Next Article in Journal
Type I and Type II Interferon Antagonism Strategies Used by Paramyxoviridae: Previous and New Discoveries, in Comparison
Next Article in Special Issue
Protective Immune Responses Induced by an mRNA-LNP Vaccine Encoding prM-E Proteins against Japanese Encephalitis Virus Infection
Previous Article in Journal
A Therapeutically Active Minibody Exhibits an Antiviral Activity in Oseltamivir-Resistant Influenza-Infected Mice via Direct Hydrolysis of Viral RNAs
Previous Article in Special Issue
Increased Ifng and Il10 Expression Correlate with Disease in Rodent Models Experimentally Infected with Modoc Virus
 
 
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Evidence of Infection with Zoonotic Mosquito-Borne Flaviviruses in Saltwater Crocodiles (Crocodylus porosus) in Northern Australia

1
School of Veterinary Science, The University of Queensland, Gatton, QLD 4343, Australia
2
Centre for Crocodile Research, Noonamah, NT 0837, Australia
3
School of Chemistry and Molecular Biosciences, The University of Queensland, St. Lucia, QLD 4072, Australia
4
Australian Infectious Diseases Research Centre, The University of Queensland, St Lucia, QLD 4072, Australia
*
Author to whom correspondence should be addressed.
Viruses 2022, 14(5), 1106; https://doi.org/10.3390/v14051106
Submission received: 28 April 2022 / Revised: 18 May 2022 / Accepted: 19 May 2022 / Published: 21 May 2022
(This article belongs to the Special Issue Flaviviruses and Flavivirus Vaccines)

Abstract

:
The risk of flavivirus infections among the crocodilian species was not recognised until West Nile virus (WNV) was introduced into the Americas. The first outbreaks caused death and substantial economic losses in the alligator farming industry. Several other WNV disease episodes have been reported in crocodilians in other parts of the world, including Australia and Africa. Considering that WNV shares vectors with other flaviviruses, crocodilians are highly likely to also be exposed to flaviviruses other than WNV. A serological survey for flaviviral infections was conducted on saltwater crocodiles (Crocodylus porosus) at farms in the Northern Territory, Australia. Five hundred serum samples, collected from three crocodile farms, were screened using a pan-flavivirus-specific blocking ELISA. The screening revealed that 26% (n = 130/500) of the animals had antibodies to flaviviruses. Of these, 31.5% had neutralising antibodies to WNVKUN (Kunjin strain), while 1.5% had neutralising antibodies to another important flavivirus pathogen, Murray Valley encephalitis virus (MVEV). Of the other flaviviruses tested for, Fitzroy River virus (FRV) was the most frequent (58.5%) in which virus neutralising antibodies were detected. Our data indicate that farmed crocodiles in the Northern Territory are exposed to a range of potentially zoonotic flaviviruses, in addition to WNVKUN. While these flaviviruses do not cause any known diseases in crocodiles, there is a need to investigate whether infected saltwater crocodiles can develop a viremia to sustain the transmission cycle or farmed crocodilians can be used as sentinels to monitor the dynamics of arboviral infections in tropical areas.

