Skip to main content
Download PDF

Interspecies co-feeding transmission of Powassan virus between a native tick, Ixodes scapularis, and the invasive East Asian tick, Haemaphysalis longicornis

Parasites & Vectors volume 17, Article number: 259 (2024) Cite this article

Abstract

Background

Powassan virus, a North American tick-borne flavivirus, can cause severe neuroinvasive disease in humans. While Ixodes scapularis are the primary vectors of Powassan virus lineage II (POWV II), also known as deer tick virus, recent laboratory vector competence studies showed that other genera of ticks can horizontally and vertically transmit POWV II. One such tick is the Haemaphysalis longicornis, an invasive species from East Asia that recently established populations in the eastern USA and already shares overlapping geographic range with native vector species such as I. scapularis. Reports of invasive H. longicornis feeding concurrently with native I. scapularis on multiple sampled hosts highlight the potential for interspecies co-feeding transmission of POWV II. Given the absence of a clearly defined vertebrate reservoir host for POWV II, it is possible that this virus is sustained in transmission foci via nonviremic transmission between ticks co-feeding on the same vertebrate host. The objective of this study was to evaluate whether uninfected H. longicornis co-feeding in close proximity to POWV II-infected I. scapularis can acquire POWV independent of host viremia.

Methods

Using an in vivo tick transmission model, I. scapularis females infected with POWV II (“donors”) were co-fed on mice with uninfected H. longicornis larvae and nymphs (“recipients”). The donor and recipient ticks were infested on mice in various sequences, and mouse infection status was monitored by temporal screening of blood for POWV II RNA via quantitative reverse transcription polymerase chain reaction (q-RT-PCR).

Results

The prevalence of POWV II RNA was highest in recipient H. longicornis that fed on viremic mice. However, nonviremic mice were also able to support co-feeding transmission of POWV, as demonstrated by the detection of viral RNA in multiple H. longicornis dispersed across different mice. Detection of viral RNA at the skin site of tick feeding but not at distal skin sites indicates that a localized skin infection facilitates transmission of POWV between donor and recipient ticks co-feeding in close proximity.

Conclusions

This is the first report examining transmission of POWV between co-feeding ticks. Against the backdrop of multiple unknowns related to POWV ecology, findings from this study provide insight on possible mechanisms by which POWV could be maintained in nature.

Graphical Abstract

Background

Powassan virus (POWV) is a member of the Orthoflavivirus genus, belonging to the Flaviviridae family. POWV is primarily identified in the northeastern and Great Lakes regions of the USA, Southern Canada, and the Russian Far East. This zoonotic virus is transmitted to humans by several Ixodes species ticks [1,2,3], and it is the only tick-borne flavivirus endemic to North America. POWV was first identified as a human pathogen in 1958 in Powassan, Ontario when the virus was isolated from the brain tissue of a child who succumbed to encephalitis [4]. Initial clinical signs of POWV disease in humans include fever, headache, vomiting, and weakness. POWV infections can also progress to an altered mental state, encephalitis, seizures, paralysis, and coma [4,5,6,7]. The fatality rate for POWV cases is estimated to be 12.5–17.9% [8], with more than half of survivors experiencing long-term neurological sequelae [2]. There appears to be an increase in human cases of POWV in the USA in recent decades [3, 8, 9]. This rise could potentially be attributed to heightened surveillance of ticks and tick-borne diseases, increased clinician awareness and improved diagnostics, or the expansion of Ixodes scapularis populations [3, 10].

Following its initial identification, POWV was shown to be maintained in nature through transmission cycles involving Ixodes species ticks and mammalian hosts [11, 12]. POWV exhibits two genetic lineages: POWV lineage I (POWV I) and POWV lineage II (POWV II), also recognized as deer tick virus. Although serologically indistinguishable, these two POWV lineages share 84% and 93–94% nucleotide and amino acid sequence identity, respectively [13, 14]. It is also becoming increasingly apparent that POWV lineages I and II are ecologically distinct. POWV I was initially thought to be sustained in enzootic cycles involving Ixodes cookei and groundhogs/mustelids [12, 15] or Ixodes marxi and squirrels [11]. POWV II is typically isolated from Ixodes scapularis, which tend to have a catholic host preference and frequently parasitize humans. While POWV II was originally thought to be maintained in an enzootic transmission cycle between human-biting I. scapularis and white-footed mice (Peromyscus leucopus) [1, 16], the virus has never been isolated from mice in the field. Furthermore, laboratory infection of P. leucopus with POWV resulted in no overt clinical disease [17], suggesting that although P. leucopus are frequently exposed to POWV, they are an unlikely reservoir host. Various studies have assessed POWV experimental infections in several animal species. One study involving groundhogs, striped skunk, and fox squirrels [18] and another focusing on snowshoe hares [19] found that none of these vertebrate hosts developed POWV I or II viremias greater than ~ 1000 PFU/mL (Plaque-Forming Unit/mL), which indicates that these vertebrates are improbable reservoir hosts for POWV. A recent study used host-specific retrotransposon targeted real-time PCR on questing I. scapularis nymphs and demonstrated that there is a strong positive correlation between ticks feeding on shrews at a given site and the prevalence of POWV II infection in the ticks [20]. These data, together with the detection of POWV II RNA in the brain of one shrew, led the authors to conclude that shrews are a potential reservoir host for POWV II [20]; however, laboratory infection of shrews with POWV, as well as transmission studies with ticks, are needed to fully assess the reservoir host capacity of shrews.

In field studies, POWV I and POWV II have been detected in alternate hard-tick species (e.g., Dermacentor andersoni, Dermacentor variabilis) other than the main Ixodes vector species outlined above [21, 22]. Furthermore, recent laboratory studies demonstrate that D. variabilis, H. longicornis and Amblyomma americanum are competent vectors for POWV II [23, 24]. These findings suggest that the transmission cycles of POWV I and POWV II could be more complex than the single tick vector species/single reservoir host paradigm [25].

Haemaphysalis longicornis is an ixodid tick native to East and Southeast Asia and eastern Russia that has become established in Australia, New Zealand, several Pacific Island nations, and most recently, the USA. This tick species was first documented outside of a US port of entry when several H. longicornis were collected in Staten Island, New York in 2014–2015 [26]. Since its initial detection in the USA, established populations of H. longicornis have been documented in 19 states along the East Coast and Appalachia, with the most recent detection reported in Georgia in September 2022 [27]. The first record of H. longicornis biting a human in the USA was reported in June 2018 [28], indicating that populations of this tick species are already present in outdoor areas frequented by humans. In recent years, several studies have evaluated the ability of H. longicornis to acquire, maintain, and transmit tick-borne pathogens endemic to North America, ultimately demonstrating through laboratory studies that this tick is a competent vector for Rickettsia rickettsii and POWV II [24, 29]. This tick’s ability to transovarially transmit Heartland virus was also reported [30].

Haemaphysalis longicornis demonstrates adaptability to a broad range of climates in both its native and invasive ranges, including seasonal climates. Ecological niche modeling predicting the potential spread of H. longicornis in North America has identified several regions beyond the Northeast and East Coast as suitable for sustaining populations of H. longicornis. These regions include the Southeast, the Midwest, the California coast, the Pacific Northwest, and southeastern Canada [31,32,33]. Both the current distribution of H. longicornis (eastern USA and Appalachia), as well as the tick’s predicted geographic range expansion, overlap with the distribution of native tick species (e.g., I. scapularis, D. variabilis, A. americanum, etc.) that are competent vectors for POWV II.

