoides midges	Ceratopogonidae	Diptera
Fleas	Multiple families	Siphonaptera
Hippoboscid flies	Hippoboscidae	Diptera
Lice	Multiple families	Psocodea
Mites	Multiple families	Acariformes and Parasitiformes
¹ Mosquitoes	Culicidae	Diptera
Muscid flies	Muscidae	Diptera
Phlebotomine sandflies	Psychodidae	Diptera
Tabanids	Tabanidae	Diptera
¹ Ticks	Argasidae, Nuttalliellidae, and Ixodidae	Ixodida
Triatomines	Reduviidae	Hemiptera
Tsetse flies	Glossinidae	Diptera

¹ Not covered in this review.

Vector competence experiments are performed to measure the ability of an arthropod to support the replication and dissemination of an arbovirus and subsequent transmission of the virus to a susceptible host [27,28,29,30,31,32,33]. The arthropod is usually exposed to the virus by allowing it to feed upon a viremic animal or infectious artificial blood meal or by direct needle inoculation into the thorax. One limitation of the intrathoracic inoculation technique is that it bypasses the natural midgut barrier, a critical component of the natural infection route [34,35]. After an appropriate extrinsic incubation period, the arthropod is allowed to feed upon a naïve vertebrate host, which is then monitored for evidence of viral infection. The extrinsic incubation period is defined as the time it takes for a virus to disseminate from the midgut to the salivary glands of the arthropod [28,30]. Infection, dissemination, and transmission rates are typically defined as the percentage of engorged arthropods that contain virus in their midguts, secondary organs (i.e., legs or wings), and saliva, respectively. The transmission rate can also be defined as the percentage of infected arthropods that successfully transmit to virus to an uninfected vertebrate host.

Importantly, the detection of a virus in an arthropod is not evidence that the virus can replicate in the arthropod. An arthropod can acquire a virus through a blood meal (either in the field or laboratory) yet be refractory to viral replication. If a virus persists in an arthropod for no longer than one or a few days, viral replication most likely did not occur. An increase in viral titer or the amount of viral RNA in an arthropod over time is strong evidence of active viral replication [36].

The role of mosquitoes and ticks in arboviral transmission has been extensively reviewed elsewhere and will not be addressed here [5,37,38,39,40,41,42,43]. This review instead summarizes our current knowledge regarding the involvement of other hematophagous arthropods in arbovirus transmission because there is a growing body of evidence to suggest that some have an important role in arboviral transmission. Thirteen arthropod groups are covered: bat flies, blackflies, cimicids, Culicoides midges, fleas, hippoboscid flies, lice, mites, muscid flies, phlebotomine sandflies, tabanids, triatomines, and tsetse flies. With the exception of mosquitoes and ticks, the only hematophagous arthropods not included in this review are blowflies (Calliphoridae). While blowflies are primarily scavengers that feed upon decaying organic matter, some species feed on blood during their larval stage [44,45]. However, blowflies are excluded from this review because the majority are not blood-feeders, and none have been implicated in the biological transmission of arboviruses.

Each group of arthropods discussed in this review is evaluated based on its association with (i) recognized arboviruses, (ii) other vertebrate-infecting viruses, and (iii) viruses not known to infect vertebrates. For the purpose of this review, a “recognized arbovirus” refers to any virus listed as such in the recent arbovirus catalog compiled by Huang and colleagues, which integrates data from the Arbovirus Catalog, the arbovirus list in Section VIII-F of the Biosafety in Microbiological and Biomedical Laboratories (6th edition), Virus Metadata Resource of International Committee on Taxonomy of Viruses, and GenBank [1]. “Other vertebrate-infecting viruses” are non-arboviral viruses known to infect vertebrates or vertebrate cell cultures or for which there is evidence for viral replication due to the detection of virus-derived small RNAs in vertebrate cells [46]. “Non-vertebrate-infecting viruses” are those for which there is no evidence of vertebrate or vertebrate cell infection.

The arthropods discussed in this review can be separated into three categories based on their roles in arbovirus transmission:

Arthropods known or suspected to be epidemiologically important biological vectors of arboviruses, i.e., blackflies, cimicids, Culicoides midges, and phlebotomine sandflies. These arthropods have yielded multiple isolations of the relevant viruses in field studies and have transmitted the virus under experimental conditions, while arthropods from no other groups have been identified as major vectors.
Arthropods that may play a minor role in the biological transmission of arboviruses, i.e., fleas and mites. These arthropods have also yielded multiple isolations of relevant virus and have transmitted the virus under experimental conditions, but other arthropods have been identified as major vectors.
Arthropods where there is insufficient evidence to suggest that they are biological vectors of arboviruses, i.e., bat flies, hippoboscid flies, lice, muscid flies, tabanids, triatomines, and tsetse flies. None of these arthropods have been shown to transmit arboviruses under experimental conditions.

This review is important because it addresses a significant gap in the arbovirology literature by systematically evaluating the potential role of non-mosquito, non-tick hematophagous arthropods in arbovirus transmission. While these arthropods have received far less attention, growing evidence suggests that some may serve as important or overlooked vectors. This review provides a foundation for future research efforts aimed at understanding the broader ecological dynamics of arboviral transmission and improving disease surveillance and control strategies.

3. Bat Flies

Bat flies (Nycteribiidae and Streblidae) are wingless ectoparasites that feed exclusively on the blood of bats [47]. The Nycteribiidae contains at least 275 species across 21 genera, whereas Streblidae comprises approximately 227 species in 31 genera. Bat flies primarily inhabit tropical and subtropical regions, with nycteribiids occurring mainly in the Eastern Hemisphere and streblids in the Western Hemisphere. Bat flies are not considered biological vectors of viruses of medical or veterinary importance, although five viruses capable of replicating in vertebrate cells have been detected in these arthropods: dengue virus 2 (DENV2; Flaviviridae), Kaeng Khoi virus (KKV; Perbunyaviridae), Mahlapitsi virus (MAHLV; Sedoreoviridae), Nelson Bay reovirus (NBV; Sedoreoviridae), and Wolkberg virus (WBV; Peribunyaviridae) (Table 2) [48,49,50,51,52,53]. DENV2 and KKV are recognized arboviruses, but the others are not [1].

3.1. Recognized Arboviruses

DENV2 was detected by RT-PCR in bat flies (Strebla wiedemanni and Trichobius parasiticus) collected from common vampire bats (Desmodus rotundus) in Mexico [53]. DENV2 was also found in tissues from the infested vampire bats. KKV was originally isolated from a wrinkle-lipped free-tailed bat (Chaerephon plicatus; aka Tadarida plicata) in Thailand [54]. Subsequent isolations were made from bat flies (Eucampsipoda sundaica) parasitizing Leschenault’s rousette fruit bats (Rousettus leschenaultii) in China and from C. plicatus bats in Cambodia [48,51,55]. KKV has also been isolated from cimicids (discussed later in this review). There is insufficient evidence to support the role of bat flies as vectors of DENV2 or KKV because vector competence studies have not been conducted.

Table 2. Arboviruses and other vertebrate-infecting viruses identified in field-collected bat flies.

Virus	Species Name	Abbreviation	Family	Recognized Arbovirus	¹ Experimental Evidence of Bat Fly-Borne Virus Transmission	Citation
Dengue virus 2	Orthoflavivirus denguei	DENV2	Flaviviridae	Yes	² -	[53]
Kaeng Khoi virus	Orthobunyavirus kaengkhoiense	KKV	Perbunyaviridae	Yes	-	[48,51]
Mahlapitsi virus	Orthoreovirus mahlapitsiense	MAHLV	Sedoreoviridae	-	-	[49]
Nelson Bay reovirus	Orthoreovirus nelsonense	NBV	Sedoreoviridae	-	-	[52,56]
Wolkberg virus	Orthobunyavirus wolkbergense	WBV	Peribunyaviridae	-	-	[50]

¹ Excludes mechanical transmission. ² No.

3.2. Other Vertebrate-Infecting Viruses

MAHLV and WBV were originally isolated from bat flies (Eucampsipoda africana) collected from Egyptian fruit bats (Rousettus aegyptiacus) in South Africa, while NBV was originally isolated from a flying fox (Pteropus poliocephalus) in Australia [49,50,56]. Additional isolations of NBV were made from bat flies (E. sundaica) and bats (R. leschenaultii) in China [52]. All three viruses replicate in vertebrate cells and NBV causes disease in experimentally inoculated mice [49,50,52,56]. Additionally, a closely related NBV-like virus has been implicated in cases of acute respiratory distress in humans [57]. Vector competence experiments have not been performed to determine whether bat flies are capable of transmitting these viruses.

3.3. Viruses Not Known to Infect Vertebrates

Other viruses have been detected in bat flies using metagenomics, but none are known to replicate in vertebrate cells [58,59,60,61,62,63]. One example is Kanyawara virus (Rhabdoviridae), which was discovered in Dipseliopoda spp. bat flies parasitizing Myonycteris spp. bats in Uganda [59]. A subsequent study detected Kanyawara virus in an oral swab from an Angolan soft-furred fruit bat (Lissonycteris angolensis) infested with bat flies [58]. In the same study, the closely related Bughendera virus (Rhabdoviridae) was detected in a Dipseliopoda spp. bat fly. Additional novel viruses were detected in bat flies in China, Mexico, Nigeria, and Uganda [60,61,62,63]. In another study, novel rhabdoviruses were detected in bat flies (Nycteribia kolenatii, Nycteribia schmidlii, and Penicillidia conspicua) in Spain by RT-PCR using rhabdovirus-specific primers, but none have been demonstrated to replicate in vertebrate cells [64].

4. Blackflies

There are more than 2200 species of blackflies (Simuliidae) and most are hematophagous [65]. These arthropods occur on every continent, except Antarctica. Blackflies are vectors of several filarial nematodes and protozoan pathogens, including the etiological agent of human onchocerciasis (also known as river blindness) [66,67,68]. More pertinent to this review, blackflies harbor many viruses, including five recognized arboviruses: eastern equine encephalitis virus (EEEV; Togaviridae), Rift Valley fever virus (RVFV; Phenuiviridae), Venezuelan equine encephalitis virus (VEEV; Togaviridae), Vesicular stomatitis Indiana virus (VSIV; Rhabdoviridae), and Vesicular stomatitis New Jersey virus (VSNJV; Rhabdoviridae) (Table 3).

4.1. Recognized Arboviruses

Blackflies have been implicated as major biological vectors of vesicular stomatitis (VS), an agriculturally important disease of hoofed livestock in the Americas [69,70,71,72]. VS is caused by VSNJV and VSIV, known collectively as vesicular stomatitis virus (VSV). VSV has been frequently detected in blackflies during epizootics, with several studies reporting high prevalence rates [73,74,75,76,77,78,79]. For example, VSV was detected in 16 of 319 pools containing 1215 blackflies in California, the U.S. [75]. Blackflies of at least four species were positive for the virus: Simulium argus, Simulium hippovorum, Simulium tescorum, and Simulium vittatum complex. Infection rates were highest for S. hippovorum and S. vittatum complex. Additionally, VSV was detected in 11 of 77 pools of blackflies in New Mexico, the U.S. [77]. Over half of the positive pools contained Simulium mediovittatum, while the others contained Simulium meridionale, Simulium notatum/griseum, Simulium robynae, and an unidentified Simulium spp. In contrast, there was a low prevalence of VSV in blackflies in Chiapas, Mexico [80]. In this study, hematophagous arthropods (blackflies, midges, mosquitoes, and sandflies) were collected on cattle ranches with a history of VS cases. VSV was detected in 102 of 874 pools, but most positive pools contained mosquitoes and sandflies. Only one VSV-positive blackfly pool was detected, indicating that these arthropods are not major vectors of VSV in the study area.

Some species of blackflies are competent vectors of VSV [81,82,83,84,85,86]. In one study, 70% of colonized Simulium vittatum intrathoracically inoculated with VSNJV contained virus in their saliva at 10 days post-inoculation [85]. Following oral exposure, 45% of S. vittatum had detectable VSNJV in their saliva. Field-collected Simulium bivittatum, Simulium longithallum, and S. vittatum from Arizona, the U.S. were orally challenged with VSNJV strains from both Arizona and Oaxaca, Mexico [84]. VSNJV was detected in the saliva of 23% and 26% of S. notatum exposed to the Oaxaca and Arizona strains, respectively; however, neither viral strain disseminated to the salivary glands of S. bivittatum or S. longithallum. VSIV was also detected in the saliva of both colonized and wild S. notatum and S. vittatum at 10 days post-oral challenge [86]. Colonized S. vittatum intrathoracically inoculated with VSNJV successfully transmitted the virus to mice, pigs, and cattle [81,82,83]. Additionally, VSNJV was transmitted to uninfected S. vittatum co-feeding on the same nonviremic mice as infected blackflies [87].