1. Introduction

Flaviviruses are members of the family Flaviviridae, which are mostly mosquito-borne viruses with a worldwide distribution. The family comprises four main genera, including flavivirus, pestivirus, pegivirus, and hepacivirus [1]. Flaviviruses are positive-sense, single-stranded RNA (ssRNA(+)),enveloped and monopartite viruses, with an unsegmented linear viral genome of approximately 9.4–13 kb, enclosed in an icosahedral nucleocapsid of 50nm in diameter [2]. Flaviviruses are generally transmitted by arthropods, including mosquitoes and ticks, and the final hosts are usually mammals, birds, and reptilians [1,3]. Some flaviviruses are pathogenic and can cause severe diseases in humans and animals [4]. Serologically, mosquito-borne flaviviruses are classified in multiple serocomplexes. The Japanese encephalitis virus (JEV) serocomplex comprises the JEV, West Nile (WNV), Usutu (USUV), Murray Valley encephalitis (MVEV), Yaounde, Alfuy (ALFV), and Saint Louis encephalitis (SLEV) viruses. The yellow fever virus (YFV) serocomplex comprises the YFV, Fitzroy River (FRV), Sepik (SEPV), Edge Hill (EHV), Bouboui, Banzi, Jugra, Saboya, and Potiskum viruses. YFV and SEPV are the two most closely related in the YFV serogroup. The dengue virus (DENV) serogroup comprises DENV1, DENV2, DENV3, and DENV4. The Spondweni virus complex comprises the Spondweni and Zika (Brazilian and African strains) viruses. The Kokobera virus (KOKV) serocomplex group comprises the KOKV, Stratford (STRV), and New Mapoon (NMV) viruses. The KOKV serocomplex group is closely related to the Aroa virus complex viruses that comprise the Iguape, Naranjal, and Bussuquara viruses. The Ntaya virus serocomplex comprises the Bagaza, Ntaya, Tembusu, Ilheus, and Rocio viruses.
In Australia, the most clinically significant flaviviruses that infect and cause disease in humans are MVEV, WNVKUN (Kunjin strain), DENV, and JEV [5,6,7]. Of these, most cases of DENV and JEV are imported. DENV is no longer endemic to Australia, but travel-associated outbreaks occur regularly, mainly in Northern Queensland [8]. There were JEV outbreaks in the Cape York Peninsula and Torres Strait Islands in 1995 and 1998, respectively [9,10,11]. Recently, JEV outbreaks have been reported in Victoria, Queensland, New South Wales, and South Australia. Currently, 34 human cases, of which three were fatal, have been recorded. Additionally, JEV cases have been reported in 70 piggeries across Australia [12,13,14]. However, other flaviviruses, such as KOKV, ALFV, STRV, and EHV, have been reported to be associated with a few human disease cases, usually with non-specific fevers [15,16].
MVEV is the leading cause of viral encephalitis in Australia and endemic in parts of Western Australia and the Northern Territory. The virus is maintained between aquatic birds and the vector mosquito Culex annulirostris [17,18]. There have been several outbreaks since 1974, which have all been linked to heavy rainfall, followed by increased mosquito activity [19,20]. WNVKUN is endemic in Northern and Central Australia, but it was considered to be of limited medical and veterinary importance until 2011, when it caused an equine encephalitis outbreak in south-eastern Australia. During that outbreak, the fatality rate was as high as 15–20% amongst clinically affected horses, including horses terminated for humane reasons [17,21]. Similar to MVEV, clinical cases of WNV infection have occurred in humans and horses following heavy rainfall, causing flooding and leading to favourable conditions for mosquito activity. Recently, WNVKUN was detected by RT-PCR in samples from saltwater crocodiles, in which the virus causes skin lesions that lead to hide depreciation [3,22,23]. While WNVKUN does not cause any apparent systemic diseases in saltwater crocodiles, there remains the potential for more virulent WNV strains to emerge, as has happened in the Americas during the past two decades [24,25].
KOKV infections have been reported in humans in Australia [26]. The clinical presentation for KOKV infection is similar to DENV fever; therefore, clinical diagnosis should be accompanied by laboratory testing [27]. In addition to fever and headache, infected individuals may experience influenza-like aches, myalgia, polyarthralgia syndrome, and polyarthritis [15,16,26,27].
Since its isolation in 1961 at Edge Hill, a suburb of Cairns in Northern Queensland, there has been only one report of EHV infection in humans, in which the clinical picture was characterised by myalgia, arthralgia, and fatigue [28]. Anti-EHV antibodies have been detected in wallabies, kangaroos, and bandicoots; thus, EHV is generally thought to be a virus of marsupials [29].
Antibodies to ALFV, FRV, SEPV, and STRATV have also been detected in patients who tested positive for other flaviviruses [16,26,30]. Further investigation is needed to confirm whether the neutralising antibody titres were due to actual infection or cross-reactivity with other flaviviruses. Currently, there is no evidence of human exposure to NMV; thus, it likely poses a low risk to humans [31]. Generally, individuals infected with most flaviviruses have low levels of viraemia, sometimes below detectable levels, even in clinically ill patients [32]. Therefore, serology remains the sole diagnostic method in such cases, with the virus neutralisation test being the gold standard diagnostic method [22,33].
Following our previous study demonstrating the role of WNVKUN in saltwater crocodile skin lesions [3], this investigation aimed to identify other flaviviruses that might infect saltwater crocodiles in northern Australia. Furthermore, an understanding of the potential role of crocodiles in the transmission cycle of flaviviruses and possible serological cross-reactivity between various flaviviruses, which might interfere with the specificity of flavivirus diagnostics in crocodiles and response to a WNV vaccine, is of public health and industry interest.

