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1 Department of Pathology, and Center for Biodefense and Emerging Infectious Diseases, University of Texas Medical Branch, Galveston, Texas, USA
2 Department of Medicine, Zhong-nan Hospital, Wuhan University, Wuhan, Hubei Province, PR China
Correspondence
Shu-Yuan Xiao
syxiao{at}utmb.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). Rarely, a poliomyelitis-like illness can develop (Centers for Disease Control and Prevention, 2002; Glass et al., 2002; Leis et al., 2002; Ohry et al., 2001). In a few patients, WNV infection is expressed as a Guillain-Barre-like syndrome with acute flaccid paralysis (Asnis et al., 2000) and significant muscle weakness leading to respiratory distress. The pathogenesis of WNV is largely unknown, but both host and viral factors, including host innate and adaptive immune responses, and tissue tropism of the virus, are likely to be important. Therefore, elucidation of the genetic determinants of tissue tropism should lead to a better understanding of at least some aspects of WNV pathogenesis.
To date, many studies have focused on examining natural WNV variants, which show differing neurovirulence or neuroinvasiveness in rodent or avian models of the disease (Beasley et al., 2001, 2002; Ceccaldi et al., 2004; Langevin et al., 2005; Brault et al., 2004). Changes in nucleotide sequences were identified among these isolates, but their significance remains uncertain, due to the lack of consistency when comparing the biological characteristics and neurovirulence of a diverse group of viruses in different hosts. Recently, we have made a series of interesting observations in a hamster model of WNV encephalitis (Xiao et al., 2001), that we believe are relevant to the study of WNV neuropathogenesis. Initially, it was observed that some hamsters surviving experimental WNV infection developed chronic viruria (Tonry et al., 2005). A subsequent study showed that adult hamsters experimentally infected with the NY385-99 strain of WNV developed chronic renal infection and persistent shedding of virus in urine for up to 8 months, despite initial rapid clearance of virus from blood and the timely appearance of high titres of neutralizing antibody (Tesh et al., 2005). WNV could be recovered from kidney tissue by co-cultivation, or demonstrated by RT-PCR as well as immunohistochemically. In addition, when the viruses recovered from urine of the persistently infected hamsters (urine isolates) were inoculated into naïve hamsters, they no longer caused neurological illness or mortality, as the parent NY385-99 virus did. Instead, they produced chronic renal infection and viruria in most of the inoculated hamsters. These observations suggested a lack of neurovirulence in these hamster-passaged urine isolates. Nucleotide sequence analysis of several first-urine-passage isolates, and of the ‘parent’ NY385-99 strain showed a number of nucleotide changes in the progeny virus strains compared with the parent virus, suggesting that some of these sequence changes were associated with the phenotype of enhanced renal tropism and loss of neurotropism (Ding et al., 2005). However, the possibility that these changes were simply random variants could not be ruled out, as it was not known if the nucleotide changes or new phenotypes persisted after additional passages. Furthermore, it was uncertain if additional passage of the urine isolates in vivo would maintain the same phenotype. In the current study, four consecutive urine isolates, representing four serial passages of WNV, were used.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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WNV strain NY385-99 was the parent virus used to initially infect hamsters. It was originally isolated from the liver of a dead snowy owl (Nyctea scandiaca) found in the Bronx Zoo in New York City, during the 1999 WNV outbreak (Xiao et al., 2001a). This virus strain had been passaged three times in Vero cells and has been used extensively in the development of our hamster model of WNV encephalitis (Sbrana et al., 2005; Tesh et al., 2002a, b, 2005; Tonry et al., 2005; Xiao et al., 2001a). The other WNV strains examined in this study were isolates of the virus obtained from the urine of chronically infected hamsters after one, two, three or four serial hamster passages over a 13 month period, as described below.
Fig. 1 shows the passage history and source of the WNV strains used in this study. Initially, 10 hamsters were inoculated intraperitoneally (i.p.) with 
104 p.f.u. of the parent NY385-99 virus. After about 7 weeks, one of the surviving hamsters (H9317B) was killed and dissected and urine was aspirated directly from the urinary bladder. Culture of the urine in Vero cells confirmed that it contained WNV. Fetal bovine serum (FBS, 25 %) was added to the remaining urine, which was frozen at –80 °C as hamster passage number one (HP1), strain H9317B. A proportion (200 µl) of this first hamster passage infectious urine was inoculated i.p. into a single naïve hamster (T35639
[GenBank]
). Twelve days after inoculation, this animal was killed and infectious urine was collected from its bladder. Virus isolated from this second urine passage (HP2) was designated T35639.