In its native geographic range, both parthenogenetic and bisexual strains of H. longicornis have been observed; however, the North American populations of this invasive tick appear to reproduce exclusively by parthenogenesis [34]. Haemaphysalis longicornis is characterized as an aggressive biter and a host generalist. In its native range, it feeds on mammalian, avian, and human hosts [34]. Since its recent establishment in North America, this tick species has been observed feeding on a diverse array of hosts. Documented instances include various domestic animals and wildlife, ranging from rodent species to dogs and white-tailed deer [27, 35]. Interestingly, field surveillance studies conducted on Staten Island, New York detected H. longicornis feeding concurrently with at least one native tick species (A. americanum or I. scapularis) on 78% of sampled hosts [36, 37]. Haemaphysalis longicornis was found co-feeding on the same host body region alongside I. scapularis on 81.3% of white-tailed deer, 95.2% of opossums, 46.2% of raccoons, 50% of striped skunks, 50% of marmots, and 60% of feral cats; however, exact proximities between the co-feeding H. longicornis and I. scapularis were not reported [36].

While horizontal and vertical transmission are the most commonly evaluated routes of tick-borne virus transmission, tick-borne viruses can also be transmitted between co-feeding ticks. Co-feeding transmission occurs when an infected tick transmits virus to an uninfected tick while simultaneously feeding on the same host, often in close proximity on the skin. The phenomenon of co-feeding transmission (i.e., nonviremic transmission) was first described in the late 1980s when uninfected ticks, co-feeding on the same host with Thogoto virus-infected ticks, acquired virus even in the absence of host viremia [38]. Prior to this work by Jones et al., it was believed that an obvious viremia and systemic infection was necessary for ticks to acquire virus during blood feeding; however, Jones’ work was paradigm shifting in that it demonstrated that tick-to-tick transmission of virus can occur via nonviremic hosts. To date, no reservoir host has been clearly defined for POWV II; therefore, it is possible that POWV II may be sustained in natural transmission cycles via nonviremic transmission between co-feeding ticks [39], as has been shown for tick-borne encephalitis virus (TBEV) transmission in Europe [25].

With the many unknowns of POWV ecology and the yet to be determined role of invasive H. longicornis in tick-borne virus transmission cycles in North America, the objective of this study was to investigate whether and to what extent nonviremic co-feeding transmission of POWV II occurs when uninfected H. longicornis co-feed on the same host as an infected I. scapularis. By investigating co-feeding transmission of POWV II in the context of two different tick species with overlapping geographic distributions in portions of North America, the present study will provide us with important insight into a mechanism through which POWV could be maintained in nature.

Methods

Ethics and biosafety

All experiments involving virus-infected and mock-infected ticks fed on mice were conducted within a dedicated room of the animal biosafety level 3 (ABSL-3) facility. Infected ticks were handled and housed in arthropod containment level 3 (ACL-3) facilities. Bloodmeals were provided to uninfected ticks for colony maintenance by feeding ticks on guinea pigs in animal biosafety level 2 (ABSL-2) facilities. All biosafety level 3 (BSL-3), ABSL-3, and ACL-3 experiments were performed in accordance with protocols approved by the Institutional Biosafety Committee and the Institutional Animal Care and Use Committee (IACUC). Animal work was conducted using protocols approved by the University of South Alabama (USA) IACUC (protocol nos. 1619216, 2065516).

Cells and virus

African green monkey kidney (VeroE6) cells were maintained in Dulbecco’s Modified Eagle Medium (DMEM, Genesee Scientific) with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin. Cells were cultured in an incubator set to 37 °C and 5% CO2 and used to generate virus stock. The Spooner strain of Powassan virus lineage II (POWV II) was acquired from the World Reference Center for Emerging Viruses and Arboviruses at the University of Texas Medical Branch. The stock had previously been passaged once on suckling mouse brains and five times on Vero cells. It was then passaged once on VeroE6 cells. The stock virus titer was determined by focus forming immunoassay as previously described [40].

Ticks and animals

Ticks

A pathogen-free colony of H. longicornis was maintained by feeding ticks on Hartley guinea pigs under ACL-2 conditions. This colony originated from ticks collected in New York state in 2018. This lineage of H. longicornis has been propagated under laboratory conditions for seven generations without supplementation. Male and female I. scapularis were provided by Oklahoma State University. This colony originated from engorged females collected in Stillwater, Oklahoma in 1991. All tick vials were stored at 21 °C with 90–95% relative humidity. The photoperiod of the room was set on a 16:8 h light:dark cycle.

Mice

The 5-week-old male and female BALB/c mice were purchased from The Jackson Laboratory (Bar Harbor, ME). Mice were allowed to acclimate to the environment for a minimum of 5 days before the commencement of the experiments, at which point they were 6–6.5-weeks-old. During the study, mice were individually housed in ventilated cage systems maintained in a 12:12 h light:dark environment. Room humidity and temperature were closely regulated for the caging environments. Food and water were provided to mice ad libitum.

Tick feeding capsules used for tick infestations on mice

Mice were randomly assigned to infection or mock groups 1 day before tick infestation, and one tick containment capsule was attached per mouse. Capsules were fashioned from 2 mL cryotubes. The base of each tube was cut to leave approximately 3 mm of remaining tube below the screw-cap lid. The tops of the lids were cut to allow for an opening. The capsule was then inserted in the middle of a round piece of elastic adhesive bandage. Mice were anesthetized with isoflurane anesthesia to effect for the capsule placement procedures. The dorsum and lateral sides of each mouse were shaved with an electric razor and the capsule adhered to the mouse skin using Kamar livestock adhesive (Kamar Inc., Steamboat Springs, CO). At this point, mice were individually housed in micro-isolator cages. On the day of tick infestation, ticks were placed inside the capsules and a piece of fine mesh fabric was placed under the lid to allow for both tick containment and air exchange. Capsule lids were secured using masking tape. Capsule integrity was checked daily throughout tick infestation. Capsules were reinforced with adhesive and bandages as needed.

RNA extractions from tick and mouse tissues

Pools of up to five fed recipient H. longicornis larvae, individual fed recipient H. longicornis nymphs, fed donor I. scapularis females, and mouse skin biopsies were homogenized with 3 mm sterile stainless-steel beads on a Qiagen Tissue Lyser II at 30 Hz for 3 min. These homogenized tissue samples as well as mouse blood were all stored in TRIzol reagent. A hybrid of TRIzol and Qiagen RNeasy Mini Kit protocols was used to perform RNA extractions. We have previously demonstrated that this hybrid protocol inactivates virus and produces high-quality RNA [24, 41]. A Nanodrop One Spectrophotometer (Thermo Fisher Scientific) was used to determine total RNA quantity and purity.

Detection of viral RNA by q-RT-PCR

Absolute quantification of POWV II RNA quantities in tick and mouse samples was determined by quantitative reverse transcription real-time PCR (q-RT-PCR) as previously described [42]. Viral RNA quantities are expressed on a Log10 scale as the number of POWV II NS5 gene fragment copies per ng of RNA after normalization to a standard curve produced using serial tenfold dilutions of a 464-base pair POWV II NS5 gene fragment to estimate viral burden. q-RT-PCR was performed using forward (5′—GATCATGAGAGCGGTGAGTGACT—3′) and reverse (5′ –GGATCTCACCTTTGCTATGAATTCA—3′) primers and a probe (/56-FAM/TGAGCACCTTCACAGCCGAGCCAG/36-TAMSp/) specific to POWV II NS5 gene, as previously described [43]. When performing the assay, 1 µL of RNA was added to the appropriate wells of a 96-well PCR plate and samples were run in triplicate; 10 µM dilutions of the forward and reverse primers and probe were used with reagents from the iTaq Universal SYBR Green One-Step Kit (BioRad). The total reaction volume was 20 µL per well. Plates were sealed and run on a LightCycler® 480 II PCR System (Roche) at the following cycle settings: 10 min at 50 °C, 1 min at 95 °C, 10 s at 95 °C, and 30 s at 60 °C for 45 cycles.

