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Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks
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Original article
Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks Mirko Slovák a , Mária Kazimírová a , Marta Siebenstichová b , Katarína Ustaníková b , Boris Klempa b,c , Tamara Gritsun d , Ernest A. Gould e,f , Patricia A. Nuttall f,g,∗ a
Institute of Zoology, Slovak Academy of Sciences, Bratislava, Slovakia Institute of Virology, Slovak Academy of Sciences, Bratislava, Slovakia c Institute of Virology, Charité Medical School, Berlin, Germany d School of Biological Sciences, University of Reading, Reading, United Kingdom e Unité des Virus Emergents, Faculté de Médecine de Marseille, Aix-Marseille Universite, France f NERC Centre for Ecology & Hydrology, Wallingford, Oxfordshire, United Kingdom g Department of Zoology, University of Oxford, United Kingdom b
a r t i c l e
i n f o
Article history: Received 23 May 2014 Received in revised form 18 July 2014 Accepted 22 July 2014 Available online xxx Keywords: TBEV Ixodes ricinus Salivary glands Trans-stadial survival Co-feeding transmission
a b s t r a c t Biotic factors contributing to the survival of tick-borne viruses in nature are poorly understood. Using tickborne encephalitis virus (TBEV) and its principal European vector, Ixodes ricinus, we examined the relative roles of salivary gland infection, co-feeding transmission, and moulting in virus survival. Virus titres in the salivary glands increased after blood-feeding in a time- and dose-dependent manner. This was observed in ticks infected by inoculation but not in ticks infected by the natural route of co-feeding. Amplification of infection prevalence occurred via co-feeding. However, when larvae or nymphs subsequently moulted, the infection prevalence dramatically declined although this was not observed when ticks were infected by inoculation. Trans-stadial survival is a hitherto overlooked parameter that may contribute to the low incidence of TBEV infection in field-collected I. ricinus ticks. © 2014 Elsevier GmbH. All rights reserved.
Introduction Ticks are important vectors of at least 50 different virus species, including tick-borne encephalitis virus (TBEV), the most important human arboviral pathogen in Europe (Gritsun et al., 2003a,b; Nuttall, 2013). Taxonomically, TBEV is subdivided into three subtypes (Pletnev et al., 2011), the geographic distribution of which correlates with the distribution of their principal vector species, i.e., Ixodes persulcatus for Far-Eastern (TBEV-FE) and Siberian (TBEV-Sib) subtypes, and Ixodes ricinus for the western European (TBEV-Eur) subtype (Gritsun et al., 2003a). Although wild rodents are frequently referred to as the reservoirs for TBEV (e.g. Achazi et al., 2011; Tonteri et al., 2011), numerous data indicate that ticks are long-term reservoirs that have shaped the evolution and pathogenic properties of TBEV (Nuttall and Labuda, 2003; Kuno and Chang, 2005; Romanova et al., 2007).
∗ Corresponding author at: Department of Zoology, The Tinbergen Building, South Parks Road, Oxford OX1 3PS, United Kingdom. Tel.: +44 1865271167. E-mail addresses: [email protected], [email protected] (P.A. Nuttall).
Ixodes vectors of TBEV are characterised by a long life cycle that can take up to six years to complete (Gray, 1998). The virus may be maintained throughout the tick life cycle as it can infect all four developmental stages (eggs, larvae, nymphs, and adults). However, to do so TBEV must survive metamorphosis from larva to nymph and nymph to adult, which occurs after each stage has taken a bloodmeal and dropped off its vertebrate host. Trans-stadial survival is therefore critical for maintenance of the TBEV transmission cycle particularly as Ixodes vectors are 3-host ticks and there is usually a delay of months before each stage feeds and can transmit the ˇ virus (Rehᡠcek, 1965; Nuttall and Labuda, 2003). Although ticks become infected when feeding on viraemic animals, comparative estimates of the basic reproductive number, R0 (a measure of the ability of an infection to survive and spread in a population) indicate that the incidence and short duration of viraemia in typical rodent hosts are unlikely to ensure adequate maintenance of the TBEV transmission cycle (Hartemink et al., 2008). On the other hand, an important transmission mechanism for TBEV occurs between infected and uninfected ticks when they co-feed on susceptible hosts even when these hosts are immune to TBEV (Alekseev and Chunikhin, 1990a; Labuda et al., 1993a, 1997; Jones et al., 1997). This process, known as non-viraemic
http://dx.doi.org/10.1016/j.ttbdis.2014.07.019 1877-959X/© 2014 Elsevier GmbH. All rights reserved.
Please cite this article in press as: Slovák, M., et al., Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks. Ticks Tick-borne Dis. (2014), http://dx.doi.org/10.1016/j.ttbdis.2014.07.019
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transmission (NVT) because it does not depend on development of a patent viraemia, is considered a highly efficient mechanism for the transmission and long-term survival of TBEV in natural foci (Labuda et al., 1993b; Randolph, 2011). Estimates of R0 for TBEV indicate the virus would not survive in nature if it were not for co-feeding of ticks and NVT (Hartemink et al., 2008; Harrison and Bennett, 2012). Transovarial transmission also contributes to TBEV survival within the vector population. Field and experimental data indicate that the infection rate of I. ricinus larvae from a TBEV-positive egg mass is below 1% (Danielová and Holubová, 1991) and few larvae (<1%) are infected before feeding on viraemic hosts in nature (Danielová et al., 2002b). Nevertheless, a large proportion of tick larvae may become infected via NVT when they co-feed with infected nymphs (and/or infected larvae) on transmission-competent hosts (Labuda et al., 1993b; Randolph et al., 1999; Randolph, 2011). Transmission of tick-borne viruses from infected ticks to vertebrate hosts occurs via saliva produced by the infected tick salivary glands (Nuttall and Labuda, 2003). TBEV has been detected in tick saliva and in the cement cone that forms around the tick mouthparts during feeding (Chunikhin et al., 1988; Alekseev et al., 1996). During blood-feeding of ixodid ticks, extensive development of their salivary glands occurs. Moreover, 33–50% of all fluid ingested by the tick is excreted back to the host (Bowman et al., 2008). Thus, tick feeding involves alternation of blood ingestion and saliva secretion, providing an opportunity for recirculation of virus in and around the tick co-feeding sites. Furthermore, increased virus titres have been detected in feeding I. persulcatus (Alekseev and Chunikhin, 1990b) and I. ricinus (Khasnatinov et al., 2009; Belova et al., 2012). Reports of increased TBEV prevalence in ticks removed from humans and animals in comparison to questing ticks (Süss et al., 2004; Romanenko and Kondratyeva, 2011) support the concept that tick feeding affects TBEV infection in ticks. Research on TBEV has largely focused on molecular and pathogenic viral characteristics. By comparison, the biotic factors contributing to successful TBEV survival in natural foci are poorly understood. Therefore, in order to understand better the nature of these complex virus–vector–host interactions and to identify the primary biotic factors likely to be responsible for the survival of TBEV in its natural environment, we compared prevalence and levels of infection in salivary glands and in moulted ticks following infection by either virus inoculation or co-feeding transmission.
Materials and methods Ticks, animals and virus Adult Ixodes ricinus ticks were collected by blanket dragging the vegetation in localities in SW Slovakia known to be free of TBEV. The F1 generation of larvae, nymphs and adults, reared in the laboratory by feeding on uninfected laboratory rabbits, were used in the experiments. California rabbits (3 kg) were obtained from the Research Institute for Animal Production (Nitra, Slovakia). Balb/c (20 g) used in virus transmission experiments and ICR mice (25 g) used in tickfeeding experiments were bred at the Institute of Experimental Pharmacology and Toxicology (Dobrá Voda, Slovakia). Rearing of ticks was carried out as described in Slovák et al. (2002). The prototype Czech TBEV strain Hypr (accession number U39292) prepared as a 10% mouse brain suspension of 1.1 × 109 PFU/ml in Leibovitz’s L-15 medium was used for inoculation of ticks. The usage of animals in the experiments was approved by the State Veterinary and Food Administration of the Slovak Republic (permit numbers 2362/06221, 928/10-221 and1335/12-221). Euthanasia of the animals at the end of the experiments was by cervical dislocation under deep anaesthesia induced using carbon dioxide.
Virus inoculation of ticks and partial feeding Unfed female ticks were inoculated with a defined concentration of 0.5 l of TBEV suspension into the coxal plate of the second pair of legs under a photomacroscope (Wild M 400, Wild Heerbrugg AG, Switzerland), using a digital microinjectorTM system (MINJ-D-CE, Tritech Research, Inc., USA), as previously described (Khasnatinov et al., 2009). Inoculated female ticks were incubated at 22 ± 1 ◦ C and 85–90% RH for a maximum of 120 days. Immediately after inoculation and at defined periods of incubation (dpi = days post inoculation), a cohort of the inoculated ticks was removed from the incubation chamber and placed in retaining chambers on ICR mice, as previously described (Labuda et al., 1993a). Five inoculated female ticks together with 3 uninfected male ticks (to facilitate female tick feeding) were placed within the retaining chamber on each mouse. The tick-infested mice were retained for 2 or 3 days, which is sufficient time for the inoculated ticks to attach and commence feeding but insufficient time for the female ticks to complete engorgement (requiring 6–10 days). Unfed I. ricinus nymphs were inoculated with 0.1 l of TBEV suspension (1.1 × 109 PFU/ml) into the coxal plate of the second pair of legs in the same manner as described for tick females. Inoculated nymphs were incubated at 22 ± 1 ◦ C and 85–90% RH for 16 or 30 days and then fed on mice. The engorged nymphs either were assayed for virus immediately after drop off or they were incubated and allowed to moult to adults. A cohort of moulted females was fed on mice for 3 days; the rest of the females were assayed for virus without being fed.