[GenBank]
Similarly, the HP2 virus (200 µl per animal) was inoculated i.p. into six additional naïve hamsters (H8533–H8538); this represented the third serial hamster passage (HP3). Over the next 6–8 months, freshly voided urine was collected periodically from these six hamsters. A fourth passage was made when urine was collected from hamster H8535 and inoculated i.p. into two new hamsters (H8912 and H8915). Urine samples, representing the fourth serial passage (HP4), were subsequently collected and tested positive for WNV. Although sequence analyses had been carried out with multiple isolates from these various hamster hosts (H8533–H8538), only viruses from a single hamster ‘lineage’, H8535, were further studied and are described in this paper.
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). The freshly voided urine was aspirated, diluted 1 : 10 in PBS containing 25 % FBS, then 200 µl of the diluted sample was inoculated into a 12.5 cm2 flask culture of Vero cells. After absorption for 1 h at 37 °C, maintenance medium consisting of minimum essential medium with Earle's salts (EMEM; Gibco), 1.5 % heat-inactivated FBS (Sigma) and penicillin–streptomycin (Sigma) was added. Cultures were held in a 37 °C incubator and observed daily for viral cytopathic effect (CPE). When viral CPE was observed, a sample (125 µl) of the cell culture fluid was removed and tested for the presence of WNV, using the VecTest WNV antigen assay kit (Medical Analysis Systems), following the manufacturer's instructions. If the sample tested positive for WNV, the remaining culture fluid was decanted and 5 ml Trizol-LS reagent (Invitrogen) was added to the flask to lyse the cells in preparation for subsequent RNA extraction.
RNA extraction, RT-PCR and sequencing.
The infected cell culture–Trizol mixture (100 µl) was mixed with 900 µl Trizol. Total RNA was extracted by the chloroform/2-propanol method (Liu et al., 2003). The resulting RNA pellet was dissolved in 30 µl RNase-free water and the RNA concentration was determined from its A260 and A280 using a Beckman Coulter UV–visible spectrophotometer. First-strand cDNA was synthesized by using SuperScript III First-Strand Synthesis System (Invitrogen), following the manufacturer's protocol. The reaction mixture (20 µl) contained 5 µl (1 µg µl–1) total RNA, 1 µl random hexamer primer (50 ng µl–1), 1 µl 10 mM dNTP mix, 3 µl DEPC-treated water, 2 µl 10x RT buffer, 4 µl 25 mM MgCl2, 2 µl 0.1 M DTT, 1 µl RNaseout (40 U µl–1) and 1 µl reverse transcriptase (200 U µl–1). The reaction was stopped by heating at 85 °C for 5 min, followed by the addition 1 µl RNaseH (2 U µl–1) and incubation at 37 °C for 20 min.
The primers used for PCR amplification have been described previously (Ding et al., 2005), with resultant fragments covering the full-length RNA genome. PCR products of 500–600 bp in length, with 50–100 bp overlap between two adjacent target regions, were obtained for each virus. The resultant DNA was separated and visualized with 1.5 % agarose gel electrophoresis, purified using the QIAquick kit (Qiagen), and directly sequenced in both directions with the amplifying primers by using the ABI 3100 Genetic Analyzer (Applied Biosystems) at the university's BioMolecular Resource Facility (Protein Chemistry Laboratory). The sequence data presented in this paper were collected at lease twice. Any new nucleotide change was confirmed by the following approaches. First, the RT-PCR procedure was repeated from the original Trizol lysate and the sequence of the PCR product was determined again. Alternatively, purified DNA was cloned into the pGEM-T Easy vector (pGEM-T Easy Vector Systems; Promega), and three clones were sequenced at the sites of interest.
Analysis of sequences.
Initial assembly of sequence data were performed using the SeqMan program of the DNASTAR software package. Nucleotide and deduced amino acid sequences of the complete genome and fragment genome of each isolate were aligned using the MEGALIGN program (DNASTAR). The analysis of protein characteristics was performed by using the Protein program in DNASTAR. All isolates were then compared with NY385-99 (GenBank accession no. AY842931
[GenBank]
). NetNGlyc software (post-translational modification and topology prediction) in ExPASy Proteomics tools (ExPASy, www.expasy.org) was used to detect the glycosylation site in the E-protein gene.
Histological and immunohistochemical (IHC) analysis.