Microinjection of female I. scapularis with POWV II

Infection via microinjection of female I. scapularis was achieved by injecting 292 nL, containing 1000 focus-forming units (FFU) of POWV II stock, into the immobilized tick’s anal aperture using glass microneedles, a digitally operated microinjector with a footswitch, and a dissecting microscope as previously described [24]. For the mock-infected groups, an equivalent volume of DMEM media was injected. Glass microneedles were made using a micropipette puller (Sutter Instrument) and glass capillaries (World Precision Instruments). Each capillary had an internal diameter of 0.530 mm and an outer diameter of 1.14 mm; these microneedles were pulled such that the tip had a smaller diameter than that of the tick anal pore. POWV II-injected I. scapularis were housed in ACL-3 facilities and were monitored for mortality twice daily for 4 days after microinjection.

In vivo tick co-feeding experiments

The female I. scapularis microinjected with POWV II are defined as “donor” ticks in this study. For all in vivo experiments, each mouse was infested with a single microinjected female I. scapularis and a single naïve male I. scapularis. Microinjected I. scapularis females were infested on mice at 25–27 days post-microinjection. Haemaphysalis longicornis larvae and nymphs from the pathogen-free tick colony are defined as “recipient” ticks in this study. For all in vivo experiments, each mouse was infested with up to 53 H. longicornis larvae or up to 16 H. longicornis nymphs. The notation d.p.a. (days post-attachment) consistently indicates the attachment day of the donor I. scapularis female ticks.

Overview of tick infestation schemes used for the in vivo experiments

A staggered tick infestation scheme [“Staggered (donor first) tick infestation”] was used for the original in vivo experiment whereby mice were infested with donor I. scapularis first. Then, at 2 d.p.a. (relative to the I. scapularis donors), mice were infested with recipient H. longicornis nymphs (Fig. 1a). A simultaneous tick infestation scheme (“Simultaneous donor and recipient tick infestation”) was used for a subsequent in vivo experiment whereby mice were infested by donor I. scapularis and recipient H. longicornis nymphs at the same time (0 d.p.a.) (Fig. 1b). Finally, a flipped version of the staggered tick infestation scheme [“Staggered (recipient first) tick infestation”] was implemented whereby mice were infested by recipient H. longicornis larvae or nymphs first (−2 d.p.a.), and then by donor I. scapularis (0 d.p.a.) (Fig. 1c, d). This tick infestation scheme was repeated in three different cohorts of mice, each assessed separately over the course of approximately 1 year.

Fig. 1
figure 1

Tick infestation strategies used for the in vivo experiments. a “Staggered (donor first) tick infestation” timeline. Each mouse was infested with a single POWV II-injected donor Ixodes scapularis at 0 d.p.a. At 2 d.p.a., each mouse was then infested with recipient Haemaphysalis longicornis nymphs. Submandibular blood was collected from each mouse at −1, 2, and 5 d.p.a. Mice were euthanized at 14 d.p.a. or earlier if they met humane endpoints. b “Simultaneous donor and recipient tick infestation” timeline. Each mouse was infested with a single POWV II-injected donor I. scapularis and recipient H. longicornis nymphs at the same time (0 d.p.a.). Submandibular blood was collected every other day from each mouse. Mice were euthanized 2–3 days after completion of H. longicornis feeding for mice that did not reach humane endpoints. c “Staggered (recipient first) tick infestation” timeline. At −2 d.p.a., each mouse was infested with recipient H. longicornis larvae or nymphs, and 2 days later (0 d.p.a.), each mouse was then infested with a single POWV II-injected donor I. scapularis. Submandibular blood was collected every other day from each mouse. Mice were euthanized 2–3 days after completion of H. longicornis feeding for mice that did not reach humane endpoints. d “Staggered (recipient first) tick infestation with daily bleeds” and 2 or 4 d.p.a. necropsy timeline. At −2 d.p.a., each mouse was infested with recipient H. longicornis larvae or nymphs, and 2 days later (0 d.p.a.), each mouse was then infested with a single POWV II-injected donor I. scapularis. Submandibular blood was collected daily from each mouse until euthanasia at 2 or 4 d.p.a. d.p.a. days post-attachment, consistently indicates the attachment day of the donor I. scapularis female ticks, POWV II Powassan virus lineage II.

Full size image

Daily mouse clinical observations and weights

Mice were anesthetized at times of tick placement, capsule repair, tick removal, and submandibular bleed using isoflurane anesthesia. All mice were observed twice daily during the study period through the use of a clinical score chart that assesses weight loss, appearance (e.g., reduced grooming, ruffled coat, ocular/nasal discharge, hunched posture, etc.), neurologic disease (e.g., weak grip, paresis, ataxia, tremors, head tilt, partial paralysis, seizures, etc.), and behavior (e.g., subdued when stimulated, unresponsive). Mice that reached any of the following humane endpoints were immediately euthanized: > 20% weight loss, paralysis, seizures, gasping respiration, hemorrhage, prostrate, or unresponsive. Euthanasia occurred via isoflurane overdose followed by terminal cardiac puncture and cervical dislocation. For the original in vivo experiment [“Staggered (donor first) tick infestation, Fig. 1a], mice were euthanized either at 14 d.p.a. or earlier (7–11 d.p.a.) when they met humane endpoints. For all other in vivo experiments, euthanasia was performed via the same method 2–3 days after completion of H. longicornis feeding.

Tick-infested mice were also checked daily during this clinical observation for tick attachment status. Attachment of the female I. scapularis “donors” was monitored two to three times a day for the first 2 days. Adult female tick attachment day (0 d.p.a.) was carefully recorded. Mice were removed from the study if their corresponding donor I. scapularis failed to attach. A naïve I. scapularis male was added to each capsule to facilitate the female I. scapularis attachment, although it is likely that I. scapularis females had already mated prior to infestation on the host as they were co-housed with I. scapularis males for ~ 1 week prior to feeding. Mice were checked daily for naïve H. longicornis “recipient” (larvae and nymphs) attachment. Engorged “recipients” were retrieved from the capsules after detaching from the skin. “Donor” ticks were collected upon reaching full engorgement and detaching from the host, or at the time of euthanasia, whichever occurred first. Blood was collected from each mouse via submandibular bleed. For the “Staggered (donor first) tick infestation” study, submandibular blood was collected at −1, 2, and 5 d.p.a. (Fig. 1a). For the “Simultaneous donor and recipient tick infestation” study (Fig. 1b) and the first “Staggered (recipient first) tick infestation” study (Fig. 1c), submandibular blood was collected from each mouse every other day after I. scapularis donor attachment. For the final “Staggered (recipient first) tick infestation with daily bleeds” study (Fig. 1d), submandibular blood was collected daily from each mouse after I. scapularis donor attachment and until euthanasia.

Necropsies

Necropsies of mock-infected and POWV-infected mice were conducted under ABSL-3 conditions. Terminal blood collected via cardiac bleed was equally divided into TRIzol Reagent storage and serum separator tubes. Brain, liver, kidney, spleen, testes, skin from the tick attachment site (inside the capsule), and skin from outside of the tick attachment site (~ 3 cm caudal to the capsule) were harvested from mice during necropsy (Fig. 3a), and 4-mm biopsy punch tools were used to harvest the skin samples. A portion of each tissue sample was stored in TRIzol Reagent and the other portion in 10% neutral-buffered formalin. Formalin was exchanged after a minimum of 24 h and was allowed a minimum of 72 h total contact time with tissues.

Statistical analyses

For the comparison of the experimental groups, analysis of variance (ANOVA) followed by Tukey as a post-test was used; p ≤ 0.05 was considered statistically significant.