Co-feeding transmission and trans-stadial survival of the virus The model of co-feeding transmission was carried out essentially as described previously (Labuda et al., 1993a). Briefly, Balb/c mice were infested with two virus-inoculated I. ricinus females 14 dpi, two uninfected males and either 20 nymphs or approximately 100 larvae placed together in one retention chamber at the same time. Following 1, 2 and 3 days of co-feeding, nymphs were subsequently removed from three of the experimental mice. Of the partially fed cohorts of nymphs, 50% were dissected and the presence of TBEV was tested separately in their salivary glands (SG) and body carcasses (remains of the tick after removal of SG), whereas the remaining nymphs were tested without dissection. The rate of co-feeding virus transmission was determined as % of positive nymphs. Infectivity of co-feeding female ticks was confirmed by plaque assay of their SG. To study trans-stadial survival, fully engorged larvae and nymphs that had dropped off the host were allowed to moult. At various time intervals following moulting to nymphs and adults, unfed and fed ticks were dissected and their SG and body carcasses were tested by infectivity assays for the presence of TBEV.
Tick dissection The mice on which ticks fed were killed and ticks were gently removed with fine forceps. Both unfed and partially fed ticks were washed in distilled water and then in sterile 0.15 M NaCl (0.9%) with antibiotics (5% Penicillin/Streptomycin mixture). SG were dissected out individually under a stereomicroscope in ice-cold sterile 0.15 M NaCl, rinsed twice in sterile 0.15 M NaCl to minimise contamination with haemolymph and virus inoculum, and stored individually in Eppendorf tubes containing 50 l of sterile 0.15 M NaCl, at −80 ◦ C. Carcasses and undissected nymphs were individually stored in Eppendorf tubes at −80 ◦ C.
Please cite this article in press as: Slovák, M., et al., Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks. Ticks Tick-borne Dis. (2014), http://dx.doi.org/10.1016/j.ttbdis.2014.07.019
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Fig. 1. Dynamics of TBEV infection in I. ricinus salivary glands. Proportion (%) of infected ticks and virus titres in SG of fasting or 2–3 days-fed adult female ticks at different times following inoculation with 5.5 × 104 PFU of TBEV. Error bars indicate SEM. Asterisks denote significant differences in mean values for partially fed ticks compared with unfed ticks at corresponding days post inoculation: ***P < 0.001, *P < 0.05 (Kolgomorov–Smirnov test). The complete dataset is presented in Table S1 (supplementary file).
Plaque assay Virus presence and infectivity titres were determined using plaque assays, which were performed on Porcine kidney stable (PS) cell monolayers. Prior to assays, SG were homogenised individually in 50 l sterile 0.15 M NaCl and the volumes were completed to 500 l with culture medium. Tick carcasses and whole nymphs were thawed and homogenised in 500 l culture medium. Samples were centrifuged at 2000 rpm for 10 min at 4 ◦ C and the supernatants were kept. PS cells were seeded in 24-well plates (2.5–3 × 105 cells/well) in medium comprising inactivated 7% foetal bovine serum (FBS) solution, Medium 199 [a combination of Medium 199 in Hank’s buffer (150 ml) and Medium 199 in Earle’s buffer (350 ml)] and Penicillin/Streptomycin. 100 l of undiluted or serially diluted tick samples were used to infect the PS cells. After incubation of the virus with cells for 4 h at 37 ◦ C in 5% CO2 , Medium 199 containing 2% carboxymethyl cellulose (CMC) (1:1) and buffered with sodium bicarbonate to pH 7.0–7.5 was added and plates were then incubated for 4 days at 37 ◦ C in 5% CO2 . Thereafter, plates were fixed with 4% formalin and plaques were detected by staining with 0.3 ml of crystal violet for 3 min. After staining, plates were washed using tap water and plaques were counted. Virus titres were expressed as log10 PFU/tick. Statistical analysis Statistical analyses (Kolmogorov–Smirnov test, regression analysis) were performed on the raw data (PFU/tick, ignoring zero values). Descriptive statistics were calculated based on logtransformed data, using Statgraphics Plus 5.1.software. Chi-square statistics was used to compare trans-stadial survival rates in nymphs after co-feeding and parenteral inoculation.
3
Fig. 2. Ratio of the total amount of infectious virus detected in SG of partially fed I. ricinus adult female ticks compared with unfed ticks at different days after inoculation (dpi).
decreased until 120 dpi when the mean titre was 2.1 log10 PFU/SG and monitoring was discontinued. The cohort of fasting ticks that were allowed to feed for 2–3 days at different times following inoculation also showed an increase in the number of ticks infected and in the SG virus titres, with increasing dpi (Fig. 1, Table S1B). All of the ticks in this cohort had infected SG by 7 dpi (4 days fasting + 3 days feeding) and the mean maximum titre in SG of 4.3 log10 PFU/SG was observed 24 dpi (21 days fasting + 3 days feeding). Comparison of the two tick cohorts revealed that, as measured by plaque assay, partial feeding increased the rate at which infection of the SG was detected, the rate of virus replication in the SG, and enhanced the total amount of infectious virus measured in the SG (Fig. S1). The amplification in SG virus yield was greatest when inoculated ticks were immediately fed on mice for 2 days (total dpi = 2) and then reduced over an extended period until ∼120 dpi when blood-feeding did not appear to affect the virus yield from SG (Fig. 2). To examine whether the dose of virus inoculated into ticks affected the enhancing effect of partial feeding, five groups of female ticks were each inoculated with 10-fold differences in virus dose. At 14 dpi, 50% ticks in each group were fed on mice for 3 days. All of the unfed ticks inoculated with TBEV doses >55 PFU per tick were positive (Fig. 3, Table S2). With lower doses of inoculated virus, the percentage of positive ticks detected was reduced but remained high (75–94%). In contrast, 100% of partially fed ticks, inoculated with virus, were positive irrespective of the virus dose. Apart from the highest dose in unfed ticks, virus titres in SG of both unfed and partially fed ticks declined with decreasing doses: in unfed ticks from 3.7 to 2.0 log10 PFU/SG and in ticks fed for 3 days, from 3.3 to 2.8 log10 PFU/SG. When virus titres in fasting ticks were
Results Dynamics of virus replication in tick salivary glands An estimated 5.5 × 104 PFU of TBEV were inoculated into the haemocoel of I. ricinus females and the infection rate and TBEV titres in SG were determined at different time intervals in fasting and fasting + feeding ticks as shown in Fig. 1 and Table S1 (see supplementary file). In the cohort of inoculated unfed ticks, infectious TBEV was detected in 25% of the SG at 2 dpi and 100% of SG were infected by 14 dpi (Fig. 1, Table S1A). The highest average titres of 2.9 log10 PFU/SG were recorded at 21 dpi after which titres gradually
Fig. 3. Dose-dependence of TBEV replication in I. ricinus. Proportion (%) of infected ticks and virus titres in SG of fasting or 3 days-fed adult female ticks, 14 days following inoculation with different doses of TBEV. Error bars indicate SEM values of virus titres as presented in Table S2 (supplementary file).
Please cite this article in press as: Slovák, M., et al., Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks. Ticks Tick-borne Dis. (2014), http://dx.doi.org/10.1016/j.ttbdis.2014.07.019
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Fig. 4. Comparative dynamics of TBEV transmission to I. ricinus nymphs during cofeeding with infected female ticks for either 1, 2 or 3 days.