EEEV, RVFV, and VEEV have occasionally been detected in blackflies. EEEV was isolated from one pool each of Simulium johannseni (aka Eusimulium johannseni) and Simulium meridionale in Wisconsin, the U.S. [88]. RVFV was isolated from at least one pool of Simulium blackflies (species not specified) in South Africa [89]. In Colombia, VEEV was isolated from 11 pools of blackflies (Simulium cauicumn, Simulium exiguuma, Simulium metallicum, Simulium mexicanuma, and Simulium payneih) [90]. Vector competence experiments have not been performed to determine whether blackflies are capable of transmitting any of these viruses.

Table 3. Arboviruses and other vertebrate-infecting viruses identified in field-collected blackflies.

Virus	Species Name	Abbreviation	Family	Recognized Arbovirus	¹ Experimental Evidence of Blackfly-Borne Virus Transmission	Citation
Eastern equine encephalitis virus	Alphavirus eastern	EEEV	Togaviridae	Yes	² -	[88]
Hepatitis B virus	Orthohepadnavirus hominoidei	HBV	Hepadnaviridae	-	-	[91]
Rift Valley fever virus	Phlebovirus riftense	RVFV	Phenuiviridae	Yes	-	[89]
Venezuelan equine encephalitis virus	Alphavirus venezuelan	VEEV	Togaviridae	Yes	-	[90]
Vesicular stomatitis Indiana virus	Vesiculovirus indiana	VSIV	Rhabdoviridae	Yes	-	[78]
Vesicular stomatitis New Jersey virus	Vesiculovirus newjersey	VSNJV	Rhabdoviridae	Yes	Yes (transmission to mice, pigs, and cattle)	[73,75,80,81,82,83]

¹ Excludes mechanical transmission. ² No.

4.2. Other Vertebrate-Infecting Viruses

Blackflies (Simulium buissoni) collected on the Marquesas Islands were tested by PCR for hepatitis B virus (HBV; Hepadnaviridae) [91]. HBV DNA was detected on the external surfaces of blackflies, but not within their internal tissues, indicating that these arthropods are refractory to HBV infection.

4.3. Viruses Not Known to Infect Vertebrates

Multiple DNA viruses were discovered in blackflies (Austrosimulium australense) from New Zealand using viral metagenomics [92]. The majority of the viruses were assigned to the family Microviridae, although viruses from the Circoviridae and Genomoviridae were also detected. None of the identified viruses are known to replicate in vertebrate cells. Two other DNA viruses identified in blackflies are invertebrate iridescent viruses 22 and 25 (Iridoviridae) [93,94]. Both viruses were originally isolated from blackflies in Wales and are assumed to have invertebrate-specific hosts. Because nearly all recognized arboviruses have RNA genomes, it is unlikely that the aforementioned viruses are unrecognized arboviruses.

5. Cimicids

The genus Cimex (family Cimicidae) contains 23 species of wingless ectoparasites that occur worldwide and feed exclusively on the blood of endotherms [95,96,97]. Two cimicid species (Cimex lectularius and Cimex hemipterus) exhibit a preference for human hosts and are commonly referred to as bed bugs. Others prefer bats or birds and are known as bat bugs and bird bugs, respectively. There is no evidence to suggest that cimicids transmit arboviruses to humans, although they are known to harbor many vertebrate-infecting viruses, including five recognized arboviruses: Fort Morgan virus (FMV; Togaviridae), Buggy Creek virus (BCRV; Togaviridae), KKV, Tonate virus (TONV; Togaviridae), and Usutu virus (USUV; Flaviviridae) (Table 4).

5.1. Recognized Arboviruses

Bird bugs, specifically cliff swallow bugs (Oeciacus vicarius), are suspected—but not conclusively confirmed—to be epidemiologically important vectors of FMV [98]. These ectoparasites primarily feed on nesting cliff swallows (Petrochelidon pyrrhonota), but also opportunistically feed on house sparrows (Passer domesticus), which frequently often occupy the nests of cliff swallows [99]. FMV was originally isolated from cliff swallow bugs, cliff swallows, and house sparrows in Colorado, the U.S. [100,101]. In a subsequent study, FMV was isolated from 80 of 1156 (6.9%) nestlings [102]. The prevalence of FMV in cliff swallow bugs is not well documented, although 75 isolates (38 alphaviruses and 37 apparent alphaviruses) were recovered from 5268 cliff swallow bugs (452 pools) collected in Colorado, with an unspecified number later identified as FMV [100,101].

Cliff swallow bugs are susceptible to FMV replication and dissemination and can efficiently transmit the virus to uninfected birds under laboratory conditions [103]. Eighteen days after feeding on a viremic bird, 80% of bugs were infected with FMV. At 24 days post-feeding, salivary glands were removed from 30 bugs and pooled (five pairs of glands in each pool). FMV was detected in five of six pools, and the transmission rate was estimated as 30%. At 47 days post-feeding, 16 bugs (in eight groups of two) were allowed to feed upon uninfected house sparrows (one bird per pair). Four birds became viremic and the transmission rate was estimated as 29%. Additionally, bugs were capable of transmitting FMV at 25, 33, 50, and 83 days post-infectious blood meal. There is no evidence to suggest that any other arthropods are competent vectors of FMV. Despite strong evidence indicating that cliff swallow bugs are likely to be the principal vectors of FMV, they cannot be classified as epidemiologically important vectors without field data demonstrating consistent FMV infection in wild populations.

BCRV (a subtype of FMV) was originally isolated from cliff swallow bugs in Oklahoma, the U.S. [104,105]. Subsequent isolations have been made from cliff swallow bugs, cliff swallows, and house sparrows at various locations across the U.S. [106,107,108,109,110,111]. In one study, BCRV was detected in 1459 of 7421 (19.7%) pools of cliff swallow bugs collected from active cliff swallow nests in Nebraska, the U.S. [106] BCRV causes clinical signs and, in some cases, fatal encephalitis in naturally infected house sparrows, whereas cliff swallows exhibit no apparent adverse effects [112]. Although cliff swallow bugs are known to harbor BCRV, vector competence experiments have not been conducted to determine whether they can transmit the virus to susceptible birds.

Other cimicid-associated arboviruses are KKV, TONV, and USUV (Table 4). KKV was isolated from bat bugs (Stricticimex parvus and Cimex insuetus) in Thailand [113]. Additional isolations have been made from bat flies and bats (discussed earlier in this review) [48,51,54,55,113]. However, KKV failed to replicate in experimentally inoculated bat bugs [114]. TONV has been isolated from cliff swallow bugs, mosquitoes, cliff swallows, house sparrows, and a fringe-lipped bat (Trachops cirrhosus) in the U.S. and South America [115,116,117,118]. TONV is clinically significant because it has been associated with fatal encephalitis and fetal abnormalities in humans [119,120,121]. However, cliff swallow bugs appear to be inefficient biological vectors of TONV. Only 2 of 124 (1.6%) bugs became infected after feeding upon a viremic bird, while 5 of 11 (45.5%) Culex tarsalis mosquitoes became infected [116]. USUV was detected in one of 96 pools of house martin bugs (Oeciacus hirundinis) collected from abandoned house martin (Delichon urbicum) nests in the Czech Republic [122]. Vector competence experiments have not been performed to determine whether bird bugs are capable of biologically transmitting USUV.

5.2. Other Vertebrate-Infecting Viruses

Field-collected C. hemipterus and C. lectularius have tested positive for HBV and hepatitis C virus (HCV; Flaviviridae), but laboratory studies suggest that bed bugs cannot biologically transmit these viruses or human immunodeficiency virus (Retroviridae) [123,124,125,126,127,128,129,130,131,132]. Avian paramyxovirus type 4 (Paramyxoviridae) and hepatitis E virus (Hepeviridae) have also been detected in field-collected bed bugs, while poliovirus (Picornaviridae) persisted for 7 days in C. lectularius that had fed upon viremic monkeys under laboratory conditions, although none of these viruses are known to be transmitted by bed bugs [133,134,135].

Table 4. Arboviruses and other vertebrate-infecting viruses identified in field-collected cimicids.

Virus	Species Name	Abbreviation	Family	Recognized Arbovirus	¹ Experimental Evidence of Efficient Cimicid-Borne Virus Transmission	Citation
Avian paramyxovirus type 4	Paraavulavirus hongkongense	APMV-4	Paramyxoviridae	² -	-	[134]
Buggy Creek virus (a subtype of Fort Morgan virus)	Alphavirus fortmorgan	BCRV	Togaviridae	Yes	-	[104,105,106]
Fort Morgan virus	Alphavirus fortmorgan	FMV	Togaviridae	Yes	Yes (transmission to birds)	[100,101,103]
Hepatitis B virus	Orthohepadnavirus hominoidei	HBV	Hepadnaviridae	-	-	[127,129,132]
Hepatitis C virus	Hepacivirus hominis	HCV	Flaviviridae	-	-	[128]
Hepatitis E virus	Paslahepevirus balayani	HEV	Hepeviridae	-	-	[133]
Kaeng Khoi virus	Orthobunyavirus kaengkhoiense	KKV	Perbunyaviridae	Yes	-	[113]
Tonate virus	Alphavirus tonate	TONV	Togaviridae	Yes	-	[116]
Usutu virus	Orthoflavivirus usutuense	USUV	Flaviviridae	Yes	-	[122]

¹ Excludes mechanical transmission. ² No.

5.3. Viruses Not Known to Infect Vertebrates

Metagenomic surveys of European bat bugs (Cimex pipistrelli) have identified several viruses, but all are phylogenetically distinct from known vertebrate-infecting viruses [136]. Similarly, metagenomics studies of bed bugs have revealed numerous viruses, but with the exception of HCV, all are phylogenetically distinct from viruses known to replicate in vertebrates [128,136,137,138,139]. A Sedoreoviridae-like virus was also detected in field-collected bed bugs using electron microscopy, but there is no evidence indicating its ability to replicate in vertebrate cells [140].

6. Culicoides Midges

Hematophagous midges (Ceratopogonidae) have a worldwide geographic distribution and are well-known nuisance biters, often causing significant discomfort in humans and other vertebrates. The family contains more than 5400 species across 125 genera [141,142]. Many species in the genus Culicoides are vectors of viruses, protozoans, and bacteria [143,144,145]. Some Culicoides-associated viruses are arboviruses of medical and veterinary significance, and notable examples include African horse sickness virus (AHSV; Sedoreoviridae), Akabane virus (AKAV; Peribunyaviridae), bluetongue virus (BTV; Sedoreoviridae), bovine ephemeral fever virus (BEFV; Rhabdoviridae), epizootic hemorrhagic disease virus (EHDV; Sedoreoviridae), Main Drain virus (MDV; Peribunyaviridae), Oropouche virus (OROV; Peribunyaviridae), and Schmallenberg virus (SBV; Peribunyaviridae) [146,147,148,149,150,151,152,153,154,155,156,157] (Table 5).

6.1. Recognized Arboviruses

AHSV is the etiological agent of African horse sickness, a highly infectious and often fatal disease of equids endemic to sub-Saharan Africa, but also present in parts of Europe and Asia [146,148,151]. AHSV exists as nine antigenic types, designated as AHSV-1 to AHSV-9. Culicoides imicola is likely the principal vector of AHSV due to its abundance in endemic regions, its history of yielding more AHSV isolations than any other arthropod species, and its susceptibility to infection under laboratory conditions [54,158,159,160,161,162]. However, other Culicoides species, including Culicoides bolitinos, have also been implicated as vectors [161,163]. Field studies have reported the detection of AHSV in Culicoides midges [54,158,159,160,161]. For example, AHSV was detected in 12 of 95 pools of Culicoides midges from South Africa [160]. Of the AHSV-positive pools, seven pools consisted exclusively of C. imicola, four pools contained a mixture of C. imicola and other species, and one pool contained a mixture of species, excluding C. imicola. In a study performed in Spain, AHSV was isolated from six of 31 pools of Culicoides midges, including four pools of C. imicola [158]. A six-year surveillance study performed in South Africa reported the isolation of AHSV from 66 of 4506 pools of Culicoides midges [159]. All isolations were made from C. imicola.

Several studies have investigated the vector competence of Culicoides midges for AHSV [162,164,165]. Wetzel and colleagues were unable to demonstrate transmission of AHSV by Culicoides midges to horses [165]. The experiments were performed using field-collected Culicoides midges of multiple species, but most were C. imicola (aka C. pallidipennis). The same article refers to an unpublished study, cited via a personal communication, in which field-collected Culicoides midges (species not specified) were allowed to feed upon a horse infected with AHSV. Twelve days later, the midges were permitted to feed upon an uninfected horse, which became ill and died of African horse sickness 12 days later. In another study, Culicoides variipennis that had fed upon AHSV-infected, embryonated chicken eggs were able to transmit the virus to uninfected eggs [164]. In a more recent study, 17 species of Culicoides midges from South Africa were orally challenged with AHSV-3, AHSV-5, and AHSV-8, then tested for virus after a 10-day extrinsic incubation period [162]. AHSV-5 was detected in 8.5% of C. imicola and 20.6% of C. bolitinos but was not detected in any other species. AHSV-8 was detected in 26.8% of C. imicola and 1.7% of C. bolitinos, while all other species tested negative. ASHV-3 was found in 3.8% of C. bolitinos, but was not detected in any other species, including C. imicola.