Results

Staggered (donor first) tick infestation

A total of 24 donor I. scapularis females were individually transferred to 24 mice. Then, at 2 days post-attachment (d.p.a.), each mouse was infested with seven to ten recipient H. longicornis nymphs (Fig. 1a). Submandibular blood was collected from each mouse at −1, 2, and 5 d.p.a. to assess mouse viremia status (Fig. 1a). In total, 2 of the media-injected control donor I. scapularis and 11 of the virus-injected donor I. scapularis engorged and fed to repletion. Mice showing mild signs of Powassan (POW) disease were euthanized at the scheduled time of 14 d.p.a., whereas other mice developed neuroinvasive POW disease and reached humane endpoints requiring euthanasia between 7 d.p.a. and 11 d.p.a. These findings demonstrate that our POWV-infected I. scapularis donors transmit infectious virus to the mice.

All recipient H. longicornis nymphs from these 13 remaining mice were individually screened for POWV II RNA by q-RT-PCR (Table 1). No viral RNA was detected in recipient H. longicornis that co-fed on mice infested with a media-injected donor I. scapularis (Table 1). POWV II RNA was detected in 59.4% of recipient H. longicornis that co-fed on mice infested with a virus-injected donor I. scapularis (Table 1). At 2 d.p.a. and/or 5 d.p.a., POWV II RNA was detected in the blood of 10 out of the 11 mice that were fed upon by the virus-injected donor I. scapularis. Therefore, it was not possible to determine whether the presence of viral RNA in recipient H. longicornis resulted from mouse viremia or co-feeding transmission of virus between donor and recipient ticks. Only one mouse had no detectable viral RNA in both blood samples collected at 2 d.p.a. and 5 d.p.a.; however, the lack of data regarding viremia status from other days during the tick infestation prevents any clear conclusion about whether this mouse supported H. longicornis acquisition of POWV II RNA via nonviremic co-feeding transmission.

Table 1 Rate of detection of POWV II RNA by q-RT-PCR in fed recipient H. longicornis from the “Staggered (donor first) tick infestation”
Full size table

Simultaneous donor and recipient tick infestation

In total, eight donor I. scapularis females were individually transferred to eight mice, and mice were each simultaneously infested with nine to ten recipient H. longicornis nymphs (Fig. 1b). Submandibular blood was collected every other day from each mouse to assess viremia status during tick co-feeding (Fig. 1b). All donor I. scapularis (media-injected and POWV II-injected) attached and fed. As expected, no viral RNA was detected in media-injected donor I. scapularis, whereas viral RNA was detected in 100% of virus-injected donor I. scapularis (Supplementary Fig. 1). All recipient H. longicornis nymphs were individually screened for POWV II RNA by q-RT-PCR upon detachment and completion of feeding (Table 2). No viral RNA was detected in recipient H. longicornis nymphs that co-fed on mice infested with a media-injected donor I. scapularis; however, POWV II RNA was detected in 44.4% of recipient H. longicornis nymphs that co-fed on mice infested with a virus-injected donor I. scapularis (Table 2). POWV II RNA was detected in the blood of mice corresponding to virus positive recipient H. longicornis nymphs on the same day or on a day before these recipient H. longicornis nymphs completed feeding. Therefore, similar to results obtained in the “Staggered (donor first) tick infestation” (Fig. 1a) described above, it is not possible to determine whether the presence of viral RNA in recipient H. longicornis resulted from mouse viremia or from co-feeding transmission of virus between the donor I. scapularis and recipient H. longicornis.

Table 2 Rate of detection of POWV II RNA by q-RT-PCR in fed recipient H. longicornis from the “Simultaneous donor and recipient tick infestation”
Full size table

Staggered (recipient first) tick infestation

In this tick infestation experiment, recipient H. longicornis were infested on mice prior to the donor I. scapularis in an effort to allow the recipient ticks to start feeding prior to the mouse developing viremia. Thus, at −2 d.p.a., 8 mice were each infested with 47–50 recipient H. longicornis larvae and another 8 mice were each infested with 9–10 recipient H. longicornis nymphs (Fig. 1c); 2 days later (0 d.p.a.), the mice were each infested with one donor I. scapularis female tick. Submandibular blood was collected every other day from each mouse to assess mouse viremia status (Fig. 1c), and 15 out of the 16 donor I. scapularis (media-injected and POWV II-injected) attached and fed. As expected, no POWV II viral RNA was detected in media-injected donor I. scapularis, whereas viral RNA was detected in 100% of POWV II-injected donor I. scapularis (Supplementary Fig. 1). Recipient H. longicornis were screened for POWV II RNA by q-RT-PCR (Table 3). No viral RNA was detected in recipient H. longicornis larvae or nymphs that co-fed on mice infested with a media-injected donor I. scapularis, whereas POWV II RNA was detected in 19.4% of recipient H. longicornis larvae (screened in pools) and 14.3% of recipient H. longicornis nymphs (screened individually) that co-fed on mice infested with a virus-injected donor I. scapularis (Table 3).

Table 3 Rate of detection of POWV II RNA by q-RT-PCR in fed recipient H. longicornis from the “Staggered (recipient first) tick infestation”
Full size table

Figure 2a shows the timeline of viremia for each of the five mice that yielded engorged H. longicornis recipients positive for POWV II RNA. The bar graphs show the temporal development of viremia as well as the days where virus-positive engorged H. longicornis recipients were collected (as indicated by the “+” symbol) from each mouse. For mouse 2, mouse 3, and mouse 5, virus-positive recipient H. longicornis were collected at timepoints when viral RNA was also detected in the blood on the same day or on the day prior. POWV II RNA was detected in five recipient H. longicornis nymphs that detached at 2 d.p.a. from mouse 4, but no POWV II RNA was detected in blood at 2 d.p.a. for this mouse (as shown in Fig. 2a with the “Not Detected” symbol), suggesting that nonviremic co-feeding transmission occurred. Similarly, viral RNA was detected in one pool of recipient H. longicornis larvae that detached from mouse 1 at 2 d.p.a., and although no blood sample was collected from mouse 1 at that timepoint, viral RNA was not detected in the blood samples collected the day before (1 d.p.a.) and the day after (3 d.p.a.) the pool of recipient H. longicornis larvae screened positive for POWV II RNA. These findings suggest that nonviremic co-feeding transmission is likely to have occurred for the virus-positive H. longicornis recipients collected from mouse 1. However, the lack of daily blood samples collected from each mouse in this experiment prevents a definitive conclusion regarding the mode of acquisition of POWV II RNA by these recipient H. longicornis nymphs.

Fig. 2
figure 2

Detection of POWV II RNA in the blood of mice that yielded recipient H. longicornis positive for POWV II RNA. a Mice 1–5 are from the “Staggered (recipient first) tick infestation.” Each of these mice were bled every other day. b Mice 6–9 are from the “Staggered (recipient first) tick infestation with daily bleeds.” These mice were bled daily and euthanized at 2 d.p.a. c Mice 10–14 are from the “Staggered (recipient first) tick infestation with daily bleeds.” These mice were bled daily and euthanized at 4 d.p.a. POWV II RNA was detected in mouse blood and recipient H. longicornis samples via q-RT-PCR. Viral RNA quantities are expressed as the number of NS5 gene fragment copies per ng of RNA after normalization to a standard curve. Each bar represents the viral RNA quantity in the blood for that mouse at a specific time point. “ND” means that POWV II RNA was Not Detected in the blood. “NB” means that No Blood was collected on that day. The “+” symbol means that fed recipient H. longicornis screened positive for POWV II RNA at that timepoint. d.p.a. days post-attachment, consistently indicates the attachment day of the donor I. scapularis female ticks, POWV II Powassan virus lineage II, RNA ribonucleic acid, q-RT-PCR quantitative reverse transcription polymerase chain reaction.