plotted as a function of inoculation dose (ignoring zero values), the regression was significant (P < 0.001). The regression formula was: Y = 1.68x + 1403 (R2 = 0.55, F = 19.44, n = 46). Significant regression in fed ticks was not detected (data not shown). Dynamics of co-feeding transmission Groups of two I. ricinus females, 14 dpi, were placed together with 20 uninfected nymphs in a retention chamber on the shaven skin of mice and allowed to co-feed for 1–3 days as described in Section “Materials and methods”. The rate of TBEV transmission was expressed as the percentage of TBEV-positive nymphs (Table 1). Maximum co-feeding transmission (61%) was recorded on day 3. The dynamics of TBEV accumulation in the SG and body carcass of nymphs compared with whole intact nymphs are presented in Fig. 4. During the 3-day co-feeding period, no virus was detected in the SG of nymphs, only in the remaining body carcasses and in whole intact nymphs. Virus titres in body carcasses and whole intact bodies of nymphs were 1.7–2.0 and 2.0–2.3 log10 PFU/tick, respectively. Dynamics of trans-stadial survival In experiments to study trans-stadial survival, uninfected nymphs were allowed to co-feed with infected adults until repletion of the nymphs (usually within 3–5 days of co-feeding). The replete nymphs were then allowed to moult into adults and the percentage of positive female ticks was determined at different time intervals during 90 days post-moult; cohorts were allowed to feed for 3 days to determine whether feeding amplified virus infection. To trace TBEV traffic inside the moulted female ticks, their SG and body carcass were investigated (Table 2). Of the 82 female ticks examined, 7 (8.5%) showed evidence of virus infection. Based on a 61% transmission rate (Table 1), the expected level of infection was 50/82 if 100% infections survived moulting. Hence the observed level of 8.5% indicates a trans-stadial survival rate of
14%. Infectious virus was detected in the SG of only two of these females, both of which also had virus positive carcasses. These two ticks with infected SG had the highest virus titres in their carcasses. Partial feeding of the adults did not increase the number of virus positive ticks or the virus titres. No infected ticks were detected after 60 days post-moult and infections in SG were not detected after 14 days post-moult. Trans-stadial survival was also examined using larvae. Uninfected larvae were allowed to co-feed with infected adults until repletion and drop off. The engorged larvae were then allowed to moult to nymphs. At various times after moulting, the nymphs were assayed for virus either as whole undissected ticks or following dissection as SG and carcass. Cohorts were partially fed on uninfected mice for 2 days to determine whether feeding amplified virus infection following moulting. Only one of the 129 nymphs that moulted from engorged larvae was positive for infectious virus. This infected nymph, examined 14 days after moulting, had a remarkably high virus titre in its SG, which was higher than the titre in its body carcass (Table 3). As was the case for moulted adults, partial feeding of moulted nymphs did not affect the prevalence or incidence of infection. For comparison with the natural route of co-feeding infection, nymphs were inoculated with virus and then allowed to feed to repletion, drop off and then moult to adults to determine transstadial survival. Two different treatments were examined: feeding nymphs at 16 dpi or at 30 dpi. In the first experiment, 23 nymphs were inoculated with an estimated 1.1 × 105 PFU TBEV of which 15 (65.2%) survived. At 16 dpi, the nymphs were allowed to feed on Balb/c mice; 12 nymphs completed engorgement. Five of the replete intact nymphs were immediately assayed for virus; all were infected (titres >5.0 log10 PFU/tick). The remaining engorged nymphs moulted into 6 female and 1 male adults; 2 of the females were allowed to feed for 2 days, 45–50 days after moulting. In the second experiment, 80 nymphs were inoculated with the same virus dose; 26 (32.5%) survived. At 30 dpi, the nymphs were allowed to feed on Balb/c mice; 25 nymphs completed engorgement. Ten of the replete nymphs were immediately dissected and then assayed for virus. Of these, 9 had infected carcasses (titres 1.3–3.3 log10 PFU) while 5 had infected SG (titres 1.5–3.1 log10 PFU); the infection prevalence was 90%. The remaining engorged nymphs moulted into 12 female and 3 male adults; 7 of the females were allowed to feed for 2 days. At 45–50 days after moulting, the adult ticks were all assayed for virus either following dissection into SG + carcass (females) or as intact (whole body) ticks (males) (Table 4). Following moulting, 19/22 (86.4%) adults were infected (100% unfed and 66.7% fed; 83.3% females and 100% males). There was no obvious difference in results obtained following feeding of nymphs either 16 or 30 dpi, or between unfed and partially fed adults, although numbers were comparatively low as a result of failure of nymphs to survive the trauma of inoculation. However, as observed following co-feeding infection, all female ticks that had infected SG also had an infected body carcass, and the infection prevalence in body carcasses (mean 83.3%) was greater than that of SG (mean 22.2%).
Table 1 Comparative dynamics of TBEV transmission to I. ricinus nymphs during co-feeding with infected female ticks. Days of co-feeding
1 2 3 a
SG
Body carcass
No. positive/total (%)
No. positive/total (%)
Mean ± SEM (log10 PFU/tick)
No. positive/total (%)
Mean ± SEM (log10 PFU/tick)
No. positive/total (%)
0/6 (0%) 0/6 (0%) 0/7 (0%)
0/6 (0%) 1/6 (17%) 4/7 (57%)
0 2.0 1.72 ± 0.29
1/6 (17%) 2/6 (33%) 4/6 (67%)
1.95 2.12 ± 0.42 2.25 ± 0.32
1/12 (8%) 3/12 (25%) 8/13 (61%)
Whole body
Total no. ticks examineda
Infectious virus was measured in homogenates of either the SG and remaining body carcass of a tick or the whole body of a tick.
Please cite this article in press as: Slovák, M., et al., Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks. Ticks Tick-borne Dis. (2014), http://dx.doi.org/10.1016/j.ttbdis.2014.07.019
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Table 2 Trans-stadial survival of TBEV from I. ricinus nymphs to females following co-feeding infection of the nymphs. Sample assayed for virus
Days after moulting (+3 days of feeding) 1
7
14
11 + 3
SG Observed Positive % positive Titrea
6 0 0 0
6 1 17 2.7
14 1 7.14 5.0
14 0 0 0
Carcass Observed Positive % positive Titrea
6 1 17 3.1
6 1 17 3.5
14 1 7.14 5.1
Total ticks Observed Positive % positiveb
6 1 17
6 1 17
14 1 7.1
Total samples 30
27 + 3
60
57 + 3
90
87 + 3
6 0 0 0
8 0 0 0
6 0 0 0
8 0 0 0
8 0 NT NT
6 0 NT NT
82 2 2.4
14 1 7.14 2.6
6 1 17 2.9
8 1 17 3.5
6 1 17 3.2
8 0 0 0
8 0 0 0
6 0 0 0
82 7 8.5
14 1 7.1
6 1 17
8 1 17
6 1 17
8 0 0
8 0 0
6 0 0
82 7 8.5
a Virus titre (log10 PFU/tick) in SG and carcass (tick remains after removal of SG) of unfed or partially fed females at different days after moulting from nymphs that had co-fed with infected females. NT is not tested. b Based on 100% infected nymphs: see text.
Discussion
Studies on TBEV replication in ticks, in which ticks are inoculated parenterally with virus, have generally placed the inoculated ticks onto susceptible animals at a set time after inoculation and allowed the ticks to complete engorgement. For example, female I. ricinus inoculated with 3.7 log10 PFU TBEV were either maintained fasting or placed on experimental mice 15 h after inoculation and then cohorts of the feeding ticks were examined at 8 h, 15 h, 38 h, 96 h post-inoculation. Virus replication in whole feeding ticks was faster and reached higher titres than in fasting ticks at the same times post-inoculation (Belova et al., 2012). The role of the SG in this blood-feeding induced amplification of virus infectivity could not be determined. We took a different approach, taking ticks at different times following inoculation, and then allowing the inoculated fasting ticks to feed only partially. This approach enabled us to take a snapshot of the virus infectivity in the infected SG at different times post-inoculation and after prolonged fasting, reflecting the situation in nature when infected ticks may fast for months and possibly >1 year before feeding. We observed that, when the infected female ticks took a bloodmeal, the level of virus infectivity increased rapidly by at least 3 orders of magnitude. This
This study focuses on three biotic factors involved in the transmission cycle of TBEV by its European vector, Ixodes ricinus: (1) infection of the tick salivary glands, (2) co-feeding transmission, and (3) trans-stadial survival. The first objective was to study the dynamics of infection of the tick salivary glands to understand the significance of their role in long-term TBEV survival and transmission. Our model of inoculation of the virus into the tick haemocoel circumvents the requirement for the virus to cross the tick midgut barrier (Nuttall and Labuda, 2003), thereby providing unimpeded access for the virus to infect the tick SG. This appeared to be a successful strategy as TBEV showed high tropism to I. ricinus SG and virus amplification was detected in SG of a high proportion of inoculated ticks, although the mechanism of virus transfer from the haemocoel to tick SG and saliva is unknown (Kaufman and Nuttall, 1996). Infectious virus was recovered from SG for at least 120 days following its initial inoculation, demonstrating the potential importance of the SG as a long-term reservoir for efficient transmission of the virus.
Table 3 Trans-stadial transmission of TBEV from I. ricinus larvae to nymphs following co-feeding infection of the larvae. Sample assayed for virus
Days after moulting (+2 days of feeding) 1
7
14
12+
Total samples 30
28+
58+
90
88+
8 0 0 0
8 0 0 0
NT – – –
NT NT NT NT
57 1 1.8
6 0 0 0
8 0 0 0
8 0 0 0
– – – –
NT NT NT NT
57 1 1.8
18 0 0
5 0 0
6 0 0
6 0 0
6 0 0
6 0 0
72 0 0
24 0 0
11 0 0
14 0 0
14 0 0
6 0 0
6 0 0
SG Observed Positive % positive Titrea
8 0 0 0
6 1 17 5.7
9 0 0 0
6 0 0 0
6 0 0 0
6 0 0 0
Carcass Observed Positive % positive Titrea
8 0 0 0
6 1 17 3.5
9 0 0 0
6 0 0 0
6 0 0 0
Whole body Observed Positive % positive
6 0 0
3 0 0
8 0 0
8 0 0
17 0 0
14 0 0
Total ticks Observed Positive %
14 0 0
9 1 11.1
60
129 1 0.8%
a Virus titre (log10 PFU/tick) in SG, carcass (tick remains after removal of SG) or whole undissected tick of unfed or partially fed nymphs at different days after moulting from larvae that had co-fed with infected females. NT is not tested.