Brain, spinal cord and kidney with adrenal, were removed after exsanguination by cardiac puncture under Halothane (Halocarbon Laboratories) anaesthesia. These were fixed in 10 % buffered formalin for 20–24 h and then transferred to 70 % alcohol. The tissues were processed as described previously (Xiao et al., 2001b), with one haemotoxylin and eosin stained and several additional unstained sections prepared. IHC staining was performed using an established protocol (Xiao et al., 2001a), with the primary WNV polyclonal antibody used at a dilution of 1 : 100 and incubated at 4 °C overnight. Stained tissue samples were examined with an Olympus BX51 microscope, with an attached DP-70 digital camera, to record the representative images.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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and 2. The mutations involved genomic regions of the nucleocapsid protein (C), envelope protein (E), non-structural proteins (NS1, NS2, NS3 and NS5) and the 3' non-coding region (NCR). Nine mutations were shared by all isolates, which are located at nt 1027 (G to A), 1465 (C to T), 3017 (T to C), 4515 (G to A), 6405 (T to C), 8235 (T to C), 8849 (A to G), 10545 (C to T) and 10655 (C to T). Among these mutations, five resulted in amino acid substitutions from Val to Met at E-21, Leu to Phe at E-167, Ile to Thr at NS1-183, Met to Ile at NS2B-99, Glu to Gly at NS5-390, respectively. Almost all nucleotide changes (99.97 %) were T–C and A–G transitions. The other types of transition such as G–T, C–G or A–T were found in only a few nucleotide changes (5/116); four out of the five were sense mutations. These results are consistent with findings of earlier reports (Anderson et al., 2001; Ebel et al., 2001). No nucleotide differences were identified between the 5'-NCR sequences of NY385-99 and its progeny, while 11 nt sequences differed in the 3'-NCR. In addition, in the 3'-NCR, nt substitutions at 10545 (C to T) and 10655 (C to T) were identified among all of the isolates from hamster urine examined in this study.
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There were three nucleotide changes between the HP3 (H8535) and HP1 (H9317B) isolates.
Phenotypic characterizations of WNV isolates recovered from persistently infected hamsters
Four groups of adult hamsters were inoculated i.p. with the WNV strains representing the parent and three different hamster passages of the same lineage: virus NY385-99 (group 1), H9317B (group 2), H8535 (group 3) and H8912 (group 4). Group 1 contained 19 hamsters; the other three groups each had 10 animals. Five animals in each group were bled daily to determine the level of viraemia and antibody response. In all groups, viraemia developed by 2 days post-infection (p.i.), reaching peak levels at 4 days p.i., and gradually declining afterward (Fig. 2). The levels of viraemia among hamsters in the four groups were similar, except for group 4 (inoculated with H8912), which showed a lower peak level than the other three groups (Fig. 2). Seroconversion began by day 7 p.i., with detectable haemagglutination inhibition (HI) antibody present in all animals by day 8 (Table 3), except for one animal in group 1; this animal did not develop detectable viraemia either, so it was considered not to be infected.
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Table 4 shows the duration of West Nile viruria in the surviving hamsters in the four groups. Urine was collected from the animals and cultured at intervals of 24, 56, 72, 95 and 128 days p.i. Although it was not possible to obtain a urine specimen from every animal on a given day, the pattern seen with the hamsters was chronic viruria. Most of the hamsters from groups 2 to 4 were still shedding WNV in their urine when tested 128 days after the initial infection, indicating persistent renal infection, while only one of four hamsters from group 1 was still positive by this day.
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). No evident abnormalities were discerned in their other organs. In contrast, no histopathological changes were observed in the brain tissue or organs of the animals in groups 2, 3 or 4. (Fig. 3b, d). Likewise, WNV antigen was identified by IHC staining in brain tissue and spinal cords of the three animals that were virus-culture positive (Fig. 3e). No viral antigen was identified in the brain or spinal cord of hamsters in the other three groups (Fig. 3f). WNV antigens were identified in kidney or adrenal cortex of some of the animals in group 3 but not in the other hamsters.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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, 2002; Ceccaldi et al., 2004; Davis et al., 2004; Langevin et al., 2005). Working with our WNV–hamster model, we discovered that some of the animals that survived the acute infection continued to shed virus in their urine, and that these urine isolates lacked neurovirulence (Tesh et al., 2005). For example, in one experiment, we observed that i.p. inoculation of the WNV strain NY385-99 into 10 adult hamsters resulted in a 40 % mortality, but that 67 % of the survivors had viruria transiently (Ding et al., 2005). When a urine isolate from one of these chronically infected hamsters was inoculated i.p. again into a group of 60 naïve hamsters, none of them died. Sequence analysis of three urine isolates from hamster passage 1 showed some nucleotide changes. However, a question raised from this previous study was whether the nucleotide sequence and phenotypic differences identified in the hamster-passaged viruses were stable. Therefore, in the current study, urine isolates representing three different hamster passages were studied.