AKAV is a teratogenic pathogen affecting cattle, sheep, and goats across Asia, Africa, Australia, and the Middle East [145,147]. In Australia, Culicoides brevitarsis is suspected to be the primary arthropod vector [166,167,168,169,170,171,172]. AKAV was isolated from 3 of 101 pools containing 5314 C. brevitarsis temporally and spatially associated with sentinel cattle, from which isolations were made in Queensland [166]. Laboratory studies have shown that AKAV disseminates to the salivary glands of C. brevitarsis, but transmission experiments were not performed [172]. In Japan, Culicoides oxystoma is implicated as the major arthropod vector [173,174,175,176]. In one study, AKAV was isolated from 3 of 84 pools of Culicoides spp. collected inside a cowshed where calves had seroconverted [174]. All positive pools contained C. oxystoma. AKAV has also been isolated from Culicoides midges in China, although the species were not identified [177]. The vector competence of C. oxystoma and five other Culicoides spp. from Japan has been evaluated [176]. All species were susceptible to oral infection, but viral titers were significantly higher in C. oxystoma. In Africa and the Middle East, C. imicola is suspected to be the primary vector of AKAV, although vector competence experiments have not been performed [178,179,180,181]. Vector competence has been assessed for Culicoides nubeculosus and Culicoides variipennis, which do not occur in the endemic range of AKAV [182]. Following intrathoracic inoculation, the virus persisted for at least nine days in both species, but only C. variipennis was susceptible to oral infection and successfully transmitted the virus through a membrane at 7–10 days post-infection.

BTV is the etiological agent of bluetongue disease, a non-contagious disease affecting domestic and wild ruminants in Africa, the Middle East, Asia, Australia, the Americas, and Europe [149,183,184]. There are at least 36 recognized serotypes of BTV, designated as BTV-1 to -36 [185]. Most serotypes are primarily transmitted by Culicoides spp. midges, but some are transmitted horizontally by direct contact between vertebrate hosts. In North America, Culicoides sonorensis is considered the primary arthropod vector of BTV, although other species, i.e., Culicoides insignis and Culicoides stellifer, may also contribute to transmission [152]. In the Caribbean and South America, C. insignis is recognized as an important vector [186,187]. In Australia, C. brevitarsis is regarded as the principal vector, although several less-widespread species may have regionally important roles [188]. In Africa, the primary vector is C. imicola, although C. bolitinos and select other species may also be involved in transmission [189]. C. imicola is also a major vector in southern Europe and the Mediterranean region, while Culicoides obsoletus and Culicoides pulicaris are important in northern Europe [190]. In Asia, the major vectors of BTV are not well defined, but several Culicoides species, including Culicoides actoni, Culicoides fulvus, and Culicoides wadai, have been implicated [185,191,192].

Many field studies have reported the presence of BTV in Culicoides midges, and the prevalence has often been high [192,193,194,195,196,197,198,199,200]. In Italy, BTV RNA was detected in 68 of 77 pools comprising 1591 Culicoides spp. midges [196]. The prevalence of BTV was also high in C. imicola, with 52 of 53 pools comprising 1305 individuals testing positive. Similarly, in Morocco, BTV RNA was found in 45 of 55 pools containing 2003 Culicoides spp. midges [193]. C. imicola again showed a notably high prevalence with 15 of 16 pools testing positive. In Kazakhstan, BTV RNA was detected in 16 of 79 pools of midges [194].

Multiple studies have investigated the vector competence of Culicoides midges for BTV, revealing varying results depending on the midges spp. and BTV serotype that were used [152,196,201,202,203,204,205,206,207,208,209,210,211,212,213,214,215,216]. C. sonorensis from the United States could transmit a live-attenuated vaccine strain of BTV-1 from vaccinated to unvaccinated sheep [213]. Another study demonstrated that a single infected C. sonorensis could transmit BTV-1 to naïve sheep with an efficiency of 80–100% [214]. C. nubeculosus and C. impunctatus from England supported the replication of BTV-3 and BTV-4 after feeding upon viremic sheep [215]. BTV-1 was detected in the saliva of 53% of C. brevitarsis from Australia experimentally inoculated with the virus and viral transmission to a naïve sheep was documented [216]. In another study, 22.7 to 82.0% of C. bolitinos and 1.9 to 9.8% C. imicola from South Africa were susceptible to infection by BTV-1, -3, and -4 after oral challenge [217]. Dissemination rates as high as 57.6%, 56.0%, and 50.5% were reported for C. imicola from Italy after oral challenge with BTV-2, -4, and -8, respectively [202].

BEFV is the etiological agent of bovine ephemeral fever, an acute febrile illness that primarily affects cattle and water buffalo across Africa, Asia, Australia, and the Middle East [150,218]. Evidence suggests that BEFV is predominantly transmitted by mosquitoes and Culicoides midges, although evidence of Culicoides-borne viral transmission under laboratory conditions is lacking. BEFV has been isolated once from C. brevitarsis in Australia, once each from C. imicola and Culicoides coarctatus in Zimbabwe, and once from a pool containing several different species of Culicoides midges in Kenya [180,219,220]. BEFV has also been detected at least once in C. puncticollis in Turkmenistan, and BEFV RNA was detected by RT-PCR in one of 166 pools of Culicoides spp. midges in Korea [221,222]. However, BEFV has been isolated more often from mosquitoes. In one study, BEFV was isolated from 10 pools of Aedes, Culex, and Uranotaenia spp. mosquitoes in Australia [223]. Three more isolations were made from Anopheles bancroftii in Australia [219,224].

Several studies have investigated whether Culicoides midges are competent vectors of BEFV [225,226,227,228]. Colonized C. sonorensis and mosquitoes (Aedes aegypti, Culex pipiens, and Culex quinquefasciatus) were challenged with BEFV by artificial blood meal and intrathoracic inoculation [225]. After oral exposure, infection and dissemination rates in C. sonorensis were 7.6% and 1.2%, respectively, while no evidence of viral replication was observed in any mosquitoes. BEFV replication occurred in all C. sonorensis challenged by intrathoracic inoculation, although the midges failed to transmit the virus to naïve calves. In an earlier study, BEFV was isolated from one of 526 (0.2%) C. brevitarsis and an unspecified number of Culicoides marksi 10 days after oral challenge [226]. BEFV was also isolated from 3 of 23 (13.0%) Culex annulirostris that had been orally challenged with the virus 12 days earlier.

EHDV is the etiological agent of epizootic hemorrhagic disease, a significant cause of morbidity and mortality among livestock and wildlife across North and South America, Africa, Asia, Australia, and the Middle East [157,185,229]. The virus occurs as seven serotypes, designated as EHDV-1 to -7. The Culicoides spp. implicated in the transmission of EHDV include C. imicola, C. sonorensis, C. obsoletus, C. brevitarsis, C. insignis, C. mohave, C. oxystoma, and C. variipennis. All of these species have tested positive for EHDV in surveillance studies [219,230,231,232,233,234]. For example, EHDV was detected by RT-PCR in 6 of 103 pools containing 964 Culicoides midges collected during an EHDV outbreak at the Minnesota Zoo in the U.S., with half the positive pools comprising C. sonorensis [230]. In Italy, 18 of 411 pools containing 5721 Culicoides midges contained EHDV RNA, including 10 pools of C. imicola [232].

Vector competence experiments have shed light on the role of Culicoides midges in EHDV transmission [152,235,236,237,238]. In one study, C. sonorensis midges were orally exposed two strains of EHDV-2, resulting in infection, dissemination, and transmission rates as high as 100%, 90%, and 79%, respectively, for both strains at 9 to 14 days after exposure [235]. Transmission rates were calculated by dividing total midges with detectable virus in saliva by total midges with disseminated infection. Additional studies have investigated the ability of Culicoides midges to experimentally transmit EHDV to vertebrate animals [236,238,239]. In one such experiment, C. sonorensis were allowed to feed on EHDV-2 inoculated deer and then, after an extrinsic incubation period of 14 to 16 days, permitted to feed on a naïve deer, which subsequently developed viremia and severe disease [240]. Biological transmission of EHDV-1 from one deer to another by C. variipennis also has been documented [239]. In another study, C. sonorensis became infected after feeding on embryonated chicken eggs inoculated with EHDV-1, -2, and -6; however, none successfully transmitted the virus to uninfected eggs [238].

MDV occurs in the western U.S. and is not a common cause of disease, although it has been isolated from the brain of a horse that died of encephalitis [156]. Most arthropod-derived isolations of MDV have been made from Culicoides variipennis (on at least 31 occasions) and undetermined Culicoides spp. (on at least 65 occasions) [241]. Experimental studies have shown that MDV replicates in both C. variipennis and C. nubeculosus after intrathoracic inoculation and oral ingestion [242]. Moreover, both species were capable of transmitting the virus to suspensions of uninfected blood suspensions 16 days after feeding upon infectious blood.

OROV occurs in Latin America and the Caribbean and is the etiological agent of Oropouche fever, the only known Culicoides-transmitted disease of humans [153,154,155,243,244]. Culicoides paraeneses is considered the primary urban vector because it is often highly abundant in areas where human outbreaks occur and the virus has been isolated from this species [245,246,247,248,249,250]. For instance, over 95% of Culicoides collected during an outbreak in Brazil in 1975 were identified as C. paraeneses, with two isolations of OROV made from the approximately 15,000 Culicoides inoculated into mice [246]. C. paraensis is a competent vector of OROV in the laboratory [251,252,253]. C. paraensis that had engorged upon viremic hamsters were able to transmit OROV to naïve hamsters after a 4 to 9 day extrinsic incubation period [252]. Viral transmission rates were calculated as 25% to 83%. In another study, OROV was experimentally transmitted from a viremic human to a hamster by C. paraensis after a 6 to 12 day extrinsic incubation period [251]. C. sonorensis is also a competent laboratory vector of OROV [254]. In this study, three species of arthropods (C. sonorensis, Cx. quinquefasciatus, and Cx. tarsalis) from the United States were provided OROV-spiked blood meals. At 14 days post-engorgement, C. sonorensis demonstrated infection, dissemination, and transmission rates of 87%, 83%, and 20%, while both mosquito species demonstrated transmission rates of <1%. Other studies have also shown that mosquitoes are incompetent vectors of OROV [255,256,257,258].

SBV is a cause of pregnancy loss and congenital malformations in domestic ruminants in Europe [259,260,261,262]. Culicoides midges, particularly those in the Obsoletus complex, appear to be major vectors. The Obsoletus complex contains as many as five species: Culicoides chiopterus, Culicoides dewulfi, Culicoides montanus, Culicoides obsoletus, and Culicoides scoticus [263,264,265]. SBV has been detected in multiple species of Culicoides midges [199,266,267,268,269,270,271,272,273,274,275,276,277]. In Belgium, SBV RNA was detected in 12 of 178 pools comprising 1976 Culicoides midges [267]. The positive pools consisted of C. obsoletus complex, C. obsoletus, C. dewulfi, Culicoides chiopterus, and Culicoides pulicaris. In France and Italy, SBV RNA was detected in 22 of 508 pools (5135 midges) and 3 of 759 pools (28,177 midges), respectively [266]. Most positive pools consisted of C. obsoletus complex and C. obsoletus. In the Netherlands, SBV RNA was detected in 12 of 230 pools of C. obsoletus complex [270].

Several studies have investigated whether Culicoides midges are susceptible to SBV infection [266,275,278,279]. SBV was detected in 8 of 147 (7.3%) C. obsoletus complex midges after an infectious artificial blood meal administered 8 days earlier [278]. In another study, colonized C. nubeculosus challenged via artificial blood meals were refractory to SBV infection [275]. In experiments performed with field-collected midges, SBV RNA was detected in the heads of 26 of 32 (81%) individual C. imicola following oral challenge 10 days earlier. However, experimental transmission of SBV to naïve animals via the bite of infected Culicoides midges has not been demonstrated.

Other arboviruses have been detected in Culicoides spp. midges (Table 5). Among them are many viruses belonging to the Peribunyaviridae, although most have been isolated from Culicoides on only a few occasions and none have been efficiently transmitted by midges under experimental conditions [54,145,175,224,280,281,282,283,284,285,286,287,288]. Other arboviruses have occasionally been identified in Culicoides and this includes viruses belonging to Sedoreoviridae and Rhabdoviridae, in addition to Barmah Forest (Togaviridae), EEEV, CCHFV, Israel turkey meningoencephalitis virus (Flaviviridae), RVFV, VEEV, VSNJV, and WNV [54,175,224,289,290,291,292,293,294,295,296,297,298,299,300,301,302,303,304,305,306,307,308,309]. The ability of Culicoides midges to support the replication and dissemination of some of these viruses has been assessed, revealing that they are predominantly refractory to infection [310,311,312].

6.2. Other Vertebrate-Infecting Viruses

Fowlpox virus (Poxviridae) was detected in Culicoides arakawae in South Korea [313]. C. arakawae transmitted the virus to naive chickens after a 7-day extrinsic incubation period [314]. However, the transmission of fowlpox virus by arthropods is assumed to be mechanical [315].