Please cite this article in press as: Slovák, M., et al., Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks. Ticks Tick-borne Dis. (2014), http://dx.doi.org/10.1016/j.ttbdis.2014.07.019
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Table 4 Trans-stadial survival of TBEV from I. ricinus nymphs to adults following virus inoculation of the nymphs. Sample assayed for virus
Moulted from 16 dpi nymph Female Unfed
+3
SG Observed Positive % positive Titrea (min–max)
4 1 25% 1.30
2 0 0%
Carcass Observed Positive % positive Titrea (min–max)
4 4 100% 2.02 (1.74–2.70)
2 2 100% 2.13 (1.48–2.78)
NT
NT
Whole body Observed Positive % positive Titrea (min–max) Total ticks Observed Positive % positive a
Moulted from 30 dpi nymph Male
Female
Unfed
+3
NT
NT
NT
Unfed
+3
5 1 20% 2.65
7 2 28.6% 2.84 (1.78–3.9)
5 5 100% 1.55 (1.0–2.02)
7 4 57.1% 1.90 (1.0–3.15)
NT
NT
NT
NT
Total samples Male Unfed
+3
NT
NT 18 4 22.2%
NT
1 1 100% >2.00
18 15 83.3% NT 3 3 100% 2.48 (1.18–3.34)
NT 4 4 100%
2 2 100%
NT
1 1 100%
4 4 100% NT
5 5 100%
7 4 57.1%
3 3 100%
22 19 86.4%
Mean virus titre (log10 PFU/tick) in SG, carcass (tick remains after removal of SG) or whole undissected tick. NT, not tested.
feeding-induced amplification of virus infectivity in SG persisted for at least 87 dpi. Thus virus replication in SG may play an important role in maximising the dose of virus transmitted from an infected tick to an uninfected vertebrate host during the prolonged feeding period (6–10 days for female I. ricinus). Feeding-induced enhancement of SG infection may explain reports of significantly higher infection prevalences of TBEV in partially fed ticks removed from humans compared with field-collected ticks from the same locality (summarised in Supplementary Table S1 of Lindblom et al., 2014). The susceptibility of infected SG to blood-feeding induced amplification of the infection appeared to persist for much of the fasting period. The reason for the apparent loss in responsiveness of SG infection by 120 dpi is unknown. Possibly the SG become refractory to virus amplification after long periods off the host as a means of protecting the SG, which play a crucial role in water balance during fasting periods (Bowman et al., 2008). Based on virus inoculation studies and partial feeding at selected times following inoculation, it appears the tick SG are both an important reservoir for TBEV survival during fasting and for amplification of the potential virus dose transmitted to a vertebrate host during blood-feeding. However, these observations based on parenteral inoculation of TBEV were not reproduced in subsequent studies involving the natural route of co-feeding infection of ticks. For co-feeding transmission, uninfected nymphs were allowed to feed with TBEV-inoculated I. ricinus females for 1–3 days, on localised skin sites of mice. The experiments demonstrated comparatively high rates of co-feeding transmission with up to ∼60% of nymphs infected, supporting previous observations that co-feeding is a highly efficient mode of virus transmission. In previous studies, the rate of engorged I. ricinus nymphs acquiring TBEV during co-feeding in close proximity to infected females on laboratory hosts varied between 45% on guinea pigs (Labuda et al., 1993a) and 93% on Balb/c mice (Labuda et al., 1996), and for natural hosts, between 10% on hedgehogs and up to 80% on yellow-necked mice (Apodemus flavicollis) (Labuda et al., 1993b, 1997). Transmission efficiency of different European strains of TBEV for nymphs cofeeding with infected adults on Balb/c mice varied between 93% for strain 198 (isolated from I. ricinus in Slovakia) (Labuda et al., 1996) and 64% for the Hypr strain (Kazimírová et al., 2012) and 61% (this study).
When I. ricinus nymphs co-fed with infected females, the percentage of nymphs acquiring the virus increased during co-feeding. However, after 3 days of co-feeding, infectious virus was detected only in their bodies and not in their SG. In previous studies aimed to understand the role of local skin infection and saliva assisted transmission in non-viraemic co-feeding transmission, the percentage of I. ricinus nymphs acquiring TBEV increased in parallel with the duration of co-feeding on natural rodent hosts as well as on laboratory mice from day 1 to day 3 (Labuda et al., 1996). Similar results were achieved by infestation of yellow-necked mice with infected I. ricinus females and short-feeding Haemaphysalis inermis nymphs (Labuda et al., unpublished). However, in all these studies only whole bodies of co-feeding nymphs were screened for the presence of TBEV, and not the SG and body carcasses. The consistently higher incidence of infection and virus titre of whole bodies compared with the sum of the virus yields from SG + carcass is most likely accounted for by infectious virus lost in the haemolymph during the tick dissection as TBEV has been detected in tick haemocytes ˇ (Rehᡠcek and Mrenová, 1966). If this is the case, the presence of virus in the haemolymph indicates that virus acquired within the bloodmeal is able to escape through the tick midgut wall. The estimated mean titre of virus in the haemolymph was approximately 125 PFU/tick which, compared with parenteral inoculation of adult females with different doses of virus (Table 3), should have been sufficient to infect the SG of most of the nymphs (assuming the SG of nymphs show similar susceptibility and infection dynamics to those of adult I. ricinus). Possibly the difference is due to the dynamics of infection of SG by inoculated virus compared with naturally acquired virus. In the case of parenteral inoculation, free extracellular virus can directly access the SG whereas virus that escapes from the midgut may infect haemocytes and/or other tissues and be unable to invade the SG directly. The observation that partial feeding of moulted nymphs and moulted larvae did not increase the prevalence of infection in the co-fed tick population is surprising, given that the virus titres in the carcasses of the infected co-fed ticks were comparable to those observed in the ticks infected by inoculation. Moreover, only 2/7 infected ticks had infectious virus detectable in their SG (Table 4). Again, this is surprising given the speed at which TBEV appeared to infect SG following virus inoculation into the haemocoel. The
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observations suggest there may be a midgut and/or salivary gland constraint on TBEV infection and/or innate immune response that is circumvented by virus inoculation into the haemocoel. For example, if virus acquired in the bloodmeal fails to escape the midgut prior to moulting, it may be destroyed. Following 3 days co-feeding of nymphs with infected adults, 61% nymphs became infected. However, the infection prevalence after moulting to adults was only 8.5%, indicating a maximal transstadial survival rate of 14% (assuming a minimal 61% infection prevalence of the engorged nymphs prior to moulting). By contrast, the trans-stadial survival rate following moulting of engorged nymphs infected by inoculation was 92.6% (based on an infection prevalence of 93.3% for the engorged nymphs), significantly higher than the maximal 14% trans-stadial survival rate for nymphs infected by co-feeding (Chi-square = 56.03; P < 0.01). The estimated trans-stadial survival rate through moulting from larva to nymph following co-feeding infection of larvae was 0.8% assuming all engorged larvae were infected. Virus infection of larvae by inoculation technically was not possible so we were unable to compare the trans-stadial survival rates following co-feeding with those following infection by inoculation. We are unaware of any comparable studies on trans-stadial survival following co-feeding infection of ticks. Trans-stadial survival rates for engorged larvae and engorged nymphs of 9.5% and 54%, respectively, were reported for Powassan virus (a member of the TBEV complex) following infection by feeding on viraemic hamsters (Costero and Grayson, 1996). Nevertheless, our data correspond with epidemiological data that reveal a low TBEV-Eur prevalence in questing I. ricinus caught in nature, which is generally between 0.1 and 1% infectivity over different areas and years of investigation, and mainly reflects the infection prevalence in unfed nymphs (e.g. Danielová et al., 2002a; Carpi et al., 2009; Lommano et al., 2012). At this stage of the research, we do not know if the low rate of trans-stadial survival is a general phenomenon for TBEV infection of I. ricinus. The prototype TBEV strain Hypr has been adapted to laboratory mice via serial passages and in contrast with many wild-type TBEV isolates the 3 untranslated region (3 UTR) has a deletion of 263 nucleotides (e.g. Wallner et al., 1995; Hayasaka et al., 2001) which is believed to be a replication enhancer region of the 3 UTR (Gritsun and Gould, 2007a,b). Virus genetic characteristics encoded in 3 UTR might be essential for the virus to reach a critical replication rate and also to circumvent the tick immune responses (Pijlman et al., 2008; Gritsun et al., 2014) thus contributing to the efficiency of TBEV dissemination and amplification in ticks and transmission to co-feeding ticks and hosts. In conclusion, although our data support modelling predictions that co-feeding (non-viraemic) transmission of TBEV is essential for survival of the TBEV in nature, they raise questions about the parameters used to estimate the basic reproductive number (R0 ) for TBEV. In particular, it is generally assumed that the infection in ticks survives from one developmental stage to the next and does not need to be parameterised (Hartemink et al., 2008; Harrison and Bennett, 2012). Given our observations, the loss of an infection during moulting could have a highly significant effect on estimates of R0 . However, it remains to be determined if strain Hypr is an exception to the rule for TBEV survival. This is currently being investigated using infectious clones of the viruses.