Full size image

Staggered (recipient first) tick infestation with daily bleeds

To assess the viremia status of the vertebrate host with more granularity, a tick infestation experiment was designed in which recipient H. longicornis were infested on mice 2 days prior to the donor I. scapularis, each mouse was bled daily, and then mice were euthanized at either 2 d.p.a. or 4 d.p.a. (Fig. 1d). Here, 11 mice were each infested with 35–53 recipient H. longicornis larvae and another 13 mice were each infested with 12–16 recipient H. longicornis nymphs; 2 days later (0 d.p.a.), the mice were each infested with 1 donor I. scapularis female tick. All donor I. scapularis (media-injected and POWV II-injected) attached and fed. No viral RNA was detected in media-injected donor I. scapularis, but POWV II RNA was detected in 100% of virus-injected donor I. scapularis (Supplementary Fig. 1). All recipient H. longicornis were screened for POWV II RNA by q-RT-PCR (Table 4). No viral RNA was detected in recipient H. longicornis larvae or nymphs that co-fed on mice infested with a media-injected donor I. scapularis. However, in mice euthanized at 2 d.p.a. and infested with a virus-injected donor I. scapularis, POWV II RNA was detected in 50% of recipient H. longicornis larvae (screened in pools) and 12.5% of recipient H. longicornis nymphs (screened individually). On mice euthanized at 4 d.p.a. and infested with a virus-injected donor I. scapularis, POWV II RNA was detected in 100% of pooled H. longicornis larvae and 4.1% of individual H. longicornis nymphs (Table 4).

Table 4 Rate of detection of POWV II RNA by q-RT-PCR in fed recipient H. longicornis from the “Staggered (recipient first) tick infestation with daily bleeds”
Full size table

Figure 2b, c represents the nine mice that yielded engorged H. longicornis recipients positive for POWV II RNA. Figure 2b, c bar graphs show the temporal development of viremia (assessed by daily blood samples collected from every mouse) relative to the days where virus-positive fed H. longicornis recipients were collected (as indicated by the “+” symbol) from each mouse. POWV II RNA was detected in recipient H. longicornis nymphs that detached at 2 d.p.a. from mouse 8, mouse 9, and mouse 13, but no viral RNA was detected in the blood of these mice at any timepoint. Recipient H. longicornis larvae detached from mouse 12 at 3 d.p.a. and 4 d.p.a. were combined into a single pool for POWV II RNA screening. Viral RNA was detected in this larval pool, but no viral RNA was detected in the blood of mouse 12 at any timepoint. Together, these findings clearly demonstrate that H. longicornis larvae and nymphs were capable of acquiring POWV II RNA from donor I. scapularis via nonviremic co-feeding transmission on mouse 8, mouse 9, mouse 12, and mouse 13 (Fig. 2b, c).

For mice in the “Staggered (recipient first) tick infestations with daily bleeds” (Fig. 1d), skin biopsies collected proximal to the tick feeding site (from inside the tick feeding capsule) and skin biopsies collected distal to the tick feeding site (from outside the tick feeding capsule) were screened by q-RT-PCR for POWV II RNA (Fig. 3a). No POWV II RNA was detected in skin biopsy #3 for mice 6–9 and 11–13 (Fig. 3b). The blood of mouse 8, mouse 9, mouse 12, and mouse 13 was also negative for POWV II RNA at each day of tick feeding (Fig. 2b, c), so the absence of viral RNA in skin biopsy 3 provides additional evidence that there was no systemic infection in these mice. Furthermore, in the nonviremic mice, detection of viral RNA in skin from the tick feeding site (biopsies 1 and 2) but not at distal skin sites (biopsy 3) indicates that a localized skin infection may facilitate transmission of POWV II RNA between donor and recipient ticks co-feeding in close proximity.

Fig. 3
figure 3

Detection of POWV II RNA in skin biopsies proximal and distal to the tick co-feeding site. a Location of skin biopsies #1, #2, and #3. b Mice 6–14 are from the “Staggered (recipient first) tick infestation with daily bleeds”. These mice were bled daily and euthanized at 2 d.p.a. or 4 d.p.a. POWV II RNA was detected in mouse skin biopsy samples via q-RT-PCR. Viral RNA quantities are expressed as the number of NS5 gene fragment copies per ng of RNA after normalization to a standard curve. Each point represents the viral RNA quantity in a skin biopsy for a specific mouse, as indicated in the legend. d.p.a. days post-attachment, consistently indicates the attachment day of the donor I. scapularis female ticks, POWV II Powassan virus lineage II, RNA ribonucleic acid, q-RT-PCR quantitative reverse transcription polymerase chain reaction.

Full size image

Discussion

Co-feeding transmission has been observed with various vector-borne pathogens, and its importance in natural transmission cycles of tick-borne encephalitis virus (TBEV), a close relative to POWV, was clearly demonstrated several decades ago [44,45,46]. TBEV primary vectors are Ixodes ricinus in Europe and Ixodes persulcatus in Fennoscandia, Russia, and Asia [47]. Nymphs and larvae of these two species are typically active in the same seasons and share the same rodent hosts, which enables intraspecies co-feeding transmission between juvenile tick stages [48]. In Europe, tick co-feeding transmission of TBEV plays an important role in the persistence of this virus within certain foci in nature [46]. Additionally, epidemiological modeling shows that co-feeding transmission alone (in the absence of vertical transmission or amplifying hosts) could be sufficient to sustain POWV in tick populations [39]. During nonviremic co-feeding transmission, an infected tick can transmit a virus directly to a naïve tick co-feeding on the same host (often in close proximity) without requiring the virus to be circulating in the blood of the vertebrate host. Therefore, co-feeding transmission can be highly efficient since the virus does not need to first replicate and disseminate within the vertebrate host to be transmitted between ticks, nor is a susceptible vertebrate host required for virus transmission via this mechanism. Labuda et al. showed that TBEV can be transmitted from infected to uninfected ticks feeding on certain virus-immune natural rodent hosts via nonviremic co-feeding transmission [44]. Despite these wild rodents being immunized against the virus, their neutralizing antibodies to TBEV did not prevent subsequent transmission of TBEV between co-feeding ticks [44]. Perhaps in North America, certain vertebrate hosts for Ixodes species ticks (e.g., P. leucopus) that are refractory to POWV infection and do not develop a patent viremia are indeed playing a role in sustaining POWV by serving as a transient bridge while the ticks serve as both the reservoirs as well as the vectors.

Under laboratory conditions, TBEV-infected adult I. ricinus co-fed with naïve Rhipicephalus appendiculatus or Haemaphysalis inermis nymphs, and TBEV-infected adult Dermacentor reticulatus co-fed with naïve H. inermis nymphs, all resulted in interspecies co-feeding transmission of TBEV [47, 48]. The present study also involved donor Ixodes species females that were infected with a flavivirus and co-fed with immature recipient ticks from a different genus. Data from this study demonstrate experimental interspecies co-feeding transmission of POWV II RNA between an adult virus-infected I. scapularis and naïve recipient H. longicornis, both in the presence and absence of host viremia. The rate of detection of POWV II RNA was highest in recipient H. longicornis that co-fed on viremic mice (Tables 1, 2). However, multiple recipient H. longicornis larvae and nymphs were able to acquire viral RNA while feeding concurrently and in close proximity to the infected I. scapularis donor with no viral RNA detected in the mouse blood. In the initial “Staggered (recipient first) tick infestation” (Fig. 1c), we showed co-feeding transmission of POWV II RNA from donor I. scapularis to recipient H. longicornis nymphs and larvae (Table 3, Fig. 2a). Although blood samples from mouse 1 and mouse 4 were not screened daily for POWV II RNA, the data suggest that these mice were nonviremic on days that the recipient H. longicornis screened positive for viral RNA (Fig. 2a). Nonviremic co-feeding transmission of viral RNA was then confirmed by the “Staggered (recipient first) tick infestation with daily bleeds” (Fig. 1d) via the detection of POWV II RNA in several recipient H. longicornis nymphs and larvae from mouse 8, mouse 9, mouse 12, and mouse 13 (Table 4) and the demonstration of the absence of viremia in these mice (Fig. 2b, c). In the “Staggered (recipient first) tick infestation” experiments, pathogen-free H. longicornis recipients were allowed to pre-feed on mice for 2 days prior to addition of the POWV-infected I. scapularis donors. Thus, it is possible that the presence of H. longicornis saliva already deposited at the tick co-feeding site facilitates transmission of virus, as previously demonstrated in studies showing that tick salivary gland extract facilitates POWV transmission and dissemination in the vertebrate host [41].