6.3. Viruses Not Known to Infect Vertebrates

Many novel viruses have been detected in Culicoides spp. midges using metagenomics [316,317,318,319,320,321,322,323,324,325,326]. In most of these studies, the abilities of the newly discovered viruses to replicate in vertebrate cell lines were not assessed. When virus isolation in vertebrate cell culture was attempted, no isolates were recovered.

Table 5. Arboviruses and other vertebrate-infecting viruses identified in field-collected Culicoides midges ¹.

Virus	Species Name	Abbreviation	Family	Recognized Arbovirus	² Experimental Evidence of Culicoides-Borne Virus Transmission	Citation
Adelaide River virus	Ephemerovirus adelaide	ARV	Rhabdoviridae	Yes	³ -	[54]
African horse sickness virus	Orbivirus alphaequi	AHSV	Sedoreoviridae	Yes	Yes (transmission to embryonated chicken and anecdotal evidence of transmission to a horse)	[54,158,159,160,161,164,165]
Aino virus	Orthobunyavirus ainoense	AINOV	Peribunyaviridae	Yes	-	[175]
Akabane virus	Orthobunyavirus akabaneense	AKAV	Peribunyaviridae	Yes	Yes (transmission through a membrane)	[175]
Banna virus	Seadornavirus bannaense	BAV	Sedoreoviridae	Yes	-	[302]
Barmah Forest virus	Alphavirus barmah	BFV	Togaviridae	Yes	-	[224]
Beatrice Hill virus	Tibrovirus beatrice	BHV	Rhabdoviridae	Yes	-	[224]
Belmont virus	Orthobunyavirus belmontense	BELV	Peribunyaviridae	Yes	-	[224]
Bivens arm virus (a subtype of Tibrogargan virus)	Tibrovirus tibrogargan	BAV	Rhabdoviridae	Yes	-	[300]
Bluetongue virus	Orbivirus caerulinguae	BTV	Sedoreoviridae	Yes	Yes (transmission to sheep)	[192,193,194,195,196,197,198,199,200,213,214]
Bovine ephemeral fever virus	Ephemerovirus febris	BEFV	Rhabdoviridae	Yes	-	[180,219,220]
Bunyip Creek virus	none	BCV	Sedoreoviridae	Yes	-	[303]
Buttonwillow virus	Orthobunyavirus buttonwillowense	BUTV	Peribunyaviridae	Yes	-	[285,296]
Chuzan virus	⁴ none	CHUV	Sedoreoviridae	-	-	[298]
Crimean-Congo hemorrhagic fever virus	Orthonairovirus haemorrhagiae	CCHFV	Nairoviridae	Yes	-	[290]
Curionopolis virus	Curiovirus curionopolis	CURV	Rhabdoviridae	Yes	-	[301]
Douglas virus (a subtype of Schmallenberg virus)	Orthobunyavirus schmallenbergense	DOUV	Peribunyaviridae	Yes	-	[280]
Eastern equine encephalitis virus	Alphavirus eastern	EEEV	Togaviridae	Yes	-	[291]
Epizootic hemorrhagic disease virus	Orbivirus ruminantium	EHDV	Sedoreoviridae	Yes	Yes (transmission to deer)	[231,239,327,328,329]
Eubenangee virus	Orbivirus eubenangeense	EUBV	Sedoreoviridae	Yes	-	[224]
Facey’s Paddock virus	Faceyspaddock orthobunyavirus	FPV	Peribunyaviridae	Yes	-	[224]
Fowlpox virus	Avipoxvirus fowlpox	FWPV	Poxviridae	-	-	[314]
Fukuoka virus	Ledantevirus fukuoka	FUKV	Rhabdoviridae	Yes	-	[308]
Ibaraki virus	none	IBAV	Sedoreoviridae	Yes	-	[175]
Ingwavuma virus	Orthobunyavirus ingwavumaense	INGV	Peribunyaviridae	Yes	-	[287]
Iquitos virus (a subtype of Oropouche virus)	Orthobunyavirus oropoucheense	IQTV	Peribunyaviridae	Yes	-	[145]
Israel turkey meningoencephalitis virus	Orthoflavivirus israelense	ITV	Flaviviridae	Yes	-	[54]
Itacaiunas virus	Curiovirus itacaiunas	ITAV	Rhabdoviridae	Yes	-	[301]
Jatobal virus	Orthobunyavirus jatobalense	JATV	Peribunyaviridae	Yes	-	[145]
Kimberley virus	Ephemerovirus kimberley	KIMV	Rhabdoviridae	Yes	-	[305]
Kotonkan virus	Ephemerovirus kotonkan	KOTV	Rhabdoviridae	Yes	-	[304]
Leanyer virus	Orthobunyavirus leanyerense		Peribunyaviridae	Yes	-	[224]
Lokern virus (a subtype of Main Drain virus)	Orthobunyavirus kernense	LOKV	Peribunyaviridae	Yes	-	[296]
Madre de Dios virus (a subtype of Oropouche virus)	Orthobunyavirus oropoucheense	MDDV	Peribunyaviridae	Yes	-	[145]
Main Drain virus	Orthobunyavirus kernense	MDV	Peribunyaviridae	Yes	Yes (transmission to uninfected blood suspensions)	[296]
Manzanilla virus	Orthobunyavirus manzanillaense	MANV	Peribunyaviridae	Yes	-	[145]
Ngaingan virus	Hapavirus ngaingan	NGAV	Rhabdoviridae	Yes	-	[309]
Oropouche virus	Orthobunyavirus oropoucheense	OROV	Peribunyaviridae	Yes	Yes (transmission to hamsters)	[246,251]
Palyam virus	Orbivirus palyamense	PALV	Sedoreoviridae	Yes	-	[299]
Peaton virus	Orthobunyavirus peachesterense	PEAV	Peribunyaviridae	Yes	-	[281,282]
Pintupo virus (a subtype of Oropouche virus)	Orthobunyavirus oropoucheense	PINTV	Peribunyaviridae	-	-	[251]
Rift Valley fever virus	Phlebovirus riftense	RVFV	Phenuiviridae	Yes	-	[289]
Sabo virus	Orthobunyavirus saboense	SABOV	Peribunyaviridae	Yes	-	[290]
Sango virus	Orthobunyavirus sangoense	SANV	Peribunyaviridae	Yes	-	[290]
Sathuperi virus (a subtype of Schmallenberg virus)	Orthobunyavirus schmallenbergense	SATV	Peribunyaviridae	Yes	-	[290]
Schmallenberg virus	Orthobunyavirus schmallenbergense	SBV	Peribunyaviridae	Yes	-	[199,266,267,268,269,270,271,272,273,274,275,276,277]
Shamonda virus (a subtype of Schmallenberg virus)	Orthobunyavirus schmallenbergense	SHAV	Peribunyaviridae	Yes	-	[290]
Shuni virus	Orthobunyavirus shuniense	SHUV	Peribunyaviridae	Yes	-	[290]
Simbu virus	Orthobunyavirus simbuense	SIMV	Peribunyaviridae	Yes	-	[145]
Sweetwater Branch virus	Tibrovirus sweetwater	SWBV	Rhabdoviridae	Yes	-	[307]
Thimiri virus	Orthobunyavirus thimiriense	THIV	Peribunyaviridae	Yes	-	[224]
Tibrogargan virus	Tibrovirus tibrogargan	TIBV	Rhabdoviridae	Yes	-	[306]
Tinaroo virus (a subtype of Akabane virus)	Orthobunyavirus akabaneense	TINV	Peribunyaviridae	Yes	-	[280]
Venezuelan equine encephalitis virus	Alphavirus venezuelan	VEEV	Togaviridae	Yes	-	[297]
Vesicular stomatitis New Jersey virus	Vesiculovirus newjersey	VSNJV	Rhabdoviridae	Yes	-	[296]
Wallal virus	Orbivirus betamitchellense	WALV	Sedoreoviridae	Yes	-	[224]
Warrego virus	Orbivirus gammamitchellense	WARV	Sedoreoviridae	Yes	-	[224]
Weldona virus (a subtype of Tete virus)	Orthobunyavirus teteense	WELV	Peribunyaviridae	Yes	-	[288]
West Nile virus	Orthoflavivirus nilense	WNV	Flaviviridae	Yes	-	[292]
Wongorr virus	Orbivirus deltamitchellense	WGRV	Sedoreoviridae	Yes	-	[224]

¹ Not an exhaustive list. ² Excludes mechanical transmission. ³ No. ⁴ Not recognized by the International Committee on Taxonomy of Viruses.

7. Fleas

Fleas (Siphonaptera) are small, wingless insects that feed on the blood of endothermic vertebrates, most commonly rodents. Over 2500 species of fleas across 19 families have been identified [330,331]. Fleas occur on every continent, except Antarctica, and are vectors of several bacterial pathogens of medical significance, most notably, Yersinia pestis, the causative agent of plague. These insects are not considered major vectors of arboviruses, although a minor role in the transmission of TBEV has been suggested (Table 6).

7.1. Recognized Arboviruses

TBEV has been isolated from several species of fleas associated with small wild mammals, including Ctenocephalides spp. fleas infesting European moles (Talpa europaea) in Slovakia [332,333]. Nonetheless, such isolations are rare compared to the numerous isolations made from ticks [334,335]. Laboratory studies have demonstrated that fleas are capable of transmitting TBEV under experimental conditions [336,337]. For example, mouse fleas (Leptopsylla segnis) that fed on viremic mice were able to transmit TBEV to uninfected mice at two and three days post-feeding [336]. TBEV persisted in some fleas for up to four days. In another study, TBEV persisted in northern rat fleas (Nosopsyllus fasciatus) for 24 h and oriental rat fleas (Xenopsylla cheopis) for six days [338]. A key limitation of these studies is that viral titers were not quantified, leaving uncertainty as to whether active viral replication occurred in the fleas.

7.2. Other Vertebrate-Infecting Viruses

Hantaan virus (HTNV; Hantaviridae) has been isolated from fleas (Leptopsylla segnis and Monopsyllus anisus) collected from the nests of HTNV-infected striped field mice (Apodemus agrarius) in China [339]. In experimental studies, HTNV persisted for no more than two days in L. segnis and M. anisus following ingestion of virus-spiked blood. These findings suggest that both species of fleas are refractory to HTNV infection. Another virus that has been detected in field-collected fleas is canine distemper virus (CDV; Paramyxoviridae), with the detections made by RT-PCR in fleas (Ceratophyllus sciurorum) collected from American mink (Neovison vison) during an outbreak of CDV on a mink farm in Denmark [340]. However, vector competence experiments have not been performed to determine whether fleas are capable of biologically transmitting CDV. Like CDV, feline leukemia virus (FeLV; Retroviridae) is a pathogen of veterinary significance [341,342]. Transmission experiments revealed that cat fleas (Ct. felis) that fed upon FeLV-infected blood could transmit the virus to uninfected blood [343]. However, the uninfected blood was offered immediately after the infectious blood meal, indicating that the transmission was mechanical. Notably, FeLV has never been isolated from naturally infected fleas. Fleas are also capable of transmitting feline calicivirus (FCV; Caliciviridae), a common pathogen of domestic cats [344]. FCV was isolated from cat fleas (Ctenocephalides felis) four days after they had engorged upon FCV-spiked blood. One of four kittens seroconverted after being fed upon by fleas that had engorged upon FCV-spiked blood. However, it is unclear whether biological transmission occurred because the length of time between blood meals was not specified. If the fleas fed upon the kittens shortly after their first blood meal, mechanical transmission most likely occurred. Isolations of FCV have never been made from naturally infected fleas.

Table 6. Arboviruses and vertebrate-infecting viruses identified in field-collected fleas.

Virus	Species Name	Abbreviation	Family	Recognized Arbovirus	¹ Experimental Evidence of Flea-Borne Virus Transmission	Citation
Canine distemper virus	Morbillivirus canis	CDV	Paramyxoviridae	² -	-	[340]
Hantaan virus	Orthohantavirus hantanense	HTNV	Hantaviridae	-	-	[339]
Tick-borne encephalitis virus	Orthoflavivirus encephalitidis	TBEV	Flaviviridae	Yes	Yes (transmission to mice, but it may have been mechanical)	[332,333,336,337]

¹ Excludes mechanical transmission. ² No.

The potential for fleas to support the replication of Marburg virus (MARV; Filoviridae) and Myxoma virus (Poxviridae) has been investigated, although neither virus has been detected in field-collected fleas [345,346]. MARV RNA was detected by quantitative RT-PCR in bat fleas (Thaumapsylla breviceps) that had fed upon viremic Egyptian fruit bats (Rousettus aegyptiacus), but the levels of viral RNA did not exceed the estimated amounts in the ingested blood, suggesting a lack of virus replication [345]. Furthermore, MARV-inoculated fleas failed to transmit the virus to uninfected bats. In a separate study, European rabbit fleas (Sphilopsyllus cuniculi) that had been immersed in a solution containing Myxoma virus successfully transmitted the virus to European rabbits (Oryctolagus cuniculus), but there is no evidence to indicate that transmission was biological [346].