2. Materials and Methods

2.1. Study Area

The samples used in this study were collected from three commercial saltwater crocodile farms (Farms A, B, and C), all located in the greater Darwin region of the Northern Territory, Australia.

2.2. Animals, Blood Sample Collection, and Processing

Blood samples were collected from 500, predominantly male, farmed saltwater crocodiles at harvest when the animals were 3–4 years old. None of the sampled crocodiles had a history of clinical diseases. During harvest, crocodiles were captured using an electrical stunner that temporarily rendered the crocodile immobile and unconscious, in order to allow for safe handling, as previously described [34,35]. After immobilisation, blood was aseptically collected from the occipital sinus using 18” needles and plain (without anticoagulant) vacutainer tubes (Becton Dickinson, Franklin Lakes, NJ, USA). Blood was allowed to clot at room temperature, and it was centrifuged at room temperature for 10 min at 1000 rpm. Collected serum samples were aliquoted in triplicate into sterile Eppendorf tubes and immediately stored at −20 °C. Serum samples were heat-inactivated at 56 °C for 30 min before they were tested via blocking ELISA and virus neutralisation.

2.3. Anti-Flavivirus Blocking ELISA

The 500 crocodile serum samples were screened for the presence of flavivirus antibodies by a competitive blocking ELISA, as previously described [33,36]. Wells of SeroclusterTM U-bottom high-binding 96-well plates (Sigma Aldrich Pty Ltd. St. Louis, MO, USA) were coated with a pre-determined dilution of WNVKUN antigen (infected C6/36 cell lysate) in carbonate coating buffer (50 µL per well) and incubated overnight at 4 °C. The plates were then washed four times with phosphate-buffered saline (PBS) with 0.05% Tween-20, and nonspecific binding sites were blocked with blocking buffer (0.05 M Tris, 1 mM EDTA, 0.15 M NaCl, 0.05% (v/v) Tween-20, 0.2% (w/v) casein, and pH 8.0), followed by one-hour incubation at room temperature. On a separate plate, 5-fold dilution in the blocking buffer was performed for all the samples, including the positive and negative controls. Fifty µL of the diluted samples were transferred to the pre-coated plates in duplicate and incubated for one hour at 28 °C. Then, 50 µL of the anti-pan-flavivirus envelope protein-specific monoclonal antibody (mAb), 6B6C-1 [37], was added to the plates on top of the serum samples. The plates were incubated for a further 1 h at 28 °C. Following four washes, as above, a horseradish peroxidase (HRP)-conjugated goat anti-mouse polyclonal antibody, pre-adsorbed against human immunoglobulins and fetal bovine serum (Jackson ImmunoResearch Laboratories, West Grove, PA, USA) and diluted in the blocking buffer, was added to the plates. Following a one-hour incubation at 28°C, the plates were washed six times. The plates were then thoroughly washed, and the antibody conjugate binding was visualised via the addition of 100 µL substrate buffer (citrate-phosphate buffer, pH 4.2 supplemented with 1 mM (2,2′-azino-bis (3-ethylbenzthiazolinesulfonic acid); ABTS), and 3 mM H2O2) and incubating the plates in the dark for one hour. The enzyme activity was quantified by measuring the optical density (OD) at a wavelength of 405 nm in an automated ELISA plate reader (SpectraMax® 190 Microplate Reader, Molecular Devices, LLC, San Jose, CA, USA). The percentage of the inhibition of binding of the mAb 6B6C-1 was calculated using the following formula [36]:
Percentage of inhibition = 100 − (TS-B)/(CS-B)] × 100
where TS = OD of test serum, CS = OD of control serum, and B = background OD. Samples with an inhibition of ≥30% were considered positive. In some instances, samples giving borderline results were retested for confirmation.