Acknowledgements This paper is dedicated to the memory of Milan Labuda, whose idea it originally was. The authors thank him for everything he taught them. The technical assistance of Mrs Zuzana Guláˇsová is highly acknowledged. The work was supported by Slovak Scientific Grant Agency, VEGA (grant no. 2/0191/12), European
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Commission (European Virus Archive, FP7 CAPACITIES project – GA no. 228292), grant agency APVV (no. APVV DO7RP–0014–11), EU grant FP7-261504 EDENext and is catalogued by the EDENext Steering Committee as EDENext240 (http://www.edenext.eu), BBSRC UK grant BBS/B/00697, MRC UK grant G0801208 and Royal Society UK grant JP100471. E A Gould is funded by EU FP7 grant no. 260644–SILVER. The contents of this publication are the sole responsibility of the authors and do not necessarily reflect the views of the European Commission. Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:10.1016/j.ttbdis.2014.07.019. References Achazi, K., Ruzek, D., Donoso Mantke, O., Schlegel, M., Ali, H.S., Wenk, M., SchmidtChanasit, J., Ohlmeyer, L., Ruhe, F., Vor, T., Kiffner, C., Kallies, R., Ulrich, R.G., Niedrig, M., 2011. Rodents as sentinels for the prevalence of tick-borne encephalitis virus. Vector-Borne Zoonotic Dis. 11, 641–647. 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Original article
Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks Mirko Slovák a , Mária Kazimírová a , Marta Siebenstichová b , Katarína Ustaníková b , Boris Klempa b,c , Tamara Gritsun d , Ernest A. Gould e,f , Patricia A. Nuttall f,g,∗ a
Institute of Zoology, Slovak Academy of Sciences, Bratislava, Slovakia Institute of Virology, Slovak Academy of Sciences, Bratislava, Slovakia c Institute of Virology, Charité Medical School, Berlin, Germany d School of Biological Sciences, University of Reading, Reading, United Kingdom e Unité des Virus Emergents, Faculté de Médecine de Marseille, Aix-Marseille Universite, France f NERC Centre for Ecology & Hydrology, Wallingford, Oxfordshire, United Kingdom g Department of Zoology, University of Oxford, United Kingdom b
a r t i c l e
i n f o
Article history: Received 23 May 2014 Received in revised form 18 July 2014 Accepted 22 July 2014 Available online xxx Keywords: TBEV Ixodes ricinus Salivary glands Trans-stadial survival Co-feeding transmission
a b s t r a c t Biotic factors contributing to the survival of tick-borne viruses in nature are poorly understood. Using tickborne encephalitis virus (TBEV) and its principal European vector, Ixodes ricinus, we examined the relative roles of salivary gland infection, co-feeding transmission, and moulting in virus survival. Virus titres in the salivary glands increased after blood-feeding in a time- and dose-dependent manner. This was observed in ticks infected by inoculation but not in ticks infected by the natural route of co-feeding. Amplification of infection prevalence occurred via co-feeding. However, when larvae or nymphs subsequently moulted, the infection prevalence dramatically declined although this was not observed when ticks were infected by inoculation. Trans-stadial survival is a hitherto overlooked parameter that may contribute to the low incidence of TBEV infection in field-collected I. ricinus ticks. © 2014 Elsevier GmbH. All rights reserved.
Introduction Ticks are important vectors of at least 50 different virus species, including tick-borne encephalitis virus (TBEV), the most important human arboviral pathogen in Europe (Gritsun et al., 2003a,b; Nuttall, 2013). Taxonomically, TBEV is subdivided into three subtypes (Pletnev et al., 2011), the geographic distribution of which correlates with the distribution of their principal vector species, i.e., Ixodes persulcatus for Far-Eastern (TBEV-FE) and Siberian (TBEV-Sib) subtypes, and Ixodes ricinus for the western European (TBEV-Eur) subtype (Gritsun et al., 2003a). Although wild rodents are frequently referred to as the reservoirs for TBEV (e.g. Achazi et al., 2011; Tonteri et al., 2011), numerous data indicate that ticks are long-term reservoirs that have shaped the evolution and pathogenic properties of TBEV (Nuttall and Labuda, 2003; Kuno and Chang, 2005; Romanova et al., 2007).
∗ Corresponding author at: Department of Zoology, The Tinbergen Building, South Parks Road, Oxford OX1 3PS, United Kingdom. Tel.: +44 1865271167. E-mail addresses: [email protected], [email protected] (P.A. Nuttall).
Ixodes vectors of TBEV are characterised by a long life cycle that can take up to six years to complete (Gray, 1998). The virus may be maintained throughout the tick life cycle as it can infect all four developmental stages (eggs, larvae, nymphs, and adults). However, to do so TBEV must survive metamorphosis from larva to nymph and nymph to adult, which occurs after each stage has taken a bloodmeal and dropped off its vertebrate host. Trans-stadial survival is therefore critical for maintenance of the TBEV transmission cycle particularly as Ixodes vectors are 3-host ticks and there is usually a delay of months before each stage feeds and can transmit the ˇ virus (Rehᡠcek, 1965; Nuttall and Labuda, 2003). Although ticks become infected when feeding on viraemic animals, comparative estimates of the basic reproductive number, R0 (a measure of the ability of an infection to survive and spread in a population) indicate that the incidence and short duration of viraemia in typical rodent hosts are unlikely to ensure adequate maintenance of the TBEV transmission cycle (Hartemink et al., 2008). On the other hand, an important transmission mechanism for TBEV occurs between infected and uninfected ticks when they co-feed on susceptible hosts even when these hosts are immune to TBEV (Alekseev and Chunikhin, 1990a; Labuda et al., 1993a, 1997; Jones et al., 1997). This process, known as non-viraemic
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transmission (NVT) because it does not depend on development of a patent viraemia, is considered a highly efficient mechanism for the transmission and long-term survival of TBEV in natural foci (Labuda et al., 1993b; Randolph, 2011). Estimates of R0 for TBEV indicate the virus would not survive in nature if it were not for co-feeding of ticks and NVT (Hartemink et al., 2008; Harrison and Bennett, 2012). Transovarial transmission also contributes to TBEV survival within the vector population. Field and experimental data indicate that the infection rate of I. ricinus larvae from a TBEV-positive egg mass is below 1% (Danielová and Holubová, 1991) and few larvae (<1%) are infected before feeding on viraemic hosts in nature (Danielová et al., 2002b). Nevertheless, a large proportion of tick larvae may become infected via NVT when they co-feed with infected nymphs (and/or infected larvae) on transmission-competent hosts (Labuda et al., 1993b; Randolph et al., 1999; Randolph, 2011). Transmission of tick-borne viruses from infected ticks to vertebrate hosts occurs via saliva produced by the infected tick salivary glands (Nuttall and Labuda, 2003). TBEV has been detected in tick saliva and in the cement cone that forms around the tick mouthparts during feeding (Chunikhin et al., 1988; Alekseev et al., 1996). During blood-feeding of ixodid ticks, extensive development of their salivary glands occurs. Moreover, 33–50% of all fluid ingested by the tick is excreted back to the host (Bowman et al., 2008). Thus, tick feeding involves alternation of blood ingestion and saliva secretion, providing an opportunity for recirculation of virus in and around the tick co-feeding sites. Furthermore, increased virus titres have been detected in feeding I. persulcatus (Alekseev and Chunikhin, 1990b) and I. ricinus (Khasnatinov et al., 2009; Belova et al., 2012). Reports of increased TBEV prevalence in ticks removed from humans and animals in comparison to questing ticks (Süss et al., 2004; Romanenko and Kondratyeva, 2011) support the concept that tick feeding affects TBEV infection in ticks. Research on TBEV has largely focused on molecular and pathogenic viral characteristics. By comparison, the biotic factors contributing to successful TBEV survival in natural foci are poorly understood. Therefore, in order to understand better the nature of these complex virus–vector–host interactions and to identify the primary biotic factors likely to be responsible for the survival of TBEV in its natural environment, we compared prevalence and levels of infection in salivary glands and in moulted ticks following infection by either virus inoculation or co-feeding transmission.
Materials and methods Ticks, animals and virus Adult Ixodes ricinus ticks were collected by blanket dragging the vegetation in localities in SW Slovakia known to be free of TBEV. The F1 generation of larvae, nymphs and adults, reared in the laboratory by feeding on uninfected laboratory rabbits, were used in the experiments. California rabbits (3 kg) were obtained from the Research Institute for Animal Production (Nitra, Slovakia). Balb/c (20 g) used in virus transmission experiments and ICR mice (25 g) used in tickfeeding experiments were bred at the Institute of Experimental Pharmacology and Toxicology (Dobrá Voda, Slovakia). Rearing of ticks was carried out as described in Slovák et al. (2002). The prototype Czech TBEV strain Hypr (accession number U39292) prepared as a 10% mouse brain suspension of 1.1 × 109 PFU/ml in Leibovitz’s L-15 medium was used for inoculation of ticks. The usage of animals in the experiments was approved by the State Veterinary and Food Administration of the Slovak Republic (permit numbers 2362/06221, 928/10-221 and1335/12-221). Euthanasia of the animals at the end of the experiments was by cervical dislocation under deep anaesthesia induced using carbon dioxide.
Virus inoculation of ticks and partial feeding Unfed female ticks were inoculated with a defined concentration of 0.5 l of TBEV suspension into the coxal plate of the second pair of legs under a photomacroscope (Wild M 400, Wild Heerbrugg AG, Switzerland), using a digital microinjectorTM system (MINJ-D-CE, Tritech Research, Inc., USA), as previously described (Khasnatinov et al., 2009). Inoculated female ticks were incubated at 22 ± 1 ◦ C and 85–90% RH for a maximum of 120 days. Immediately after inoculation and at defined periods of incubation (dpi = days post inoculation), a cohort of the inoculated ticks was removed from the incubation chamber and placed in retaining chambers on ICR mice, as previously described (Labuda et al., 1993a). Five inoculated female ticks together with 3 uninfected male ticks (to facilitate female tick feeding) were placed within the retaining chamber on each mouse. The tick-infested mice were retained for 2 or 3 days, which is sufficient time for the inoculated ticks to attach and commence feeding but insufficient time for the female ticks to complete engorgement (requiring 6–10 days). Unfed I. ricinus nymphs were inoculated with 0.1 l of TBEV suspension (1.1 × 109 PFU/ml) into the coxal plate of the second pair of legs in the same manner as described for tick females. Inoculated nymphs were incubated at 22 ± 1 ◦ C and 85–90% RH for 16 or 30 days and then fed on mice. The engorged nymphs either were assayed for virus immediately after drop off or they were incubated and allowed to moult to adults. A cohort of moulted females was fed on mice for 3 days; the rest of the females were assayed for virus without being fed.