7.3. Viruses Not Known to Infect Vertebrates

Eighteen novel viruses were detected by metagenomic analysis in fleas from three genera (Stephanocircus, Pygiopsylla, and Macropsylla) collected in Australia; however, none were closely related to viruses known to infect vertebrates [347]. In another metagenomic study, novel viruses from at least eight families were detected in fleas infesting bats (Myotis daubentonii, Rhinolophus spp., and Rousettus leschenaultia) in China, but none of the newly discovered viruses are known to infect vertebrates [63].

8. Hippoboscid Flies

Hippoboscid flies (Hippoboscidae), commonly referred to as louse flies and keds, are obligatory blood-feeders of birds and mammals [348,349]. At least 213 species of hippoboscid flies have been identified. These insects have a worldwide distribution but are most common in the tropics and subtropics. Hippoboscid flies have been implicated in the transmission of various bacterial and protozoan pathogens but are not regarded as significant vectors of viruses [348,350,351]. However, two recognized arboviruses and three other vertebrate-infecting viruses have been detected in these insects (Table 7).

8.1. Recognized Arboviruses

Two recognized arboviruses, BTV and WNV, have been detected in hippoboscid flies [352,353,354]. BTV was detected by RT-PCR in Hippobosca equina in Kazakhstan and WNV was detected by RT-PCR in Icosta americana in New Jersey, the U.S and Ontario, Canada. BTV has been experimentally transmitted to sheep by hippoboscid flies (Melophagus ovinus), but the mode of transmission was most likely mechanical [355]. There is no evidence to suggest that either BTV or WNV is biologically transmitted by hippoboscid flies.

8.2. Other Vertebrate-Infecting Viruses

Three additional vertebrate-infecting viruses have been detected in hippoboscid flies: Aksy-Durug Melophagus sigmavirus (Rhabdoviridae), border disease virus (Flaviviridae), and bovine viral diarrhea virus (Flaviviridae) [354,356,357]. Aksy-Durug Melophagus sigmavirus was detected in M. ovinus in Russia using viral metagenomics and shown to replicate in pig embryo kidney (PEK) cells, although it has not been isolated from any naturally infected vertebrates [357]. Border disease virus was identified in M. ovinus in China, and bovine viral diarrhea virus was identified in H. equina in Kazakhstan, but there is no evidence to suggest that hippoboscid flies are biological vectors of either virus [354,356].

Table 7. Arboviruses and other vertebrate-infecting viruses identified in field-collected hippoboscid flies.

Virus	Species Name	Abbreviation	Family	Recognized Arbovirus	¹ Experimental Evidence of Hippoboscid-Borne Virus Transmission	Citation
Aksy-Durug Melophagus sigmavirus	² none	ADMSV	Rhabdoviridae	³ -	-	[357]
Bluetongue virus	Orbivirus caerulinguae	BTV	Sedoreoviridae	Yes	-	[354]
Border disease virus	Pestivirus ovis	BDV	Flaviviridae	-	-	[356]
Bovine viral diarrhea virus	Pestivirus bovis	BVDV	Flaviviridae	-	-	[354]
West Nile virus	Orthoflavivirus nilense	WNV	Flaviviridae	Yes	-	[352,353]

¹ Excludes mechanical transmission. ² Not recognized by the International Committee on Taxonomy of Viruses. ³ No.

8.3. Viruses Not Known to Infect Vertebrates

Multiple viruses with apparent arthropod-specific host ranges have been detected in hippoboscid flies [357,358]. For example, a metagenomic analysis of M. ovinus in Russia resulted in the identification of four viruses from three families (Iflaviridae, Reoviridae, and Solemoviridae), none of which could replicate in vertebrate cells [357].

9. Lice

Lice are wingless ectoparasites belonging to multiple families within the order Psocodea, which comprises approximately 2500 species. Lice occur worldwide and are often classified into two major groups: sucking lice (superfamily Anoplura) and chewing lice (formerly the order Mallophaga, now but divided into three suborders: Amblycera, Ischnocera, and Rhynchophthirina) [359,360]. All sucking lice are obligate blood-feeders that parasitize mammals. Chewing lice (also known as biting lice) primarily parasitize birds, but also mammals, feeding on feathers, skin, fur, and scales. Although lice are not considered biological vectors of arboviruses, they transmit various bacterial pathogens of medical significance [361,362].

9.1. Recognized Arboviruses

No recognized arboviruses have been detected in naturally infected lice. However, KFDV was isolated from suckling lice (Hoplopleura maniculata and Neohaematopinus echinatus) that had fed on KFDV-inoculated jungle palm squirrels (Funambulus tristriatus) in the laboratory [363].

9.2. Other Vertebrate-Infecting Viruses

Southern elephant seal virus (SESV; Togaviridae) was originally isolated from hematophagous lice (Lepidophthirus macrorhini) infesting southern elephant seals (Mirounga leonina) in Australia [364]. Antibodies to SESV were detected in seals from which the lice were collected and the virus replicates in BHK-21 and Vero cells. Nonetheless, there is no evidence to suggest that lice are biological vectors of SESV. Similarly, there is also no evidence indicating that lice are biological vectors of poliovirus, as the virus was not recovered from sucking lice (Pediculus capitis and Pediculus vestimenti) that had fed upon viremic monkeys [135].

9.3. Viruses Not Known to Infect Vertebrates

Relatively little research has been conducted to identify and characterize the virome of lice [365,366,367,368,369]. Some studies have examined blood-feeding lice, while others investigated non-blood-feeding species. Most of the viruses identified to date are only distantly related to known vertebrate-infecting viruses

10. Mites

Mites are small arachnids belonging to two superorders (Acariformes and Parasitiformes) [370,371,372]. More than 48,200 species of mites have been identified, and they are globally distributed, occurring in almost every type of environment. Some mites feed upon the blood, skin, and keratin of vertebrates, while others feed on insects or plants. Mites are vectors of bacterial pathogens of medical significance, including the etiological agents of rickettsialpox and scrub typhus [373]. More pertinent to this review, mites harbor a diverse range of viruses (Table 8).

10.1. Recognized Arboviruses

Studies performed in the 1940s provide evidence that mites are biological vectors of St. Louis encephalitis virus (SLEV; Flaviviridae), but subsequent work could not verify these findings and Culex spp. mosquitoes are now considered the principal vectors of the virus. SLEV was isolated from naturally infected chicken mites (Dermanyssus gallinae) in Missouri, the U.S. and persisted for five months in naturally infected mites transferred to the laboratory for propagation [374,375]. D. gallinae became infected with SLEV after feeding upon viremic chickens and were able to transmit the virus to naïve chickens [376,377]. Naturally infected mites also transmitted the virus to naïve chickens. However, a later study reported that SLEV persists for only 2 h in D. gallinae [378]. Ornithonyssus sylviarum and Ornithonyssus bursa bird mites were also evaluated, with SLEV persisting for only 2 and 36 h, respectively. SLEV was not transmitted to naïve chickens by D. gallinae, O. bursa, and O. sylviarum that had fed upon viremic chickens one to three days earlier. Mites are now considered to have an inconsequential role in the maintenance of SLEV in nature because of the conflicting data produced in the transmission experiments, sparse number of SLEV-infected fleas collected in the field, and abundance of data revealing that Culex spp. mosquitoes are the principal vectors.

EEEV and Western equine encephalitis virus (WEEV; Togaviridae) have been isolated from mites in the U.S. [379,380,381,382,383]. Isolations of EEEV were made from chicken mites (D. gallinae) in Tennessee and isolations of WEEV were made from American bird mites (Dermanyssus americanus) in Colorado, chicken mites (D. gallinae) and tropical fowl mites (Liponyssus bursa) in Texas, and tropical rat mites (Ornithonyssus sylviarum; aka Liponyssus sylviarum) in California. However, very few isolations of EEEV and WEEV have been made from mites compared to mosquitoes [291,384,385].

Vector competence experiments have been performed to determine whether mites can transmit EEEV and WEEV [386,387]. In one study, mites (D. gallinae, L. bursa, and L. sylviarum) were allowed to feed upon chickens infected with EEEV or WEEV [386]. In most cases, virus persisted in the mites no longer than two days, but there were exceptions, i.e., EEEV was isolated from a pool of D. gallinae at 15 days post-engorgement. In large-scale transmission experiments where several thousand mites that had fed upon viremic chickens were allowed to feed upon 125 naive chickens, there were only two instances of transmission; one where EEEV was transmitted to a chicken by D. gallinae after a 26-day extrinsic incubation and another where WEEV was transmitted to a chicken by D. gallinae after a 13-day extrinsic incubation. The authors concluded that mites are not major vectors of EEEV or WEEV. In another study, D. gallinae fed upon chicks that had been inoculated with EEEV [387]. EEEV persisted in the mites for 30 days, but viral titers did not increase with time, indicating that virus replication had not occurred in the mites. Mites that had fed on viremic chickens were allowed to feed on naive chickens at 3, 7, 11, 15, or 30 days post-engorgement. A subset of mites that engorged 3 and 7 days earlier could transmit EEEV, but the other mites did not. Although transmission was documented, the apparent inability of EEEV to replicate in mites suggests that viral transmission was mechanical. Two other arboviruses, VEEV and Langat virus (Flaviviridae), are also incapable of replicating in mites under experimental conditions [388,389].

Cocal virus (Rhabdoviridae) was originally isolated from a pool of Gigantolaelaps mites infesting rice rats in Trinidad and has since been detected in other insects and livestock [390]. SFTSV was detected by RT-PCR in mites (Laelaps echidninus, Leptotrombidium delicense, and Leptotrombidium scutellare) in China [391,392]. In another study, SFTSV was detected by RT-PCR in mites (Blankaartia acuscutellaris, Gahrliepia walchia, Leptotrombidium spp., and Schoengastia kanhaensis) infesting rodents in Thailand [393]. Transmission experiments have not been performed to determine whether mites are competent vectors of Cocal virus or SFTSV.

10.2. Other Vertebrate-Infecting Viruses

HTNV is not a recognized arbovirus, but mites have been suggested to play a minor role in its biological transmission [394]. HTNV was isolated from four species of field-collected mites (Eulaelaps stabularis, Haemolaelaps glasgowi, Laelaps cynognathus, and Leptotrombidium scutellare) in China [339,395,396,397,398]. Transstadial and transovarial transmission of HTNV by mites of several species have been reported [396,397,399,400,401]. HTNV was shown to persist in L. scutellare for up to 115 days and E. stabularis and H. glasgowi for up to 168 days [397,400,401]. Experimental studies have demonstrated that mites are capable of transmitting HTNV to uninfected mice [395,397,400,402]. In one study, HTNV was isolated from tissues harvested from mice that had been fed upon by infected mites (L. scutellare) fifteen days earlier [402]. Junin virus (Arenaviridae) is another vertebrate-infecting virus that has been detected in mites, with isolations made from spiny rat mites (Echinolaelaps echidninus) in Argentina [403].

Table 8. Arboviruses and other vertebrate-infecting viruses identified in field-collected mites.

Virus	Species Name	Abbreviation	Family	Recognized Arbovirus	¹ Experimental Evidence of Mite-Borne Virus Transmission	Citation
Cocal virus	Vesiculovirus cocal	COCV	Rhabdoviridae	Yes	² -	[390]
Eastern equine encephalitis virus	Alphavirus eastern	EEEV	Togaviridae	Yes	Yes (transmission to chickens, but highly inefficient)	[383,387]
Hantaan virus	Orthohantavirus hantanense	HTNV	Hantaviridae	-	Yes (transmission to mice; a minor role in HTNV transmission has been suggested)	[339,395,396,397,398,400,402]
Junin virus	Mammarenavirus juninense	JUNV	Arenaviridae	-	-	[403]
Severe fever with thrombocytopenia syndrome virus	Bandavirus dabieense	SFTSV	Phenuiviridae	Yes	-	[393]
St. Louis encephalitis virus	Orthoflavivirus louisense	SLEV	Flaviviridae	Yes	Conflicting data on whether mites can transmit SLEV to chickens, although they are considered inconsequential vectors	[374,375,376,377,378]
Western equine encephalitis virus	Alphavirus western	WEEV	Togaviridae	Yes	Yes (transmission to chickens, but highly inefficient)	[379,380,381,382]

¹ Excludes mechanical transmission. ² No.

10.3. Viruses Not Known to Infect Vertebrates

Many novel viruses have been detected in mites using metagenomics, although none are known to replicate in vertebrates [63,404].

11. Muscid Flies

Muscid flies (Muscidae) are found on every inhabited continent and comprise approximately 9000 species and 190 genera [405]. Many muscid flies are not primarily hematophagous while others, i.e., the horn fly (Hematobia irritans) and stable fly (Stomoxys calcitrans), are obligate blood-feeders [405,406]. Hematophagous muscid flies are mechanical vectors of various bacterial pathogens but are not recognized arboviral vectors, although they are known to harbor various vertebrate-infecting viruses (Table 9).