In the present study, POWV II RNA was detected in the skin from the tick feeding site (skin biopsies #1 and #2) but not at distal tick-free skin sites (skin biopsy #3) in the nonviremic mice (Fig. 3). These findings suggest that a localized skin infection and the spatiotemporal feeding proximity of the donor I. scapularis with the recipient H. longicornis supported nonviremic co-feeding transmission of POWV II RNA. Previous studies also highlighted the importance of ticks feeding in close proximity and localized virus infection at the site of tick attachment during the early stages of flavivirus transmission via tick co-feeding. When TBEV-infected and uninfected I. ricinus were infested on natural murine host species and either allowed to co-feed in close proximity inside the same capsule or retained in separate capsules and fed at a greater distance, TBEV was preferentially recruited to the tick-infested skin sites compared with tick-free skin, and TBEV transmission between co-feeding ticks was correlated with localized skin infection at the tick feeding sites [49]. Furthermore, co-feeding transmission of TBEV between infected and uninfected ticks was most efficient when ticks fed in close proximity but did still occur when ticks co-fed at a greater distance [49]. In nature, ticks often feed closely clustered together on their host. This is especially the case for many species of immature ticks that feed on rodents, whereby ~ 90% are commonly found on the ears, neck, and head of their murine hosts [50,51,52]. The present study evaluated tick co-feeding within a small (~ 1–1.5 cm) area, but it will be important for future studies to examine the outcome of co-feeding transmission of POWV when ticks feed at greater distances on the same vertebrate host. Additionally, while our data suggest that a localized skin infection plays an important role in the nonviremic co-feeding transmission of POWV, the mechanism underlying this phenomenon needs to be defined. Therefore, we have initiated studies investigating the mechanistic aspects of nonviremic co-feeding transmission.

Recent studies demonstrate that D. variabilis, A. americanum, and H. longicornis are competent vectors for POWV II under laboratory conditions [23, 24]. By detecting POWV II RNA in engorged H. longicornis that co-fed with POWV-infected I. scapularis, the present study provides evidence that multiple tick species, including invasive H. longicornis, could contribute to POWV maintenance in nature via interspecies co-feeding transmission; however, future investigations need to focus on detecting infectious virus in the recipient ticks to fully demonstrate virus transmission via tick co-feeding. Furthermore, all of the recipient H. longicornis in this study were processed as engorged larvae/nymphs, and as such, we did not screen recipient ticks post-molt for the presence of virus; however, previous studies have shown evidence of transstadial transmission of flavivirus occurring in H. longicornis [24, 53]. Despite these limitations, the present study clearly showed that interspecies tick co-feeding transmission of POWV II RNA occurs: (1) on viremic and nonviremic vertebrate hosts, (2) when various life stages of H. longicornis serve as the recipient, and (3) when donor and recipient ticks are infested on the host in different sequences. Perhaps most relevant to our current understanding of POWV ecology, whereby a definitive vertebrate reservoir host for POWV remains undefined, is our discovery that interspecies tick co-feeding transmission of POWV can indeed occur on a nonviremic host. Together, these data suggest that natural transmission cycles of POWV could involve a combination of multiple tick species and nonviremic co-feeding transmission, with or without a vertebrate reservoir host. Forms of vertical transmission of POWV (e.g., transovarial and transstadial transmission) have also been demonstrated in laboratory-maintained ticks [54], and recently in field-collected ticks [55]. To date, the relative importance of horizontal, vertical, and co-feeding transmission to the maintenance of POWV in nature is unknown. Therefore, future studies delineating the efficiency of these various routes of transmission (e.g., co-feeding transmission, transovarial transmission) will be critical toward improving our understanding of POWV ecology.

Conclusions

Our findings demonstrate that under laboratory conditions, POWV II RNA can be transmitted from infected I. scapularis to naïve H. longicornis via co-feeding transmission, both in the presence and absence of host viremia. Results from this study suggest that nonviremic co-feeding transmission could play a role in the maintenance of POWV in nature; however, future laboratory, field, and modeling studies are needed to better understand POWV persistence and transmission in tick populations.

Availability of data and materials

All data supporting the conclusions of this study are provided within the manuscript or supplementary information file.

Abbreviations

ABSL-2:

Animal biosafety level 2

BSL-3:

Biosafety level 3

ABSL-3:

Animal biosafety level 3

ACL-3:

Arthropod containment level 3

POWV II:

Powassan virus lineage II

RNA:

Ribonucleic acid

q-RT-PCR:

Quantitative reverse transcription polymerase chain reaction

d.p.a.:

Days post-attachment (of the donor I. scapularis female)

TBEV:

Tick-borne encephalitis virus

References

  1. Telford SR, Armstrong PM, Katavolos P, Foppa I, Garcia AS, Wilson ML, Spielman A. A new tick-borne encephalitis-like virus infecting New England deer ticks, Ixodes dammini. Emerg Infect Dis. 1997;3:165–70.

    Article  PubMed  PubMed Central  Google Scholar 

  2. Ebel GD. Update on Powassan virus: emergence of a North American tick-borne flavivirus. Annu Rev Entomol. 2010;55:95–110.

    Article  CAS  PubMed  Google Scholar 

  3. Campbell O, Krause PJ. The emergence of human Powassan virus infection in North America. Ticks Tick Borne Dis. 2020;11:101540.

    Article  PubMed  Google Scholar 

  4. McLean DM, Donohue WL. Powassan virus: isolation of virus from a fatal case of encephalitis. Can Med Assoc J. 1959;80:708–11.

    CAS  PubMed  PubMed Central  Google Scholar 

  5. Tavakoli NP, Wang H, Dupuis M, Hull R, Ebel GD, Gilmore EJ, Faust PL. Fatal case of deer tick virus encephalitis. N Engl J Med. 2009;360:2099–107.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  6. Cavanaugh CE, Muscat PL, Telford SR, Goethert H, Pendlebury W, Elias SP, Robich R, Welch M, Lubelczyk CB, Smith RP. Fatal deer tick virus infection in Maine. Clin Infect Dis. 2017;65:1043–6.

    Article  PubMed  Google Scholar 

  7. Yu Q, Matkovic E, Reagan-Steiner S, Denison AM, Osborn R, Salamat SM. A fatal case of Powassan virus encephalitis. J Neuropathol Exp Neurol. 2020;79:1239–43.

    Article  CAS  PubMed  Google Scholar 

  8. Kakoullis L, Vaz VR, Kaur D, Kakoulli S, Panos G, Chen LH, Behlau I. Powassan virus infections: a systematic review of published cases. Trop Med Infect Dis. 2023;8(12):508. https://doi.org/10.3390/tropicalmed8120508.

    Article  PubMed  PubMed Central  Google Scholar 

  9. CDC. Powassan virus human disease cases by year of illness onset, 2004–2022. 2023. https://www.cdc.gov/powassan/statistics-data/historic-data.html. Accessed 16 Jan 2024.

  10. Eisen RJ, Eisen L. The blacklegged tick, Ixodes scapularis: an increasing public health concern. Trends Parasitol. 2018;34:295–309.

    Article  PubMed  PubMed Central  Google Scholar 

  11. McLean DM, Larke RP. Powassan and silverwater viruses: ecology of two ontario arboviruses. Can Med Assoc J. 1963;88:182–5.

    CAS  PubMed  PubMed Central  Google Scholar 

  12. McLean DM, Best JM, Mahalingam S, Chernesky MA, Wilson WE. Powassan virus: summer infection cycle, 1964. Can Med Assoc J. 1964;91:1360–2.