Co-feeding transmission and trans-stadial survival of the virus The model of co-feeding transmission was carried out essentially as described previously (Labuda et al., 1993a). Briefly, Balb/c mice were infested with two virus-inoculated I. ricinus females 14 dpi, two uninfected males and either 20 nymphs or approximately 100 larvae placed together in one retention chamber at the same time. Following 1, 2 and 3 days of co-feeding, nymphs were subsequently removed from three of the experimental mice. Of the partially fed cohorts of nymphs, 50% were dissected and the presence of TBEV was tested separately in their salivary glands (SG) and body carcasses (remains of the tick after removal of SG), whereas the remaining nymphs were tested without dissection. The rate of co-feeding virus transmission was determined as % of positive nymphs. Infectivity of co-feeding female ticks was confirmed by plaque assay of their SG. To study trans-stadial survival, fully engorged larvae and nymphs that had dropped off the host were allowed to moult. At various time intervals following moulting to nymphs and adults, unfed and fed ticks were dissected and their SG and body carcasses were tested by infectivity assays for the presence of TBEV.
Tick dissection The mice on which ticks fed were killed and ticks were gently removed with fine forceps. Both unfed and partially fed ticks were washed in distilled water and then in sterile 0.15 M NaCl (0.9%) with antibiotics (5% Penicillin/Streptomycin mixture). SG were dissected out individually under a stereomicroscope in ice-cold sterile 0.15 M NaCl, rinsed twice in sterile 0.15 M NaCl to minimise contamination with haemolymph and virus inoculum, and stored individually in Eppendorf tubes containing 50 l of sterile 0.15 M NaCl, at −80 ◦ C. Carcasses and undissected nymphs were individually stored in Eppendorf tubes at −80 ◦ C.
Please cite this article in press as: Slovák, M., et al., Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks. Ticks Tick-borne Dis. (2014), http://dx.doi.org/10.1016/j.ttbdis.2014.07.019
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Fig. 1. Dynamics of TBEV infection in I. ricinus salivary glands. Proportion (%) of infected ticks and virus titres in SG of fasting or 2–3 days-fed adult female ticks at different times following inoculation with 5.5 × 104 PFU of TBEV. Error bars indicate SEM. Asterisks denote significant differences in mean values for partially fed ticks compared with unfed ticks at corresponding days post inoculation: ***P < 0.001, *P < 0.05 (Kolgomorov–Smirnov test). The complete dataset is presented in Table S1 (supplementary file).
Plaque assay Virus presence and infectivity titres were determined using plaque assays, which were performed on Porcine kidney stable (PS) cell monolayers. Prior to assays, SG were homogenised individually in 50 l sterile 0.15 M NaCl and the volumes were completed to 500 l with culture medium. Tick carcasses and whole nymphs were thawed and homogenised in 500 l culture medium. Samples were centrifuged at 2000 rpm for 10 min at 4 ◦ C and the supernatants were kept. PS cells were seeded in 24-well plates (2.5–3 × 105 cells/well) in medium comprising inactivated 7% foetal bovine serum (FBS) solution, Medium 199 [a combination of Medium 199 in Hank’s buffer (150 ml) and Medium 199 in Earle’s buffer (350 ml)] and Penicillin/Streptomycin. 100 l of undiluted or serially diluted tick samples were used to infect the PS cells. After incubation of the virus with cells for 4 h at 37 ◦ C in 5% CO2 , Medium 199 containing 2% carboxymethyl cellulose (CMC) (1:1) and buffered with sodium bicarbonate to pH 7.0–7.5 was added and plates were then incubated for 4 days at 37 ◦ C in 5% CO2 . Thereafter, plates were fixed with 4% formalin and plaques were detected by staining with 0.3 ml of crystal violet for 3 min. After staining, plates were washed using tap water and plaques were counted. Virus titres were expressed as log10 PFU/tick. Statistical analysis Statistical analyses (Kolmogorov–Smirnov test, regression analysis) were performed on the raw data (PFU/tick, ignoring zero values). Descriptive statistics were calculated based on logtransformed data, using Statgraphics Plus 5.1.software. Chi-square statistics was used to compare trans-stadial survival rates in nymphs after co-feeding and parenteral inoculation.
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Fig. 2. Ratio of the total amount of infectious virus detected in SG of partially fed I. ricinus adult female ticks compared with unfed ticks at different days after inoculation (dpi).
decreased until 120 dpi when the mean titre was 2.1 log10 PFU/SG and monitoring was discontinued. The cohort of fasting ticks that were allowed to feed for 2–3 days at different times following inoculation also showed an increase in the number of ticks infected and in the SG virus titres, with increasing dpi (Fig. 1, Table S1B). All of the ticks in this cohort had infected SG by 7 dpi (4 days fasting + 3 days feeding) and the mean maximum titre in SG of 4.3 log10 PFU/SG was observed 24 dpi (21 days fasting + 3 days feeding). Comparison of the two tick cohorts revealed that, as measured by plaque assay, partial feeding increased the rate at which infection of the SG was detected, the rate of virus replication in the SG, and enhanced the total amount of infectious virus measured in the SG (Fig. S1). The amplification in SG virus yield was greatest when inoculated ticks were immediately fed on mice for 2 days (total dpi = 2) and then reduced over an extended period until ∼120 dpi when blood-feeding did not appear to affect the virus yield from SG (Fig. 2). To examine whether the dose of virus inoculated into ticks affected the enhancing effect of partial feeding, five groups of female ticks were each inoculated with 10-fold differences in virus dose. At 14 dpi, 50% ticks in each group were fed on mice for 3 days. All of the unfed ticks inoculated with TBEV doses >55 PFU per tick were positive (Fig. 3, Table S2). With lower doses of inoculated virus, the percentage of positive ticks detected was reduced but remained high (75–94%). In contrast, 100% of partially fed ticks, inoculated with virus, were positive irrespective of the virus dose. Apart from the highest dose in unfed ticks, virus titres in SG of both unfed and partially fed ticks declined with decreasing doses: in unfed ticks from 3.7 to 2.0 log10 PFU/SG and in ticks fed for 3 days, from 3.3 to 2.8 log10 PFU/SG. When virus titres in fasting ticks were
Results Dynamics of virus replication in tick salivary glands An estimated 5.5 × 104 PFU of TBEV were inoculated into the haemocoel of I. ricinus females and the infection rate and TBEV titres in SG were determined at different time intervals in fasting and fasting + feeding ticks as shown in Fig. 1 and Table S1 (see supplementary file). In the cohort of inoculated unfed ticks, infectious TBEV was detected in 25% of the SG at 2 dpi and 100% of SG were infected by 14 dpi (Fig. 1, Table S1A). The highest average titres of 2.9 log10 PFU/SG were recorded at 21 dpi after which titres gradually
Fig. 3. Dose-dependence of TBEV replication in I. ricinus. Proportion (%) of infected ticks and virus titres in SG of fasting or 3 days-fed adult female ticks, 14 days following inoculation with different doses of TBEV. Error bars indicate SEM values of virus titres as presented in Table S2 (supplementary file).
Please cite this article in press as: Slovák, M., et al., Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks. Ticks Tick-borne Dis. (2014), http://dx.doi.org/10.1016/j.ttbdis.2014.07.019
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Fig. 4. Comparative dynamics of TBEV transmission to I. ricinus nymphs during cofeeding with infected female ticks for either 1, 2 or 3 days.
plotted as a function of inoculation dose (ignoring zero values), the regression was significant (P < 0.001). The regression formula was: Y = 1.68x + 1403 (R2 = 0.55, F = 19.44, n = 46). Significant regression in fed ticks was not detected (data not shown). Dynamics of co-feeding transmission Groups of two I. ricinus females, 14 dpi, were placed together with 20 uninfected nymphs in a retention chamber on the shaven skin of mice and allowed to co-feed for 1–3 days as described in Section “Materials and methods”. The rate of TBEV transmission was expressed as the percentage of TBEV-positive nymphs (Table 1). Maximum co-feeding transmission (61%) was recorded on day 3. The dynamics of TBEV accumulation in the SG and body carcass of nymphs compared with whole intact nymphs are presented in Fig. 4. During the 3-day co-feeding period, no virus was detected in the SG of nymphs, only in the remaining body carcasses and in whole intact nymphs. Virus titres in body carcasses and whole intact bodies of nymphs were 1.7–2.0 and 2.0–2.3 log10 PFU/tick, respectively. Dynamics of trans-stadial survival In experiments to study trans-stadial survival, uninfected nymphs were allowed to co-feed with infected adults until repletion of the nymphs (usually within 3–5 days of co-feeding). The replete nymphs were then allowed to moult into adults and the percentage of positive female ticks was determined at different time intervals during 90 days post-moult; cohorts were allowed to feed for 3 days to determine whether feeding amplified virus infection. To trace TBEV traffic inside the moulted female ticks, their SG and body carcass were investigated (Table 2). Of the 82 female ticks examined, 7 (8.5%) showed evidence of virus infection. Based on a 61% transmission rate (Table 1), the expected level of infection was 50/82 if 100% infections survived moulting. Hence the observed level of 8.5% indicates a trans-stadial survival rate of
14%. Infectious virus was detected in the SG of only two of these females, both of which also had virus positive carcasses. These two ticks with infected SG had the highest virus titres in their carcasses. Partial feeding of the adults did not increase the number of virus positive ticks or the virus titres. No infected ticks were detected after 60 days post-moult and infections in SG were not detected after 14 days post-moult. Trans-stadial survival was also examined using larvae. Uninfected larvae were allowed to co-feed with infected adults until repletion and drop off. The engorged larvae were then allowed to moult to nymphs. At various times after moulting, the nymphs were assayed for virus either as whole undissected ticks or following dissection as SG and carcass. Cohorts were partially fed on uninfected mice for 2 days to determine whether feeding amplified virus infection following moulting. Only one of the 129 nymphs that moulted from engorged larvae was positive for infectious virus. This infected nymph, examined 14 days after moulting, had a remarkably high virus titre in its SG, which was higher than the titre in its body carcass (Table 3). As was the case for moulted adults, partial feeding of moulted nymphs did not affect the prevalence or incidence of infection. For comparison with the natural route of co-feeding infection, nymphs were inoculated with virus and then allowed to feed to repletion, drop off and then moult to adults to determine transstadial survival. Two different treatments were examined: feeding nymphs at 16 dpi or at 30 dpi. In the first experiment, 23 nymphs were inoculated with an estimated 1.1 × 105 PFU TBEV of which 15 (65.2%) survived. At 16 dpi, the nymphs were allowed to feed on Balb/c mice; 12 nymphs completed engorgement. Five of the replete intact nymphs were immediately assayed for virus; all were infected (titres >5.0 log10 PFU/tick). The remaining engorged nymphs moulted into 6 female and 1 male adults; 2 of the females were allowed to feed for 2 days, 45–50 days after moulting. In the second experiment, 80 nymphs were inoculated with the same virus dose; 26 (32.5%) survived. At 30 dpi, the nymphs were allowed to feed on Balb/c mice; 25 nymphs completed engorgement. Ten of the replete nymphs were immediately dissected and then assayed for virus. Of these, 9 had infected carcasses (titres 1.3–3.3 log10 PFU) while 5 had infected SG (titres 1.5–3.1 log10 PFU); the infection prevalence was 90%. The remaining engorged nymphs moulted into 12 female and 3 male adults; 7 of the females were allowed to feed for 2 days. At 45–50 days after moulting, the adult ticks were all assayed for virus either following dissection into SG + carcass (females) or as intact (whole body) ticks (males) (Table 4). Following moulting, 19/22 (86.4%) adults were infected (100% unfed and 66.7% fed; 83.3% females and 100% males). There was no obvious difference in results obtained following feeding of nymphs either 16 or 30 dpi, or between unfed and partially fed adults, although numbers were comparatively low as a result of failure of nymphs to survive the trauma of inoculation. However, as observed following co-feeding infection, all female ticks that had infected SG also had an infected body carcass, and the infection prevalence in body carcasses (mean 83.3%) was greater than that of SG (mean 22.2%).