11.1. Recognized Arboviruses

WNV was isolated from stable flies infesting viremic American white pelicans (Pelecanus erythrorhynchos) in Montana, the U.S. [407]. The authors speculated that stable flies provide a source of infection via ingestion or mechanical transmission. ASFV DNA was detected in stable flies (Stomoxys calcitrans) collected on domestic pig farms in Lithuania [408]. In another study, ASFV DNA persisted for three days in stable flies that had engorged upon infected blood [409]. Transmission of ASFV to pigs via the ingestion of infected stable flies has also been reported [410]. However, there is no evidence to support biological transmission of ASFV by stable flies. Isolations of VSIV were made from house flies (Musca domestica) in Colorado, the U.S., but these insects are generally non-hematophagous [74].

Table 9. Arboviruses and other vertebrate-infecting viruses identified in field-collected muscid flies ¹.

Virus	Species Name	Abbreviation	Family	Recognized Arbovirus	² Experimental Evidence of Muscid-Borne Virus Transmission	Citation
African swine fever virus	Asfivirus haemorrhagiae	ASFV	Asfarviridae	Yes	³ -	[408]
Bovine leukosis virus	Deltaretrovirus bovleu	BLV	Retroviridae	-	-	[411]
West Nile virus	Orthoflavivirus nilense	WNV	Flaviviridae	Yes	-	[407]

¹ Excludes non-hematophagous species. ² Excludes mechanical transmission. ³ No.

11.2. Other Vertebrate-Infecting Viruses

Bovine leukosis virus (Retroviridae) has been detected in field collected horn flies, but these arthropods were unable to transmit bovine leukosis virus from infected to uninfected cattle under natural grazing conditions [411]. In another study, stable flies were fed artificial blood meals containing porcine reproductive and respiratory syndrome virus (PRRSV; Arteriviridae) [412]. PSSRV was detectable by qRT-PCR in the flies for up to 3 days, but the amount of viral nucleic acid decreased with time, indicating a lack of viral replication. Infectious virus persisted in the flies for only 24 h, and PRRSV has not been isolated from field-collected stable flies.

11.3. Viruses Not Known to Infect Vertebrates

Drosophila melanogaster Nora virus (Picornaviridae) has been detected in colonized horn flies, but there is no evidence to suggest that the virus replicates in vertebrate cells [413]. Densoviruses have also been detected in horn flies [414].

12. Phlebotomine Sandflies

Phlebotomine sandflies are small, hematophagous, mostly nocturnal insects belonging to the subfamily Phlebotominae (Psychodidae), which contains 1060 species and subspecies [415]. These insects have a wide geographic distribution, with the majority distributed throughout the tropics and subtropics, although none have been reported on the Pacific Islands or New Zealand. Phlebotomine sandflies are of major medical and veterinary importance, primarily because they are the principal vectors of Leishmania protozoan parasites [416,417,418,419]. More relevant to this review, phlebotomine sandflies harbor numerous arboviruses, most of which belong to the Phenuiviridae and Rhabdoviridae (Table 10) [420,421].

12.1. Recognized Arboviruses

Multiple viruses from the genera Phlebovirus (Phenuiviridae) have been detected in phlebotomine sandflies, although most have been detected once or on a few occasions. Several sandfly-associated phleboviruses cause human disease, including Sandfly fever Naples virus (SFNV), Sandfly fever Sicilian virus (SFSV), and Toscana virus (TOSV) [422,423,424,425,426,427,428,429,430,431,432,433,434,435]. SFNV and SFSV are endemic in the Mediterranean region and parts of Asia, where they cause an incapacitating febrile illness, while TOSV is a relatively common cause of meningitis and encephalitis in the Mediterranean region.

SFNV has been isolated from many pools of Phlebotomus spp. sandflies [54,436,437,438,439]. For example, SFNV was recovered from 4 of 59 pools containing 33,100 Phlebotomus spp. sandflies in India [437]. One positive pool contained P. papatasi, while the remaining three were not identified to species. In another study, 11 isolates were recovered from 26,734 Phlebotomus spp. sandflies in India [436,437,438]. Most isolates were presumed to be from P. papatasi, which constituted 99.2% of the total collection. Similarly, SFSV has been isolated from multiple pools of Phlebotomus spp. sandflies [54,422,431,440,441,442]. For instance, SFSV was identified in 49 of 304 pools containing 12,485 Phlebotomus spp. sandflies in Iran [441]. There is limited information confirming the vector competence of Phlebotomus spp. sandflies for SFNV and SFSV. However, in a study performed during World War II, eight of nine U.S. soldiers experimentally inoculated with an unknown phlebovirus (likely SFNV or SFSV) developed febrile illness [443]. P. papatasi were allowed to feed upon the patients and, after an extrinsic incubation period of 8 to 18 days, were then permitted to feed upon two uninfected volunteers. One of these individuals developed a febrile illness 3–4 days later, providing evidence of sandfly-borne phlebovirus transmission.

TOSV has been isolated from multiple pools of P. perniciosus and P. perfiliewi, with additional isolations also made from other Phlebotomus spp. (P. longicuspis, P. major, P. neglectus, P. papatasi, P. sergenti, and P. tobbi) and select Sergentomyia spp. (S. dentata and S. minuta) [444,445,446,447,448,449,450,451]. In one study, TOSV was isolated from 37 of 232 pools containing 16,374 sand flies (P. perniciosus and P. perfiliewi) in Italy [447]. The ability of P. perniciosus to support the replication of TOSV has been evaluated [452]. TOSV was detected in 95% of P. perniciosus following intrathoracic inoculation 2 to 12 days earlier. Viral titers increased over time. After oral exposure with of TOSV, up to 51% of P. perniciosus contained the virus at 4 to 14 days post-challenge, with minimal change in viral titers over time. Dissemination and transmission rates were not determined. In the same study, the ability of P. perniciosus to support the replication of Adria virus (a subtype of Salehabad virus), which is another sandfly-associated phlebovirus, was also assessed, yielding similar results to those reported for TOSV.

In a more recent study, the vector competence of four species of sandflies (Phlebotomus papatasi, Phlebotomus sergenti, Phlebotomus tobbi, and Sergentomyia schwetzi) for TOSV was evaluated [453]. Sandflies were orally challenged with two viral strains (denoted as TOSV-A and -B, respectively). At eight days post-challenge, TOSV-A was not detected in any sandflies. Likewise, at eight days post-challenge, TOSV-B was not detected in any P. papatasi or S. schwetzi, while TOSV-B infection rates of 52.8% and 6.2% were reported in P. tobbi and P. sergenti, respectively. Dissemination rates among infected P. tobbi and P. sergenti were 94.7% and 100%, respectively. Studies have not been performed to determine whether sandflies can experimentally transmit TOSV.

The ability of sandflies to support the replication of two other phleboviruses, Ntepes virus and RVFV, has been investigated [454,455,456,457]. Ntepes virus occurs in Kenya and has been isolated from sandflies (Phlebotomine spp. and Sergentomyia spp.) on three occasions [458,459]. Infection rates in P. duboscqi orally challenged with Ntepes virus 6, 10, and 15 days earlier were 10.7%, 5.4%, and 3.4%, respectively. None of the sandflies contained virus in their legs or salivary glands [454]. RVFV is primarily transmitted by mosquitoes, although it has been detected on at least one occasion in sandflies in a pool of Phlebotomus spp. in Nigeria (Genbank Accession No. ON856673). The vector competence of five species of sandflies (Lutzomyia longipalpis, Phlebotomus duboscqi, P. papatasi, P. sergenti, and S. schwetzi) for RVFV has been investigated [455,456,457]. In one experiment, 72 of 145 (49.7%) P. duboscqi that fed upon a viremic hamster became infected, including eight (5.5%) that developed disseminated infections [457]. Six sandflies with disseminated infections were allowed to feed on naïve hamsters, and five (83.3%) transmitted the virus. In contrast, only 6 of 236 (2.5%) and none of 378 (0%) L. longipalpis challenged with RVFV by intrathoracic inoculation and oral ingestion, respectively, transmitted the virus to susceptible hamsters [456]. Dissemination rates of 6% to 14% were reported in P. duboscqi, P. papatasi, P. sergenti, and S. schwetzi that fed upon RVFV-infected hamsters [455]. Additionally, 14% of P. papatasi and 50% of P. duboscqi, but none of the S. schwetzi, challenged with RVFV by intrathoracic inoculation were able to transmit the virus to susceptible hamsters.

Aside from SFNV, SFSV, and TOSV, only four phleboviruses have been detected in ≥20 pools of phlebotomine sandflies: Aguacate virus, Arbia virus (a subtype of Salehabad virus), Punta Toro virus, and Saddaguia virus (a subtype of SFNV). Aguacate virus occurs in Panama, where it was isolated from 31 of 3455 pools containing 252,512 Lutzomyia spp. sandflies [460]. Thirty-five isolates of Arbia virus were obtained from an unknown number of pools containing 10,843 P. perfiliewi and P. perniciosus in Italy and Spain [54,451]. Punta Toro virus was isolated on 49 occasions in a study where 219,512 sandflies from Panama were tested for virus [54,461]. Most isolations were made from Lutzomyia trapidoi. Saddaguia virus was isolated from 23 of 135 pools containing 3992 sandflies of various species, although most of the sample population were Phlebotomus perniciosus, Phlebotomus perfiliewi, and Phlebotomus longicuspis [462]. Vector competence experiments have not been performed with any of these viruses.

Multiple viruses belonging to the genus Vesiculovirus (Rhabdoviridae) have been detected in phlebotomine sandflies (Table 10) [463,464]. However, only three of these viruses have been isolated from over five pools: Chandipura virus (CHPV), VSIV, and VSNJV. CHPV is a human pathogen that has caused large outbreaks of encephalitis in India but also occurs in parts of Africa [465,466]. As already noted, VSIV and VSNJV are livestock pathogens that occur in the Americas [70,465,466].

Several studies report the detection of CHPV in Phlebotomus spp. and Sergentomyia spp. sandflies [467,468,469,470,471,472]. For example, CHPV was detected in 2 of 29 pools containing 625 Phlebotominae spp. sandflies in India [467]. In another study, three of nine pools of 277 Sergentomyia spp. sandflies collected during an outbreak of CHPV in India were positive for the virus [469]. Vector competence experiments have investigated whether sandflies are capable of transmitting CHPV [473,474]. CHPV replication and dissemination occurred in P. papatasi after intrathoracic inoculation, and clinical signs were observed in one of four mice fed upon by the infected P. papatasi after a seven-day extrinsic incubation period [474]. In another study, 65% of P. argentipes became infected after oral exposure to CHPV [473]. P. argentipes challenged with CHPV by intrathoracic inoculation could transmit the virus to naive mice and the estimated minimum transmission rate was 32%. Notably, experiments assessing the ability of orally challenged sandflies to transmit CHPV have not been performed.

VSIV and VSNJV has been detected in multiple pools of Lutzomyia spp. sandflies [54,80,475,476]. Additionally, evidence suggests that sandflies are competent vectors of these viruses [475,477,478,479,480]. Lutzomyia trapidoi became infected with VSIV after feeding upon viremic hamsters and could transmit the virus to naive mice after a 3-to-5-day extrinsic incubation period [477]. Lutzomyia shannoni supported the replication of VSNJV after challenge by both intrathoracic inoculation and membrane feeding [479]. VSNJV-inoculated sandflies could transmit the virus to a naive hamster after a 3-day extrinsic incubation period. Orally challenged sandflies could transmit the virus to mice after a 6-day extrinsic incubation period. Transmission electron microscopy revealed the presence of VSNJV in the salivary glands of L. shannoni five to six days after oral challenge [480]. High titers of VSNJV were reported in naturally infected L. shannoni [475].

Vector competence experiments have been performed with several other sandfly-associated vesiculoviruses. Carajas and Maraba viruses replicated in L. longipalpis following intrathoracic inoculation [481]. However, replication of Carajas virus did not occur in L. longipalpis after oral challenge, while sandflies were not orally challenged with Maraba virus. Vesicular stomatitis Alagoas virus replication occurred in L. longipalpis following intrathoracic inoculation and the infected sandflies could transmit the virus to naïve mice [482].

In addition to vesiculoviruses, many other viruses in the Rhabdoviridae have been detected in phlebotomine sandflies, including Charleville, Inhangapi, Iriri, Niakha, Santa Barbara, and Sripur viruses (Table 10) [464,483,484,485,486]. However, none of these viruses have been isolated from phlebotomine sandflies on more than five occasions and none have been assessed in vector competence experiments. Phlebotomine sandflies also harbor viruses various from the family Peribunyaviridae, including Caimito, Chilibre, Guamá, Rio Preto da Eva, Pacui, Santarém, Tapara, and Uriurana viruses [460,464,487,488]. None of these viruses have been isolated from more than 2 pools of phlebotomine sandflies, except for Pacui virus, which was recovered from 100 of 658 pools of 61,437 Lutzomyia flaviscutellata in Brazil [487]. The ability of phlebotomine sandflies to experimentally transmit these peribunyaviruses has not been evaluated.