2.4. Virus Culture

Nine viruses were screened in this study. These included MVEV (strain 1–51), KOKV (strain MRM32), STRATV (strain C338), ALFV (strain CY2269), FRV (strain K78296), WNV-KUNV (NSW2011), NMV (CY1014), EHV (C281), and SEPV (MK7979) (Table 1). All virus stocks were titrated by the TCID50 method, as previously described [38]. Briefly, each virus stock was diluted 10-fold in Dulbecco’s modified Eagle’s medium (DMEM; Thermofisher Scientific, Seventeen Mile Rocks, Qld, Australia) supplemented with 2% fetal bovine serum (FBS) (Gibco, Sigma Aldrich Pty Ltd., St. Louis, MO, USA), one-time penicillin–streptomycin, and 2 mM L-glutamine (Gibco, Gibco, Sigma Aldrich Pty Ltd., St. Louis, MO, USA). The diluted virus was transferred into a 96-well tissue culture plate (Costar, Corning), pre-seeded with baby hamster kidney cells (BSR) in DMEM supplemented with 5% FBS, one-time penicillin (50 U/mL), streptomycin (50 µg/mL), and 2 mM L-glutamine (Gibco, Sigma Aldrich Pty Ltd., St. Louis, MO, USA). The plates were incubated at 37 °C and 5% CO2 for five days, after which the plates were scored for cytopathic effect (CPE). The cell monolayers were fixed with 20% acetone supplemented with 0.02% (w/v) bovine serum albumin (BSA) by overnight incubation at 4 °C. The fixative was then discarded, and the plates were dried overnight at room temperature. ELISA was performed on fixed plates using the mAb 4G2 (anti-flavivirus E protein), as previously described [3,33,39]. The virus titres were expressed in TCID50/mL, as previously described [38,40].
Table 1. Summary of viruses used.
Table 1. Summary of viruses used.
Virus.StrainAccession NumberLocation of IsolationSource of IsolateYear of IsolationPropagation Cell LineReference
MVEVMVE-1-51AF161266.1Mooroopna, VictoriaFatal human case1951C6/36[41]
KOKVMRM32 strainL48969.1Kowanyama, QLDC. annulirostris and C. pullus1960C6/36[42]
STRATVSTRV C338KM225263.1Cairns, QLDAedes vigilas1961C6/36[43]
ALFVCY2269-Pompuraaw, QLDCulex annulirostris1999C6/36 and PS-EK[44,45]
FRVK78296-Fitzroy Crossing, QLDOchlerotatus normanensis2011C6/36[30]
WNVNSW2011JN887352.1New South WalesHorse brain2011C6/36[46]
NMVCY1014KC788512.1New Mapoon, QLDCulex annulirostris1998PS-EK[47]
EHVC281AF275876.2Edge Hill (Cairns) QLDCulex annulirostris1961C6/36[48]
SEPVMK7979NC_008719.1Sepik, PNG 1966PS-EK[49]