Table 1 Comparative dynamics of TBEV transmission to I. ricinus nymphs during co-feeding with infected female ticks. Days of co-feeding
1 2 3 a
SG
Body carcass
No. positive/total (%)
No. positive/total (%)
Mean ± SEM (log10 PFU/tick)
No. positive/total (%)
Mean ± SEM (log10 PFU/tick)
No. positive/total (%)
0/6 (0%) 0/6 (0%) 0/7 (0%)
0/6 (0%) 1/6 (17%) 4/7 (57%)
0 2.0 1.72 ± 0.29
1/6 (17%) 2/6 (33%) 4/6 (67%)
1.95 2.12 ± 0.42 2.25 ± 0.32
1/12 (8%) 3/12 (25%) 8/13 (61%)
Whole body
Total no. ticks examineda
Infectious virus was measured in homogenates of either the SG and remaining body carcass of a tick or the whole body of a tick.
Please cite this article in press as: Slovák, M., et al., Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks. Ticks Tick-borne Dis. (2014), http://dx.doi.org/10.1016/j.ttbdis.2014.07.019
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Table 2 Trans-stadial survival of TBEV from I. ricinus nymphs to females following co-feeding infection of the nymphs. Sample assayed for virus
Days after moulting (+3 days of feeding) 1
7
14
11 + 3
SG Observed Positive % positive Titrea
6 0 0 0
6 1 17 2.7
14 1 7.14 5.0
14 0 0 0
Carcass Observed Positive % positive Titrea
6 1 17 3.1
6 1 17 3.5
14 1 7.14 5.1
Total ticks Observed Positive % positiveb
6 1 17
6 1 17
14 1 7.1
Total samples 30
27 + 3
60
57 + 3
90
87 + 3
6 0 0 0
8 0 0 0
6 0 0 0
8 0 0 0
8 0 NT NT
6 0 NT NT
82 2 2.4
14 1 7.14 2.6
6 1 17 2.9
8 1 17 3.5
6 1 17 3.2
8 0 0 0
8 0 0 0
6 0 0 0
82 7 8.5
14 1 7.1
6 1 17
8 1 17
6 1 17
8 0 0
8 0 0
6 0 0
82 7 8.5
a Virus titre (log10 PFU/tick) in SG and carcass (tick remains after removal of SG) of unfed or partially fed females at different days after moulting from nymphs that had co-fed with infected females. NT is not tested. b Based on 100% infected nymphs: see text.
Discussion
Studies on TBEV replication in ticks, in which ticks are inoculated parenterally with virus, have generally placed the inoculated ticks onto susceptible animals at a set time after inoculation and allowed the ticks to complete engorgement. For example, female I. ricinus inoculated with 3.7 log10 PFU TBEV were either maintained fasting or placed on experimental mice 15 h after inoculation and then cohorts of the feeding ticks were examined at 8 h, 15 h, 38 h, 96 h post-inoculation. Virus replication in whole feeding ticks was faster and reached higher titres than in fasting ticks at the same times post-inoculation (Belova et al., 2012). The role of the SG in this blood-feeding induced amplification of virus infectivity could not be determined. We took a different approach, taking ticks at different times following inoculation, and then allowing the inoculated fasting ticks to feed only partially. This approach enabled us to take a snapshot of the virus infectivity in the infected SG at different times post-inoculation and after prolonged fasting, reflecting the situation in nature when infected ticks may fast for months and possibly >1 year before feeding. We observed that, when the infected female ticks took a bloodmeal, the level of virus infectivity increased rapidly by at least 3 orders of magnitude. This
This study focuses on three biotic factors involved in the transmission cycle of TBEV by its European vector, Ixodes ricinus: (1) infection of the tick salivary glands, (2) co-feeding transmission, and (3) trans-stadial survival. The first objective was to study the dynamics of infection of the tick salivary glands to understand the significance of their role in long-term TBEV survival and transmission. Our model of inoculation of the virus into the tick haemocoel circumvents the requirement for the virus to cross the tick midgut barrier (Nuttall and Labuda, 2003), thereby providing unimpeded access for the virus to infect the tick SG. This appeared to be a successful strategy as TBEV showed high tropism to I. ricinus SG and virus amplification was detected in SG of a high proportion of inoculated ticks, although the mechanism of virus transfer from the haemocoel to tick SG and saliva is unknown (Kaufman and Nuttall, 1996). Infectious virus was recovered from SG for at least 120 days following its initial inoculation, demonstrating the potential importance of the SG as a long-term reservoir for efficient transmission of the virus.
Table 3 Trans-stadial transmission of TBEV from I. ricinus larvae to nymphs following co-feeding infection of the larvae. Sample assayed for virus
Days after moulting (+2 days of feeding) 1
7
14
12+
Total samples 30
28+
58+
90
88+
8 0 0 0
8 0 0 0
NT – – –
NT NT NT NT
57 1 1.8
6 0 0 0
8 0 0 0
8 0 0 0
– – – –
NT NT NT NT
57 1 1.8
18 0 0
5 0 0
6 0 0
6 0 0
6 0 0
6 0 0
72 0 0
24 0 0
11 0 0
14 0 0
14 0 0
6 0 0
6 0 0
SG Observed Positive % positive Titrea
8 0 0 0
6 1 17 5.7
9 0 0 0
6 0 0 0
6 0 0 0
6 0 0 0
Carcass Observed Positive % positive Titrea
8 0 0 0
6 1 17 3.5
9 0 0 0
6 0 0 0
6 0 0 0
Whole body Observed Positive % positive
6 0 0
3 0 0
8 0 0
8 0 0
17 0 0
14 0 0
Total ticks Observed Positive %
14 0 0
9 1 11.1
60
129 1 0.8%
a Virus titre (log10 PFU/tick) in SG, carcass (tick remains after removal of SG) or whole undissected tick of unfed or partially fed nymphs at different days after moulting from larvae that had co-fed with infected females. NT is not tested.