    CAS  PubMed  PubMed Central  Google Scholar 

  13. Beasley DW, Suderman MT, Holbrook MR, Barrett AD. Nucleotide sequencing and serological evidence that the recently recognized deer tick virus is a genotype of Powassan virus. Virus Res. 2001;79:81–9.

    Article  CAS  PubMed  Google Scholar 

  14. Kuno G, Artsob H, Karabatsos N, Tsuchiya KR, Chang GJ. Genomic sequencing of deer tick virus and phylogeny of Powassan-related viruses of North America. Am J Trop Med Hyg. 2001;65:671–6.

    Article  CAS  PubMed  Google Scholar 

  15. McLean DM, Cobb C, Gooderham SE, Smart CA, Wilson AG, Wilson WE. Powassan virus: persistence of virus activity during 1966. Can Med Assoc J. 1967;96:660–4.

    CAS  PubMed  PubMed Central  Google Scholar 

  16. Ebel GD, Campbell EN, Goethert HK, Spielman A, Telford SR. Enzootic transmission of deer tick virus in New England and Wisconsin sites. Am J Trop Med Hyg. 2000;63:36–42.

    Article  CAS  PubMed  Google Scholar 

  17. Mlera L, Meade-White K, Saturday G, Scott D, Bloom ME. Modeling Powassan virus infection in Peromyscus leucopus, a natural host. PLoS Negl Trop Dis. 2017;11:e0005346.

    Article  PubMed  PubMed Central  Google Scholar 

  18. Nemeth NM, Root JJ, Hartwig AE, Bowen RA, Bosco-Lauth AM. Powassan virus experimental infections in three wild mammal species. Am J Trop Med Hyg. 2021;104:1048–54.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Zarnke RL, Yuill TM. Powassan virus infection in snowshoe hares (Lepus americanus). J Wildl Dis. 1981;17:303–10.

    Article  CAS  PubMed  Google Scholar 

  20. Goethert HK, Mather TN, Johnson RW, Telford SR. Incrimination of shrews as a reservoir for Powassan virus. Commun Biol. 2021;4:1319.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  21. Thomas LA, Kennedy RC, Eklund CM. Isolation of a virus closely related to Powassan virus from Dermacentor andersoni collected along North cache la Poudre river. Colo Proc Soc Exp Biol Med. 1960;104:355–9.

    Article  CAS  PubMed  Google Scholar 

  22. Hart C, Hassett E, Vogels CBF, Shapley D, Grubaugh ND, Thangamani S. Powassan virus lineage I in field-collected Dermacentor variabilis ticks, New York, USA. Emerg Infect Dis. 2023;29:415–7.

    Article  PubMed  PubMed Central  Google Scholar 

  23. Sharma R, Cozens DW, Armstrong PM, Brackney DE. Vector competence of human-biting ticks Ixodes scapularis, Amblyomma americanum and Dermacentor variabilis for Powassan virus. Parasit Vectors. 2021;14:466.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  24. Raney WR, Herslebs EJ, Langohr IM, Stone MC, Hermance ME. Horizontal and vertical transmission of Powassan virus by the invasive Asian Longhorned tick, Haemaphysalis longicornis, under laboratory conditions. Front Cell Infect Microbiol. 2022;12:923914.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  25. Brackney DE, Vogels CBF. The known unknowns of Powassan virus ecology. J Med Entomol. 2023;60:1142–8.

    Article  PubMed  Google Scholar 

  26. Bajwa W, Kennedy A, Vincent Z, Heck G, Riaj S, Shah Z, Tsynman L, Casal C, Haynes S, Cornman H, Egizi A, Stromdahl E, Nadolny R. Earliest records of the Asian Longhorned tick (Acari: Ixodidae) in Staten Island, New York, and subsequent population establishment, with a review of its potential medical and veterinary importance in the United States. J Med Entomol. 2024;61(3):764–71. https://doi.org/10.1093/jme/tjae019.

    Article  PubMed  Google Scholar 

  27. Agriculture U. D. O. National Haemaphysalis longicornis (Asian Longhorned tick) Situation Report as of March 7, 2024; 2024. https://www.aphis.usda.gov/sites/default/files/longhorned-tick-sitrep.pdf. Accessed 4 Apr 2024.

  28. Wormser GP, McKenna D, Piedmonte N, Vinci V, Egizi AM, Backenson B, Falco RC. First recognized human bite in the United States by the Asian Longhorned tick, Haemaphysalis longicornis. Clin Infect Dis. 2020;70:314–6.

    Article  PubMed  Google Scholar 

  29. Stanley HM, Ford SL, Snellgrove AN, Hartzer K, Smith EB, Krapiunaya I, Levin ML. The ability of the invasive Asian Longhorned tick Haemaphysalis longicornis (Acari: Ixodidae) to acquire and transmit Rickettsia rickettsii (Rickettsiales: Rickettsiaceae), the agent of rocky mountain spotted fever, under laboratory conditions. J Med Entomol. 2020;57:1635–9.

    Article  CAS  PubMed  Google Scholar 

  30. Raney WR, Perry JB, Hermance ME. Transovarial transmission of heartland virus by invasive Asian Longhorned ticks under laboratory conditions. Emerg Infect Dis. 2022;28:726–9.

    Article  PubMed  PubMed Central  Google Scholar 

  31. Raghavan RK, Barker SC, Cobos ME, Barker D, Teo EJM, Foley DH, Nakao R, Lawrence K, Heath ACG, Peterson AT. Potential spatial distribution of the newly introduced long-horned tick, Haemaphysalis longicornis in North America. Sci Rep. 2019;9:498.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  32. Rochlin I. Modeling the Asian Longhorned tick (Acari: Ixodidae) suitable habitat in North America. J Med Entomol. 2019;56:384–91.

    Article  PubMed  Google Scholar 

  33. Namgyal J, Couloigner I, Lysyk TJ, Dergousoff SJ, Cork SC. Comparison of habitat suitability models for Haemaphysalis longicornis neumann in North America to determine its potential geographic range. Int J Environ Res Public Health. 2020;17(21):8285. https://doi.org/10.3390/ijerph17218285.

    Article  PubMed  PubMed Central  Google Scholar 

  34. Hoogstraal H, Roberts FH, Kohls GM, Tipton VJ. Review of Haemaphysalis (kaiseriana) Longicornis Neumann (resurrected) of Australia, New Zealand, New Caledonia, Fiji, Japan, Korea, and Northeastern China and USSR, and its parthenogenetic and bisexual populations (Ixodoidea, Ixodidae). J Parasitol. 1968;54:1197–213.

    Article  CAS  PubMed  Google Scholar 

  35. Ben BC, Occi J, Bonilla DL, Egizi AM, Fonseca DM, Mertins JW, Backenson BP, Bajwa WI, Barbarin AM, Bertone MA, Brown J, Connally NP, Connell ND, Eisen RJ, Falco RC, James AM, Krell RK, Lahmers K, Lewis N, Little SE, Neault M, Pérez de León AA, Randall AR, Ruder MG, Saleh MN, Schappach BL, Schroeder BA, Seraphin LL, Wehtje M, Wormser GP, Yabsley MJ, Halperin W. Multistate infestation with the exotic disease-vector tick Haemaphysalis longicornis—United States, August 2017-September 2018. MMWR Morb Mortal Wkly Rep. 2018;67:1310–3.

    Article  Google Scholar 

  36. Tufts DM, Goodman LB, Benedict MC, Davis AD, VanAcker MC, Diuk-Wasser M. Association of the invasive Haemaphysalis longicornis tick with vertebrate hosts, other native tick vectors, and tick-borne pathogens in New York City, USA. Int J Parasitol. 2021;51:149–57.