2.5. Virus Neutralisation Assays

Crocodile serum samples that were positive in the 6B6C-1 mAb blocking ELISA were tested in virus micro-neutralisation assays using the baby hamster kidney-derived cell line (BSR). Serum samples were titrated in doubling dilution down the plate (1:20 to 1:2560), which was made in plain DMEM. The virus was diluted in DMEM supplemented with 2% FBS, one-time penicillin–streptomycin, and two mM L-glutamine, with a concentration of 100 TCID50 units in 50 µL. Fifty µL of the suspension was added to each well in the 96-well plates, and the plates were incubated at 37 °C and 5% CO2 for one hour. After a one-hour incubation, 1 × 104 BSR cells, resuspended in 50 µL in the above-mentioned cell culture medium, were added to each well of the 96-well plates. The plates were incubated at 37 °C in a humidified CO2 incubator for five days. Then, the plates were scored for CPE, and the cells were fixed with the 20% acetone fixative buffer, as described above. Fixed-cell ELISA was performed using mAb 4G2 to identify virus-infected wells, and the neutralising titre was expressed as the reciprocal of the highest serum dilution where the virus replication was neutralised, i.e., there was no virus replication [36,39].

2.6. Data Analysis

Data analysis and generation of graphs were performed using GraphPad Prism 9 (GraphPad Software, Inc., San Diego, CA, USA) for Windows (version 9.2.0 (332) 2020). Neutralising antibody titres were presented as a reciprocal value corresponding to 20 being the lowest dilution and 2560 being the highest.
The criterion of four-fold greater neutralising antibody titre was used to take the potential cross-reactivities that may lead to false-positive results, in the case of samples positive for two or more flaviviruses, into account [50].

3. Results

3.1. Sample Set Description

A total of 500 serum samples were collected, of which 150 samples were collected from Farm A, 200 from Farm B, and 150 from Farm C, all of which were located near Darwin in the Northern Territory, Australia (Table S1).

3.2. Blocking ELISA Results

By pan-flavivirus blocking ELISA with the 6B6C-1 mAb, 130 (26%, n = 500) samples tested positive. From the 130 positive samples, 67 (44.7%, n = 150) were from Farm A, with 20.5% (n = 200) from Farm B and 14.7% (n = 150) from Farm C (Figure 1 and Table S1).