Please cite this article in press as: Slovák, M., et al., Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks. Ticks Tick-borne Dis. (2014), http://dx.doi.org/10.1016/j.ttbdis.2014.07.019
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Table 4 Trans-stadial survival of TBEV from I. ricinus nymphs to adults following virus inoculation of the nymphs. Sample assayed for virus
Moulted from 16 dpi nymph Female Unfed
+3
SG Observed Positive % positive Titrea (min–max)
4 1 25% 1.30
2 0 0%
Carcass Observed Positive % positive Titrea (min–max)
4 4 100% 2.02 (1.74–2.70)
2 2 100% 2.13 (1.48–2.78)
NT
NT
Whole body Observed Positive % positive Titrea (min–max) Total ticks Observed Positive % positive a
Moulted from 30 dpi nymph Male
Female
Unfed
+3
NT
NT
NT
Unfed
+3
5 1 20% 2.65
7 2 28.6% 2.84 (1.78–3.9)
5 5 100% 1.55 (1.0–2.02)
7 4 57.1% 1.90 (1.0–3.15)
NT
NT
NT
NT
Total samples Male Unfed
+3
NT
NT 18 4 22.2%
NT
1 1 100% >2.00
18 15 83.3% NT 3 3 100% 2.48 (1.18–3.34)
NT 4 4 100%
2 2 100%
NT
1 1 100%
4 4 100% NT
5 5 100%
7 4 57.1%
3 3 100%
22 19 86.4%
Mean virus titre (log10 PFU/tick) in SG, carcass (tick remains after removal of SG) or whole undissected tick. NT, not tested.
feeding-induced amplification of virus infectivity in SG persisted for at least 87 dpi. Thus virus replication in SG may play an important role in maximising the dose of virus transmitted from an infected tick to an uninfected vertebrate host during the prolonged feeding period (6–10 days for female I. ricinus). Feeding-induced enhancement of SG infection may explain reports of significantly higher infection prevalences of TBEV in partially fed ticks removed from humans compared with field-collected ticks from the same locality (summarised in Supplementary Table S1 of Lindblom et al., 2014). The susceptibility of infected SG to blood-feeding induced amplification of the infection appeared to persist for much of the fasting period. The reason for the apparent loss in responsiveness of SG infection by 120 dpi is unknown. Possibly the SG become refractory to virus amplification after long periods off the host as a means of protecting the SG, which play a crucial role in water balance during fasting periods (Bowman et al., 2008). Based on virus inoculation studies and partial feeding at selected times following inoculation, it appears the tick SG are both an important reservoir for TBEV survival during fasting and for amplification of the potential virus dose transmitted to a vertebrate host during blood-feeding. However, these observations based on parenteral inoculation of TBEV were not reproduced in subsequent studies involving the natural route of co-feeding infection of ticks. For co-feeding transmission, uninfected nymphs were allowed to feed with TBEV-inoculated I. ricinus females for 1–3 days, on localised skin sites of mice. The experiments demonstrated comparatively high rates of co-feeding transmission with up to ∼60% of nymphs infected, supporting previous observations that co-feeding is a highly efficient mode of virus transmission. In previous studies, the rate of engorged I. ricinus nymphs acquiring TBEV during co-feeding in close proximity to infected females on laboratory hosts varied between 45% on guinea pigs (Labuda et al., 1993a) and 93% on Balb/c mice (Labuda et al., 1996), and for natural hosts, between 10% on hedgehogs and up to 80% on yellow-necked mice (Apodemus flavicollis) (Labuda et al., 1993b, 1997). Transmission efficiency of different European strains of TBEV for nymphs cofeeding with infected adults on Balb/c mice varied between 93% for strain 198 (isolated from I. ricinus in Slovakia) (Labuda et al., 1996) and 64% for the Hypr strain (Kazimírová et al., 2012) and 61% (this study).
When I. ricinus nymphs co-fed with infected females, the percentage of nymphs acquiring the virus increased during co-feeding. However, after 3 days of co-feeding, infectious virus was detected only in their bodies and not in their SG. In previous studies aimed to understand the role of local skin infection and saliva assisted transmission in non-viraemic co-feeding transmission, the percentage of I. ricinus nymphs acquiring TBEV increased in parallel with the duration of co-feeding on natural rodent hosts as well as on laboratory mice from day 1 to day 3 (Labuda et al., 1996). Similar results were achieved by infestation of yellow-necked mice with infected I. ricinus females and short-feeding Haemaphysalis inermis nymphs (Labuda et al., unpublished). However, in all these studies only whole bodies of co-feeding nymphs were screened for the presence of TBEV, and not the SG and body carcasses. The consistently higher incidence of infection and virus titre of whole bodies compared with the sum of the virus yields from SG + carcass is most likely accounted for by infectious virus lost in the haemolymph during the tick dissection as TBEV has been detected in tick haemocytes ˇ (Rehᡠcek and Mrenová, 1966). If this is the case, the presence of virus in the haemolymph indicates that virus acquired within the bloodmeal is able to escape through the tick midgut wall. The estimated mean titre of virus in the haemolymph was approximately 125 PFU/tick which, compared with parenteral inoculation of adult females with different doses of virus (Table 3), should have been sufficient to infect the SG of most of the nymphs (assuming the SG of nymphs show similar susceptibility and infection dynamics to those of adult I. ricinus). Possibly the difference is due to the dynamics of infection of SG by inoculated virus compared with naturally acquired virus. In the case of parenteral inoculation, free extracellular virus can directly access the SG whereas virus that escapes from the midgut may infect haemocytes and/or other tissues and be unable to invade the SG directly. The observation that partial feeding of moulted nymphs and moulted larvae did not increase the prevalence of infection in the co-fed tick population is surprising, given that the virus titres in the carcasses of the infected co-fed ticks were comparable to those observed in the ticks infected by inoculation. Moreover, only 2/7 infected ticks had infectious virus detectable in their SG (Table 4). Again, this is surprising given the speed at which TBEV appeared to infect SG following virus inoculation into the haemocoel. The
Please cite this article in press as: Slovák, M., et al., Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks. Ticks Tick-borne Dis. (2014), http://dx.doi.org/10.1016/j.ttbdis.2014.07.019
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observations suggest there may be a midgut and/or salivary gland constraint on TBEV infection and/or innate immune response that is circumvented by virus inoculation into the haemocoel. For example, if virus acquired in the bloodmeal fails to escape the midgut prior to moulting, it may be destroyed. Following 3 days co-feeding of nymphs with infected adults, 61% nymphs became infected. However, the infection prevalence after moulting to adults was only 8.5%, indicating a maximal transstadial survival rate of 14% (assuming a minimal 61% infection prevalence of the engorged nymphs prior to moulting). By contrast, the trans-stadial survival rate following moulting of engorged nymphs infected by inoculation was 92.6% (based on an infection prevalence of 93.3% for the engorged nymphs), significantly higher than the maximal 14% trans-stadial survival rate for nymphs infected by co-feeding (Chi-square = 56.03; P < 0.01). The estimated trans-stadial survival rate through moulting from larva to nymph following co-feeding infection of larvae was 0.8% assuming all engorged larvae were infected. Virus infection of larvae by inoculation technically was not possible so we were unable to compare the trans-stadial survival rates following co-feeding with those following infection by inoculation. We are unaware of any comparable studies on trans-stadial survival following co-feeding infection of ticks. Trans-stadial survival rates for engorged larvae and engorged nymphs of 9.5% and 54%, respectively, were reported for Powassan virus (a member of the TBEV complex) following infection by feeding on viraemic hamsters (Costero and Grayson, 1996). Nevertheless, our data correspond with epidemiological data that reveal a low TBEV-Eur prevalence in questing I. ricinus caught in nature, which is generally between 0.1 and 1% infectivity over different areas and years of investigation, and mainly reflects the infection prevalence in unfed nymphs (e.g. Danielová et al., 2002a; Carpi et al., 2009; Lommano et al., 2012). At this stage of the research, we do not know if the low rate of trans-stadial survival is a general phenomenon for TBEV infection of I. ricinus. The prototype TBEV strain Hypr has been adapted to laboratory mice via serial passages and in contrast with many wild-type TBEV isolates the 3 untranslated region (3 UTR) has a deletion of 263 nucleotides (e.g. Wallner et al., 1995; Hayasaka et al., 2001) which is believed to be a replication enhancer region of the 3 UTR (Gritsun and Gould, 2007a,b). Virus genetic characteristics encoded in 3 UTR might be essential for the virus to reach a critical replication rate and also to circumvent the tick immune responses (Pijlman et al., 2008; Gritsun et al., 2014) thus contributing to the efficiency of TBEV dissemination and amplification in ticks and transmission to co-feeding ticks and hosts. In conclusion, although our data support modelling predictions that co-feeding (non-viraemic) transmission of TBEV is essential for survival of the TBEV in nature, they raise questions about the parameters used to estimate the basic reproductive number (R0 ) for TBEV. In particular, it is generally assumed that the infection in ticks survives from one developmental stage to the next and does not need to be parameterised (Hartemink et al., 2008; Harrison and Bennett, 2012). Given our observations, the loss of an infection during moulting could have a highly significant effect on estimates of R0 . However, it remains to be determined if strain Hypr is an exception to the rule for TBEV survival. This is currently being investigated using infectious clones of the viruses.
Acknowledgements This paper is dedicated to the memory of Milan Labuda, whose idea it originally was. The authors thank him for everything he taught them. The technical assistance of Mrs Zuzana Guláˇsová is highly acknowledged. The work was supported by Slovak Scientific Grant Agency, VEGA (grant no. 2/0191/12), European
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Commission (European Virus Archive, FP7 CAPACITIES project – GA no. 228292), grant agency APVV (no. APVV DO7RP–0014–11), EU grant FP7-261504 EDENext and is catalogued by the EDENext Steering Committee as EDENext240 (http://www.edenext.eu), BBSRC UK grant BBS/B/00697, MRC UK grant G0801208 and Royal Society UK grant JP100471. E A Gould is funded by EU FP7 grant no. 260644–SILVER. The contents of this publication are the sole responsibility of the authors and do not necessarily reflect the views of the European Commission. Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:10.1016/j.ttbdis.2014.07.019. References Achazi, K., Ruzek, D., Donoso Mantke, O., Schlegel, M., Ali, H.S., Wenk, M., SchmidtChanasit, J., Ohlmeyer, L., Ruhe, F., Vor, T., Kiffner, C., Kallies, R., Ulrich, R.G., Niedrig, M., 2011. Rodents as sentinels for the prevalence of tick-borne encephalitis virus. Vector-Borne Zoonotic Dis. 11, 641–647. 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Please cite this article in press as: Slovák, M., et al., Survival dynamics of tick-borne encephalitis virus in Ixodes ricinus ticks. Ticks Tick-borne Dis. (2014), http://dx.doi.org/10.1016/j.ttbdis.2014.07.019