    Article  CAS  PubMed  Google Scholar 

  37. Tufts DM, VanAcker MC, Fernandez MP, DeNicola A, Egizi A, Diuk-Wasser MA. Distribution, host-seeking phenology, and host and habitat associations of Haemaphysalis longicornis ticks, Staten Island, New York, USA. Emerg Infect Dis. 2019;25:792–6.

    Article  PubMed  PubMed Central  Google Scholar 

  38. Jones LD, Davies CR, Steele GM, Nuttall PA. A novel mode of arbovirus transmission involving a nonviremic host. Science. 1987;237:775–7.

    Article  CAS  PubMed  Google Scholar 

  39. Nonaka E, Ebel GD, Wearing HJ. Persistence of pathogens with short infectious periods in seasonal tick populations: the relative importance of three transmission routes. PLoS ONE. 2010;5:e11745.

    Article  PubMed  PubMed Central  Google Scholar 

  40. Rossi SL, Nasar F, Cardosa J, Mayer SV, Tesh RB, Hanley KA, Weaver SC, Vasilakis N. Genetic and phenotypic characterization of Sylvatic dengue virus type 4 strains. Virology. 2012;423:58–67.

    Article  CAS  PubMed  Google Scholar 

  41. Hermance ME, Thangamani S. Tick saliva enhances Powassan virus transmission to the host, influencing its dissemination and the course of disease. J Virol. 2015;89:7852–60.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  42. Hermance ME, Widen SG, Wood TG, Thangamani S. Ixodes scapularis salivary gland microRNAs are differentially expressed during Powassan virus transmission. Sci Rep. 2019;9:13110.

    Article  PubMed  PubMed Central  Google Scholar 

  43. Dupuis AP, Peters RJ, Prusinski MA, Falco RC, Ostfeld RS, Kramer LD. Isolation of deer tick virus (Powassan virus, lineage II) from Ixodes scapularis and detection of antibody in vertebrate hosts sampled in the Hudson Valley, New York State. Parasit Vectors. 2013;15:185.

    Article  Google Scholar 

  44. Labuda M, Kozuch O, Zuffová E, Elecková E, Hails RS, Nuttall PA. Tick-borne encephalitis virus transmission between ticks cofeeding on specific immune natural rodent hosts. Virology. 1997;235:138–43.

    Article  CAS  PubMed  Google Scholar 

  45. Labuda M, Nuttall PA, Kozuch O, Elecková E, Williams T, Zuffová E, Sabó A. Non-viraemic transmission of tick-borne encephalitis virus: a mechanism for arbovirus survival in nature. Experientia. 1993;49:802–5.

    Article  CAS  PubMed  Google Scholar 

  46. Randolph SE. Transmission of tick-borne pathogens between co-feeding ticks: Milan Labuda’s enduring paradigm. Ticks Tick Borne Dis. 2011;2:179–82.

    Article  PubMed  Google Scholar 

  47. Labuda M, Jones LD, Williams T, Danielova V, Nuttall PA. Efficient transmission of tick-borne encephalitis virus between cofeeding ticks. J Med Entomol. 1993;30:295–9.

    Article  CAS  PubMed  Google Scholar 

  48. Ličková M, Fumačová Havlíková S, Sláviková M, Slovák M, Drexler JF, Klempa B. Dermacentor reticulatus is a vector of tick-borne encephalitis virus. Ticks Tick Borne Dis. 2020;11:101414.

    Article  PubMed  Google Scholar 

  49. Labuda M, Austyn JM, Zuffova E, Kozuch O, Fuchsberger N, Lysy J, Nuttall PA. Importance of localized skin infection in tick-borne encephalitis virus transmission. Virology. 1996;219:357–66.

    Article  CAS  PubMed  Google Scholar 

  50. Randolph SE. Patterns of distribution of the tick Ixodes trianguliceps Birula on its hosts. J Anim Ecol. 1975;44:451.

    Article  Google Scholar 

  51. Schmidt KA, Ostfeld RS, Schauber EM. Infestation of Peromyscus leucopus and Tamias striatus by Ixodes scapularis (Acari: Ixodidae) in relation to the abundance of hosts and parasites. J Med Entomol. 1999;36:749–57.

    Article  CAS  PubMed  Google Scholar 

  52. Craine NG, Randolph SE, Nuttall PA. Seasonal variation in the role of grey squirrels as hosts of Ixodes ricinus, the tick vector of the Lyme disease spirochaete, in a British woodland. Folia Parasitol. 1995;42:73–80.

    CAS  Google Scholar 

  53. Xu Y, Wang J. The vector competence of Asian Longhorned ticks in Langat virus transmission. Viruses. 2024;16(2):304. https://doi.org/10.3390/v16020304.

    Article  PubMed  PubMed Central  Google Scholar 

  54. Costero A, Grayson MA. Experimental transmission of Powassan virus (Flaviviridae) by Ixodes scapularis ticks (Acari: Ixodidae). Am J Trop Med Hyg. 1996;55:536–46.

    Article  CAS  PubMed  Google Scholar 

  55. Lange RE, Prusinski MA, Dupuis AP, Ciota AT. Direct evidence of Powassan virus vertical transmission in Ixodes scapularis in nature. Viruses. 2024;16(3):456. https://doi.org/10.3390/v16030456.

    Article  PubMed  PubMed Central  Google Scholar 

Download references

Acknowledgement

We thank Michele Schuler, Leigh Ann Wiggins, and Hannah Bryant for assistance with animal experiments and animal husbandry. We also thank Ahmed Garba and Madeline Stone for assisting with RNA extractions, and Michael Levin for providing the initial H. longicornis ticks (distributed by BEI Resources, https://www.beiresources.org) that seeded our in-house colony of ticks used in this study.

Funding

This study was supported by the University of South Alabama College of Medicine and the National Institute of Allergy and Infectious Diseases, National Institutes of Health (award nos. R21AI163693, R01AI171398).

Author information

Authors and Affiliations

  1. Department of Microbiology and Immunology, Frederick P. Whiddon College of Medicine, University of South Alabama, Mobile, AL, USA

    Clemence Obellianne, Parker D. Norman, Eliane Esteves & Meghan E. Hermance

Authors
  1. Clemence Obellianne
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Parker D. Norman
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. Eliane Esteves
    View author publications

    You can also search for this author in PubMed Google Scholar

  4. Meghan E. Hermance
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

Conceptualization: MH; data curation: CO; formal analysis: CO, EE, and MH; funding acquisition: MH; methodology: CO, EE, PN, and MH; project administration: M.H.; supervision: M.H.; validation: C.O., E.E., and M.H.; writing (original draft): C.O.; and writing (review and editing): CO, EE, PN, and MH.

Corresponding author

Correspondence to Meghan E. Hermance.

Ethics declarations

Ethical approval and consent to participate

The animal study was reviewed and approved by University of South Alabama Institutional Animal Care and Use Committee.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interest.

Additional information

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary Information

13071_2024_6335_MOESM1_ESM.tif

Supplementary Material 1: Figure 1: Detection of POWV II RNA in donor I. scapularis females. POWV II RNA was detected in donor I. scapularis females by q-RT-PCR. Viral RNA was detected in 100% of virus-injected donor I. scapularis used in the co-feeding experiments. Viral RNA quantities are expressed as the number of NS5 gene fragment copies per ng of RNA after normalization to a standard curve. Note: The donor females were collected at varying stages of feeding (i.e., some were partially fed and others were fully engorged). Each cohort of donor females was maintained in ACL-3 facilities for ~ 1–4 weeks prior to processing and viral RNA extraction.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Obellianne, C., Norman, P.D., Esteves, E. et al. Interspecies co-feeding transmission of Powassan virus between a native tick, Ixodes scapularis, and the invasive East Asian tick, Haemaphysalis longicornis. Parasites Vectors 17, 259 (2024). https://doi.org/10.1186/s13071-024-06335-0

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s13071-024-06335-0

Keywords

Parasites & Vectors

ISSN: 1756-3305

Contact us