12.2. Other Vertebrate-Infecting Viruses

Phlebotomine sandflies harbor several non-arboviral viruses capable of replicating in vertebrate cells, including Hedi virus (Phenuiviridae), Saboya virus (Flaviviridae), Santa Barbara virus (Rhabdoviridae), and Wuxiang virus (Phenuiviridae) (Table 10). Hedi virus was isolated from Phlebotomus chinensis in China and replicates in BHK-21 cells [489]. Saboya virus was originally isolated from a gerbil (Tatera kempi) in Senegal and later detected in other rodent species and hematophagous arthropods, including phlebotomine sandflies, in Senegal and Guinea [54,468,490,491,492]. Santa Barbara virus has been isolated on two occasions, from a pool of Psychodidae sandflies (species not specified) and a mouse (species not specified) in Brazil [464]. Wuxiang virus was isolated from 10 pools of Phlebotomus chinensis in China and is capable of replicating in BHK-21 cells [493].

12.3. Viruses Not Known to Infect Vertebrates

Several viruses with apparent insect-specific host ranges belonging to the families Flaviviridae, Phenuiviridae, and Rhabdoviridae have been identified in phlebotomine sandflies [494,495,496,497]. Viruses from other taxonomic groups have also been detected in sandflies and they too appear to be insect-specific [498,499].

Table 10. Arboviruses and other vertebrate-infecting viruses identified in field-collected phlebotomine sandflies ¹.

Virus	Species Name	Abbreviation	Family	Recognized Arbovirus	² Experimental Evidence of Sandfly-Borne Virus Transmission	Citation
Adana virus	Phlebovirus adanaense	ADAV	Phenuiviridae	Yes	³ -	[500]
Aguacate virus	Phlebovirus aguacateense	AGUV	Phenuiviridae	Yes	-	[460,501]
Alcube virus	Phlebovirus alcubeense	ACBV	Phenuiviridae	Yes	-	[502]
Ambé virus	Phlebovirus ambeense	ABEV	Phenuiviridae	Yes	-	[488,503]
Arbia virus (a subtype of Salehabad virus)	Phlebovirus salehabadense	ARBV	Phenuiviridae	Yes	-	[54,451]
Ariquemes virus (a subtype of Candiru virus)	Phlebovirus candiruense	CDUV	Phenuiviridae	Yes	-	[504]
Arrábida virus (a subtype of Sandfly fever Naples virus)	Phlebovirus napoliense	ARRV	Phenuiviridae	Yes	-	[505]
Balkan virus (a subtype of Sandfly fever Naples virus)	Phlebovirus napoliense	BALKV	Phenuiviridae	Yes	-	[506,507]
Bregalaka virus (a subtype of Salehabad virus)	Phlebovirus salehabadense	BREV	Phenuiviridae	Yes	-	[508]
Buenaventura virus	Phlebovirus buenaventuraense	BUEV	Phenuiviridae	Yes	-	[54,461,509]
Cacao virus	Phlebovirus cacaoense	CACV	Phenuiviridae	Yes	-	[54,460]
Caimito virus	Pacuvirus caimitoense	CAIV	Peribunyaviridae	Yes	-	[54]
Campana virus	Phlebovirus campanaense	CMAV	Phenuiviridae	Yes	-	[460,461]
Capira virus (a subtype of Punta Toro virus)	Phlebovirus toroense	CAPIV	Phenuiviridae	Yes	-	[461]
Carajas virus	Vesiculovirus carajas	CJSV	Rhabdoviridae	Yes	-	[481]
Chagres virus	Phlebovirus chagresense	CHGV	Phenuiviridae	Yes	-	[54,460],
Chandipura virus	Vesiculovirus chandipura	CHPV	Rhabdoviridae	Yes	Yes (transmission to mice)	[467,468,470,473,474,510]
Changuinola virus	Orbivirus changuinolaense	CGLV	Sedoreoviridae	Yes	-	[511]
Charleville virus	Sripuvirus charleville	CHVV	Rhabdoviridae	Yes	-	[483]
Chilibre virus	Pacuvirus chilibreense	CHIV	Peribunyaviridae	Yes	-	[54]
Corfou virus	Phlebovirus corfouense	CFUV	Phenuiviridae	Yes	-	[54]
Dashli virus	Phlebovirus dashliense	DASV	Phenuiviridae	Yes	-	[512]
Durania virus	Phlebovirus duraniaense	DRNV	Phenuiviridae	Yes	-	[503]
Fermo virus (a subtype of Sandfly fever Naples virus)	Phlebovirus napoliense	FERV	Phenuiviridae	Yes	-	[451]
Frijoles virus	Phlebovirus limboense	FRIV	Phenuiviridae	Yes	-	[54,513]
Granada virus (a subtype of Sandfly fever Naples virus)	Phlebovirus napoliense	GRV; GRAV	Phenuiviridae	Yes	-	[514]
Guamá virus	Orthobunyavirus guamaense	GMAV	Peribunyaviridae	Yes	-	[54,464,487,515]
Hedi virus	Phlebovirus hediense	HEDV	Phenuiviridae	-	-	[489]
Inhangapi virus	Arurhavirus inhangapi	INHV	Rhabdoviridae	Yes	-	[484]
Iriri virus	Curiovirus iriri	IRIRV	Rhabdoviridae	Yes	-	[307]
Isfahan virus	Vesiculovirus isfahan	ISFV	Rhabdoviridae	Yes	-	[516,517]
Ixcanal virus	Phlebovirus ixcanalense	IXCV	Phenuiviridae	Yes	-	[503]
Joá virus (a subtype of Frijoles virus)	Phlebovirus limboense	JOAV	Phenuiviridae	Yes	-	[488]
Karimabad virus	Phlebovirus karimabadense	KARV	Phenuiviridae	Yes	-	[54,441,518]
Koutango virus	Orthoflavivirus koutangoense	KOUV	Flaviviridae	Yes	-	[519]
Maraba virus	Vesiculovirus maraba	MARV	Rhabdoviridae	Yes	-	[481]
Medjerda Valley virus	Phlebovirus medjerdaense	MVV	Phenuiviridae	Yes	-	[520]
Morreton virus	Vesiculovirus morreton	MORV	Rhabdoviridae	Yes	-	[307]
Munguba virus	Phlebovirus mungubaense	MUNV	Phenuiviridae	Yes	-	[488]
Nique virus	Phlebovirus niqueense	NIQV	Phenuiviridae	Yes	-	[504]
Oriximiná virus	Phlebovirus oriximinaense	ORXV	Phenuiviridae	Yes	-	[504]
Pacui virus	Pacuvirus pacuiense	PACV	Peribunyaviridae	Yes	-	[487]
Perinet virus	Vesiculovirus perinet	PERV	Rhabdoviridae	Yes	-	[54]
Punique virus	Phlebovirus puniqueense	PUNV	Phenuiviridae	Yes	-	[439,521,522,523]
Punta Toro virus	Phlebovirus toroense	PTV	Phenuiviridae	Yes	-	[54,461]
Radi virus	Vesiculovirus radi	RADV	Rhabdoviridae	Yes	-	[54]
Rift Valley fever virus	Phlebovirus riftense	RVFV	Phenuiviridae	Yes	Yes (transmission to hamsters)	[456] ⁴ Genbank Accession No. ON856673
Rio Preto da Eva virus	Pacuvirus evaense	RPEV	Peribunyaviridae	Yes	-	[524]
Saboya virus	Orthoflavivirus saboyaense	SABV	Flaviviridae	-	-	[54,468,490,492]
Saddaguia virus (a subtype of Sandfly fever Naples virus)	Phlebovirus napoliense	SADV	Phenuiviridae	Yes	-	[462]
Salehabad virus	Phlebovirus salehabadense	SALV	Phenuiviridae	Yes	-	[525]
Sandfly fever Naples virus	Phlebovirus napoliense	SFNV	Phenuiviridae	Yes	Possibly (transmission of a virus, likely SFNV or SFSV, to human volunteers)	[54,443,526,527,528,529]
Sandfly fever Sicilian virus	Phlebovirus siciliaense	SFSV	Phenuiviridae	Yes	Possibly (transmission of a virus, likely SFNV or SFSV, to human volunteers)	[54,422,431,440,441,442,443]
Sandfly fever Turkey virus	⁵ none	SFTV	Phenuiviridae	Yes	-	[417, 428]
Santa Barbara virus	Arurhavirus santabarbara	SBAV	Rhabdoviridae	-	-	[464]
Santarém virus	none	STMV	Peribunyaviridae	Yes	-	[54]
Tapará virus	Phlebovirus taparaense	TPRV	Phenuiviridae	Yes	-	[488]
Tehran virus	Phlebovirus tehranense	THEV	Phenuiviridae	Yes	-	[54,441,530]
Toscana virus	Phlebovirus toscanaense	TOSV	Phenuiviridae	Yes	-	[425,428,450,531]
Turuna virus	Phlebovirus turunaense	TUAV	Phenuiviridae	Yes	-	[504]
Uriurana virus	Phlebovirus uriuranaense	URIV	Phenuiviridae	Yes	-	[488]
Vesicular stomatitis Alagoas virus	Vesiculovirus alagoas	VSAV	Rhabdoviridae	Yes	Yes (transmission to mice)	[482]
Vesicular stomatitis Indiana virus	Vesiculovirus indiana	VSIV	Rhabdoviridae	Yes	Yes (transmission to hamsters)	[54,80,477]
Vesicular stomatitis New Jersey virus	Vesiculovirus newjersey	VSNJV	Rhabdoviridae	Yes	Yes (transmission to hamsters)	[80,475,476,479]
Wuxiang virus	none	WUXV	Phenuiviridae	-	-	[493,532]
Yug Bogdanovac virus	Vesiculovirus bogdanovac	YBV	Rhabdoviridae	Yes	-	[54]
Zaba virus (a subtype of Salehabad virus)	Phlebovirus salehabadense	ZABAV	Phenuiviridae	Yes	-	[508]
Zerdali virus	Phlebovirus zerdaliense	ZERV	Phenuiviridae	Yes	-	[533]

¹ Not an exhaustive list. ² No. ³ Excludes mechanical transmission. ⁴ This GenBank entry was obtained by searching the “Nucleotide” section of the NCBI database using the terms “Rift Valley fever virus” and “Phlebotomus”. ⁵ Not recognized by the International Committee on Taxonomy of Viruses.

13. Tabanids

Tabanids (Tabanidae) are hematophagous flies, belonging to 4665 species, 177 genera, and five subfamilies (Adersinnae, Chrysopsinae, Pangoniinae, Scepsidinae, and Tabaninae) [534]. Tabanids have a wide geographic distribution and are found in almost all habitats. The best known and most widely studied tabanids are horse flies (Tabanus spp.) and deer flies (Chrysops spp.), although sometimes all tabanids are referred to as horse flies [535]. Horse flies and deer flies are nuisance biters of humans, livestock, and wildlife and are also mechanical vectors of several viruses of veterinary importance [536,537,538,539,540]. Moreover, one recognized arbovirus and other several vertebrate-infecting viruses have also been detected in tabanids (Table 11).

13.1. Recognized Arboviruses

Tabanids are not regarded as biological vectors of arboviruses, although Jamestown Canyon virus (JCV; Peribunyaviridae), a recognized arbovirus, was isolated from orange-sided horseflies (Hybomitra lasiophthalma) and ring-clawed deerflies (Chrysops cincticornis) in Wisconsin, the U.S. [541]. The ability of JCV to replicate in C. cincticornis and H. lasiophthalma has not been assessed, but JCV and Keystone virus (Peribunyaviridae) could not replicate in greenhead horse flies (Tabanus nigrovittatus) or deer flies (Chrysops atlanticus, Chrysops fuliginosus, and Chrysops obsoletus) under laboratory conditions [542].

Table 11. Arboviruses and other vertebrate-infecting viruses identified in field-collected tabanids.

Virus	Species Name	Abbreviation	Family	Recognized Arbovirus	¹ Experimental Evidence of Tabanid-Borne Virus Transmission	Citation
Jamestown Canyon virus	Orthobunyavirus jamestownense	JCV	Peribunyaviridae	Yes	² -	[541]
Kamenushka Hybomitra narna-like virus	³ none	KHNV	Narnaviridae	-	-	[543]
Medvezhye Chrysops ifla-like virus	none	MCIV	Iflaviridae	-	-	[543]
Medvezhye Chrysops narna-like virus 2	none	MCNV2	Narnaviridae	-	-	[543]
Medvezhye pound Haematopota permuto-like virus	none	MHPV	Permutotetraviridae	-	-	[543]
Medvezhye tabanus Toti-like virus	none	none assigned	Totiviridae	-	-	[543]
Melisia Chrysops solemo-like virus	none	MCSV	Solemoviridae	-	-	[543]
Volxa Hybomitra toti-like virus	none	VHTV	Totiviridae	-	-	[543]
Xanka Hybomitra negev-like virus	none	XHNV	unclassified	-	-	[543]

¹ Excludes mechanical transmission. ² No. ³ Not recognized by the International Committee on Taxonomy of Viruses.

13.2. Other Vertebrate-Infecting Viruses

The viromes of Chrysops, Haematopota Hybomitra, and Tabanus spp. tabanids from Russia were characterized, resulting in the detection of many novel viruses and one recognized virus (Big Sioux River virus; Dicistroviridae) [543]. Big Sioux River virus and eight of the newly discovered viruses were able to replicate in pig embryo kidney (PEK) cells, but it is unknown whether they are biologically transmitted by tabanids to vertebrates (Table 11).