As shown in the results, nucleotide changes in viruses isolated from persistently infected hamsters ranged from 0.082 to 0.262 % as compared with the parent virus NY385-99. The changes were distributed at a total of 116 sites. Most changes were in the coding regions, some causing amino acid substitutions at the genomic regions of C, E, NS1, NS2 and NS5. Only three additional nucleotide changes were identified in the hamster passage three virus, H8535. However, significantly more changes occurred in the hamster passage four virus, H8912. One of the contributing factors maybe the much longer time period (9 months) elapsing between the isolation of virus strains H8912 and H8535, as opposed to the time between isolation of strains H8535 and H9317B (about 2 months) (Fig. 1
).
The E protein of flaviviruses is considered to be crucial in mediating the virus–host interaction (Crill & Roehrig, 2001; Heinz et al., 1994; Helenius, 1995; Rey et al., 1995); it is also thought to be responsible for viral entry and fusion (Allison et al., 1995). Neutralizing monoclonal antibodies directed against this protein are capable of blocking virus-mediated cell–cell fusion (Guirakhoo et al., 1991; Heinz & Roehrig, 1990; Randolph & Stollar, 1990; Summers et al., 1989). It has been reported that loss of the glycosylation site in the E protein plays a crucial role in the attenuation of WNV (Beasley et al., 2004, 2005; Shirato et al., 2004). However, this phenomenon was not found to be the case in the isolates studied here, due to the lack of putative glycosylation sites in either the parent virus or the hamster urine isolates.
Our comparative hamster experiments did not show any significant differences in the levels of HI antibody production following infection with the four viruses (Table 3). While the hamster passage 1 and 3 viruses (H9317B and H8535) resulted in viraemia levels similar to that produced by NY385-99, the passage four virus H8912 resulted in a peak viraemia on day 4 p.i., which appeared lower than that of the parent virus (Fig. 2). In terms of persistent renal infection, although initially nearly all the surviving animals shed virus in the urine for up to 2 months, only the animals in groups 2, 3 and 4 developed long-term persistent infection, compared with only 25 % of animals in group 1 (Table 4). In addition to the different clinical outcomes of the four hamster groups, pathological and immunohistochemical analyses also showed that encephalitis, accompanied by positive viral antigens, was only seen in hamsters infected with the parent virus NY385-99 and not with other viruses.
Another implication of our findings relates to viral adaptation. It appears that WNV selected by passage through hamster kidney possesses the attenuated phenotype. We speculate that the nucleotide changes identified in the hamster urine isolates are not only associated with WNV adaptation to the new host (hamster), but are also related to tissue (kidney) adaptation. To prove this point, it will be necessary to recover viruses from different organs of the same infected hamsters (i.e. brain and kidney), and to study their genetic and phenotypic differences in order to verify that the viruses recovered from brain tissue still possess their original neuroinvasiveness/virulence.
As to the source(s) of the nucleotide changes identified among the various WNV isolates, it remains to be determined if these represent de novo mutations or are the results of ‘tissue selection’. In the latter case, these ‘new’ urine isolates are merely the product of selection through a ‘tissue sieve’ effect. Presumably, the original parent virus stock used to inoculate the hamsters was a mixture of quasispecies. Evidently, the dominant one was neurotropic and possessed neuroinvasiveness, but a minority did not. This could partially explain why even with a relatively large dosage of the parent virus, not all of the inoculated hamsters developed neurological symptoms (the mortality rate was only about 30–50 %). In the current experiment, all but one of the inoculated hamsters developed viruria soon after inoculation, but only some of them remained persistently infected and continued shedding virus in their urine. It is possible that many of the quasispecies possess renal tropism, but only some of them have the ability to persistently remain in the kidney tissue. These quasispecies were selected and ‘amplified’ through urine. Nevertheless, after multiple passages, additional de novo mutations occurred, as shown by the new nucleotide changes in the hamster passage four virus (Table 1).
In summary, we have demonstrated that most of the nucleotide changes in the first-passage urine isolate H9317B were consistently inherited by its progeny viruses after additional hamster passages. At least some of these changes appear to be responsible for the change of viral phenotype, namely, adaptation to hamster renal tissue and loss of neurovirulence. The findings from these hamster infection experiments further confirmed that these urine isolates, or hamster-passaged viruses, had lost neurovirulence and had gained increased capacity for persistence of renal infection. Future studies will focus on validating individual mutations or combinations of these mutations in infectious clones, which ultimately will offer information regarding molecular determinants of viral tropism and/or virulence.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Received 16 May 2008;
accepted 9 August 2008.
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