13.3. Viruses Not Known to Infect Vertebrates

Tabanus rufidens flavivirus (Flaviviridae) was first identified in horse flies (Tabanus rufidens) in Japan, but it is most closely related phylogenetically to insect-specific flaviviruses and cannot replicate in BHK-21 cells [544]. Other viruses with apparent insect-specific host ranges have also been detected in tabanids [358,543].

14. Triatomines

Triatomines (also known as kissing bugs) are obligate blood-feeders belonging to the subfamily Triatominae (Reduviidae) [545,546,547]. The subfamily contains 14 genera and at least 140 species. Triatomines primarily occur in the Americas but are also present in parts of Africa and Asia. Triatomines primarily feed on vertebrate blood but occasionally feed on the hemolymph of arthropods and are best known as the vectors of Trypanosoma cruzi, the protozoan that causes Chagas disease [548,549].

14.1. Recognized Arboviruses

Triatomines are not considered to be arboviral vectors, although SLEV and VEEV persisted for 30 and 124 days, respectively, in triatomines (Rhodnius prolixus) that had fed upon viremic mice [550]. Fifteen days post-engorgement, R. prolixus transmitted VEEV, but not SLEV, to mice. SLEV and VEEV have never been detected in field-collected triatomines.

14.2. Other Vertebrate-Infecting Viruses

No vertebrate-infecting viruses have been detected in field-collected triatomines.

14.3. Viruses Not Known to Infect Vertebrates

The first virus to be detected in naturally infected triatomines is Triatoma virus (Dicistroviridae), which appears to have a host range restricted to triatomines [551,552,553,554]. Other triatomine-associated viruses have since been discovered, including Meccus longipennis virus 1 (Iflaviridae), Drosophila melanogaster Nora virus (Noraviridae), and Rhodnius prolixus viruses 1 to 4 (unclassified), but none are known to replicate in vertebrates [555,556].

15. Tsetse Flies

The family Glossinidae comprises exclusively large, blood-feeding insects, known as tsetse flies. Approximately 31 species and subspecies are recognized, and all occur throughout much of sub-Saharan Africa [557,558,559]. These flies are vectors of trypanosome parasites, which cause sleeping sickness (or human African trypanosomiasis) in humans and nagana (or animal African trypanosomiasis) in livestock. Tsetse flies are not recognized arboviral vectors and are not known to harbor any vertebrate-infecting viruses. However, several viruses with apparent arthropod-specific host ranges have been detected in tsetse flies [560,561,562].

16. Conclusions

In conclusion, growing evidence suggests that lesser-studied hematophagous arthropods play important roles in arbovirus transmission. Notably, at least 132 recognized species/subtypes of arboviruses have been detected in non-mosquito, non-tick hematophagous arthropods (Table 2, Table 3, Table 4, Table 5, Table 6, Table 7, Table 8, Table 9, Table 10 and Table 11). Nonetheless, significant gaps remain in our understanding of their involvement in arboviral transmission. It is particularly important that we further investigate arthropods known or suspected to be epidemiologically important arbovirus vectors, namely, blackflies, cimicids, Culicoides midges, and phlebotomine sandflies. For example, blackflies are competent vectors of VSV under laboratory conditions and a high prevalence of VSV has been detected in blackflies during epizootics, but their role in the natural transmission of VSV is unclear. It is not known whether the majority of VSV-infected blackflies collected during outbreaks acquired the virus by biological or mechanical transmission. Further, it is not known whether blackflies contribute to the maintenance of VSV during non-epidemic periods or if they transmit the virus to wildlife. Additionally, a definitive reservoir host of VSV has yet to be identified. As such, blackflies have not been conclusively identified as epidemiologically important biological vectors of VSV and are often referred to as suspected or incriminated vectors. Experimental studies specifically designed to resolve these uncertainties are needed. Similarly, there is strong evidence that cimicids are the principal vectors of FMV; however, they cannot be classified as epidemiologically important vectors without field data demonstrating consistent FMV infection in wild cimicid populations. Addressing this knowledge gap will require targeted field investigations within the known geographic range of FMV.

Culicoides midges have been implicated as epidemiologically important biological vectors of several arboviruses, but additional research is required to fully characterize their role in transmission. Notably, there is a paucity of transmission experiments. For instance, Culicoides midges are considered primary vectors of AHSV, yet only three transmission experiments have been performed. Two studies used a natural host, but only one (an unpublished study) demonstrated successful transmission. Similarly, Culicoides midges are believed to be the primary vectors of AKAV, although most experimental infection studies focused on replication and dissemination rather than actual transmission. The exception is a study involving Culicoides species found outside the endemic range of AKAV. Another apparent Culicoides-borne arbovirus is BEFV, but successful experimental transmission of the virus is yet to be reported. EHDV is also likely transmitted primarily by Culicoides midges; however, most vector competence experiments have focused on North American Culicoides species and EHDV serotypes, leaving other species and serotypes unexplored. Transmission of MDV to uninfected blood suspensions by Culicoides midges has been documented, but transmission experiments involving vertebrate hosts have not been performed. Culicoides midges can experimentally transmit OROV to hamsters, but a high prevalence of OROV in field-collected Culicoides midges has not been consistently reported. Finally, SBV is assumed to be primarily Culicoides-borne, but all experimental infection studies have focused on replication and dissemination, without investigating transmission. These gaps highlight the need for well-designed transmission studies to clarify the role of Culicoides midges in the natural transmission cycles of these viruses.

Phlebotomine sandflies are suspected vectors of several phleboviruses, but their role in arbovirus transmission remains poorly characterized. Although numerous phleboviruses have been detected in phlebotomine sandflies, and some have been efficiently transmitted under laboratory conditions, no phlebovirus has been consistently detected in naturally infected sandflies and conclusively demonstrated to be transmitted to vertebrate hosts. As such, sandflies should be regarded as suspected or incriminated vectors of phleboviruses rather than epidemiologically important vectors. For example, phlebotomine sandflies are suspected to be the principal vectors SFNV, SFSV, and TOSV, all of which are significant human pathogens, yet comprehensive vector competence data are lacking. In a study performed during World War II, phlebotomine sandflies were shown to transmit a phlebovirus to soldier volunteers; however, the virus, while suspected to SFNV or SFSV, was never definitively identified. While experimental studies have shown that sandflies can support the replication and dissemination of TOSV, their ability to transmit the virus has not been tested. In addition to phleboviruses, sandflies are also suspected vectors of several vesiculoviruses, including CHPV. Although phlebotomine sandflies can transmit CHPV to mice under laboratory conditions, the insects were infected by intrathoracic inoculation, which, unlike oral challenge, bypasses the natural midgut barrier.

Two arthropod groups discussed in this review, namely, fleas and mites, may play a minor role in the biological transmission of arboviruses. Fleas have been implicated as minor vectors of TBEV based on two studies reporting the isolation of TBEV from multiple flea species and two other studies demonstrating experimental transmission by fleas. TBEV has also been shown to persist in fleas for up to six days. However, a key limitation of these studies is the lack of viral titer quantification, making it unclear whether TBEV actively replicates in fleas. In contrast, TBEV has been repeatedly isolated from field-collected ticks and efficiently transmitted by ticks under laboratory conditions, firmly establishing ticks as their primary vectors. Mites have been implicated as minor vectors of HTNV, with multiple studies reporting the isolation of the virus from field-collected mites and at least four studies demonstrating experimental transmission. Although HTNV is not officially classified as an arbovirus, its persistence in mites for up to 168 days indicates possible replication. Further studies are needed to determine whether HTNV titers increase over time in mites and whether the virus replicates in mite-derived cell lines. SLEV, EEEV, and WEEV have also been isolated from field-collected mites and transmitted by these arthropods under laboratory conditions, but transmission has either been inefficient or unable to be verified in follow-up studies, and it is firmly established that mosquitoes are their primary vectors. For all other arthropod groups (bat flies, hippoboscid flies, lice, muscid flies, tabanids, triatomines, and tsetse flies), there is insufficient evidence to suggest that they are biological vectors of arboviruses because none have been shown to transmit arboviruses under experimental conditions.

As previously mentioned, all known arboviruses, except for ASFV, have RNA genomes. RNA viruses evolve more rapidly than DNA viruses because their RNA-dependent RNA polymerases are inherently error-prone and lack proof-reading capabilities, resulting in higher mutation rates and, subsequently, an increased likelihood of adapting to two distinct biological environments (arthropod vectors and vertebrate hosts) [563,564,565,566,567]. Additionally, many RNA viruses have segmented genomes, allowing for reassortment, which further promotes genetic diversity and adaptation. In this regard, most arboviruses detected in Culicoides spp. midges, and phlebotomine sandflies belong to the Peribunyaviridae and Sedoreoviridae. The rapid evolutionary dynamics of RNA viruses likely contribute to the fact that the reservoirs hosts of many of the viruses discussed in this review remain unknown.

ZOVER is a database of zoonotic and vector-borne viruses launched in 2021 [568]. It integrates two previously established databases of bat- and rodent-associated viruses and compiles current information on viruses associated with mosquitoes and ticks. However, no equivalent resources exists for the other arthropod groups discussed in this review. This lack of a centralized database for arboviruses detected in non-mosquito, non-tick vectors presents a significant gap in our ability to identify underrecognized or emerging transmission cycles that may threaten human and animal health. Comprehensive risk assessments depend on a thorough understanding of all potential vector species. In the absence of data on these lesser-studied arthropods, our capacity to model transmission dynamics, forecast outbreaks, and implement effective vector control measures is reduced. Advancing our knowledge of the vector competence, host associations, and ecological roles of these groups is critical for a more complete understanding of arbovirus transmission and for strengthening both medical and veterinary preparedness.

Funding

The author is supported by grants from the National Institutes of Health (R01AI114720 and R03 AI177172).

Conflicts of Interest

The author has no conflicts of interest to declare.

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Blitvich, B.J. The Role of Hematophagous Arthropods, Other Than Mosquitoes and Ticks, in Arbovirus Transmission. Viruses 2025, 17, 932. https://doi.org/10.3390/v17070932

AMA Style

Blitvich BJ. The Role of Hematophagous Arthropods, Other Than Mosquitoes and Ticks, in Arbovirus Transmission. Viruses. 2025; 17(7):932. https://doi.org/10.3390/v17070932

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Blitvich, Bradley J. 2025. "The Role of Hematophagous Arthropods, Other Than Mosquitoes and Ticks, in Arbovirus Transmission" Viruses 17, no. 7: 932. https://doi.org/10.3390/v17070932

APA Style

Blitvich, B. J. (2025). The Role of Hematophagous Arthropods, Other Than Mosquitoes and Ticks, in Arbovirus Transmission. Viruses, 17(7), 932. https://doi.org/10.3390/v17070932

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The Role of Hematophagous Arthropods, Other Than Mosquitoes and Ticks, in Arbovirus Transmission

Abstract

1. Arboviruses

2. Hematophagous Arthropods and Vector Competence

3. Bat Flies

3.1. Recognized Arboviruses

3.2. Other Vertebrate-Infecting Viruses

3.3. Viruses Not Known to Infect Vertebrates

4. Blackflies

4.1. Recognized Arboviruses

4.2. Other Vertebrate-Infecting Viruses

4.3. Viruses Not Known to Infect Vertebrates

5. Cimicids

5.1. Recognized Arboviruses

5.2. Other Vertebrate-Infecting Viruses

5.3. Viruses Not Known to Infect Vertebrates

6. Culicoides Midges

6.1. Recognized Arboviruses

6.2. Other Vertebrate-Infecting Viruses

6.3. Viruses Not Known to Infect Vertebrates

7. Fleas

7.1. Recognized Arboviruses

7.2. Other Vertebrate-Infecting Viruses

7.3. Viruses Not Known to Infect Vertebrates

8. Hippoboscid Flies

8.1. Recognized Arboviruses

8.2. Other Vertebrate-Infecting Viruses

8.3. Viruses Not Known to Infect Vertebrates

9. Lice

9.1. Recognized Arboviruses

9.2. Other Vertebrate-Infecting Viruses

9.3. Viruses Not Known to Infect Vertebrates

10. Mites

10.1. Recognized Arboviruses

10.2. Other Vertebrate-Infecting Viruses

10.3. Viruses Not Known to Infect Vertebrates

11. Muscid Flies

11.1. Recognized Arboviruses

11.2. Other Vertebrate-Infecting Viruses

11.3. Viruses Not Known to Infect Vertebrates

12. Phlebotomine Sandflies

12.1. Recognized Arboviruses

12.2. Other Vertebrate-Infecting Viruses

12.3. Viruses Not Known to Infect Vertebrates

13. Tabanids

13.1. Recognized Arboviruses

13.2. Other Vertebrate-Infecting Viruses

13.3. Viruses Not Known to Infect Vertebrates

14. Triatomines

14.1. Recognized Arboviruses

14.2. Other Vertebrate-Infecting Viruses

14.3. Viruses Not Known to Infect Vertebrates

15. Tsetse Flies

16. Conclusions

Funding

Conflicts of Interest

References

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