3.3. Virus Neutralisation Assay

When the subset (n = 130) of the samples that were positive in the 6B6C-1 blocking ELISA were tested using VNT, 76 samples (58.5%) had neutralising antibodies to FRV. Of these, 35 (52.2%, n = 67) samples were from Farm A, 25 (60.9%, n = 41) were from Farm B, and 16 (72.7%, n = 22) were from Farm C. Fifty-three of the FRV positive samples had a neutralising titre of 20, 21 samples had a titre of 40, three samples had a titre of 80, and three samples had a titre of 160. Forty-one samples (31.5%, n = 130) had neutralising activity for WNV. Of the 41 samples, 29 (43.3%, n = 67) were from Farm A, 9 (22.0%, n = 41) from Farm B, and 3 (13.6%, n = 22) from Farm C. The WNV neutralising titres varied between 20 and 2560. Twenty-four (18.5%, n = 130) samples had neutralising antibodies to STRATV, of which 22 (16.9%, n = 67) came from Farm A, with one from Farm B (4.5%, n = 41) and one from Farm C (4.5%, n = 22). SEPV was the fourth most frequent of the tested flaviviruses at nine (6.9%, n = 130). At the farm level, neutralising antibodies to SEPV were detected in six samples (9.0%, n = 67) at Farm A, with two samples (4.9%, n = 41) at Farm B and one sample (4.5%, n = 22) at Farm C. The highest titre observed was 640, in a sample from Farm A. Neutralising antibodies to MVEV, EHV, and ALFV were each present in two samples (1.5%, n=130), and they were all from Farm A. Neutralising antibodies to KOKV and NMV were each present in one sample (0.8%, n = 130), both from Farm A.
Overall, based on the initial sample size of 500, composed of 150 samples from Farm A, 200 from Farm B, and 150 from Farm C, samples with neutralising antibodies to FRV were the most frequent, followed by WNV. The least frequently detected viruses were KOKV and NMV (Figure 2).
Figure 2. Proportion of samples that have neutralising antibodies against various viruses tested in the virus neutralisation assay. The overall frequency was determined based on the 500 samples collected from three crocodile farms. The frequency per farm was estimated based on n = 150 at Farm A, n = 200 at Farm B, and n = 150 at Farm C. Of the 130 samples positive in the 6B6C-1 blocking ELISA, 27 (20.8%, n = 130) did not have neutralising antibodies to any of the tested viruses. They were designated as unknown flaviviruses. Farm B had the highest number of animals positive for unknown flaviviruses (n = 12), followed by Farm A (n = 9) and Farm C (n = 6) (Figure 3 and Table S1).
Figure 2. Proportion of samples that have neutralising antibodies against various viruses tested in the virus neutralisation assay. The overall frequency was determined based on the 500 samples collected from three crocodile farms. The frequency per farm was estimated based on n = 150 at Farm A, n = 200 at Farm B, and n = 150 at Farm C. Of the 130 samples positive in the 6B6C-1 blocking ELISA, 27 (20.8%, n = 130) did not have neutralising antibodies to any of the tested viruses. They were designated as unknown flaviviruses. Farm B had the highest number of animals positive for unknown flaviviruses (n = 12), followed by Farm A (n = 9) and Farm C (n = 6) (Figure 3 and Table S1).
Viruses 14 01106 g002
Thirty-nine samples had neutralising antibodies to more than one of the flaviviruses tested for (Figure 4 and Table S2). Twenty-six samples (20%, n = 130) had neutralising antibodies to any two tested flaviviruses, while 13 (10%, n = 130) had neutralising antibodies to any three of the tested flaviviruses.
The verdict (diagnosis) for VNT was based on the analysis of the set of neutralising antibodies to the virus from the same serogroup. Out of the 130 flavivirus-positive samples, 16 samples (12.3%) had neutralising antibodies to both WNV and FRV, three samples (2.3%) had antibodies to WNV and STRATV, three samples (2.3%) had antibodies to FRV and STRATV, two samples (1.5%) had antibodies to FRV and EHV, one sample (0.8%) had antibodies to WNV and SEPV, one sample (0.8%) had antibodies to ALV and FRV, one sample (0.8%) had antibodies to STRATV and SEPV, one sample (0.8%) had antibodies to FRV and SEPV, and one sample (0.8%) had antibodies to MVEV and FRV.
Seven samples out of the 130 flavivirus-positive sera (5.4%) had antibodies to FRV, WNV, and STRATV, two samples (1.5%) had antibodies to WNV, STRATV, and SEPV, one sample (0.8%) had antibodies to NMV, STRATV, and WNV, one sample (0.8%) had antibodies to FRV, STRATV, and SEPV, one sample (0.8%) had antibodies to ALV, FRV, and STRATV, and one sample (0.8%) had antibodies to FRV, MVEV, and STRATV.
Cross-reactivity between viruses of the same serogroup was often observed, as opposed to coinfection (Table S1). For example, sample A131 was positive for ALFV, with a neutralisation titre of 320 and comparably low titres of 20 (16-fold less) to the closely related flaviviruses MVEV and WNV. This trend was observed for other flaviviruses of the same serogroup (Table S1). In contrast, we observed cases of possible coinfection, where samples had similar neutralising antibody titres to viruses of the same serogroup or distant serogroups. Sample A69 gave, by far, the strongest neutralising antibody titre to SEPV at 640, but it also had strong neutralising titres against WNV, FRV, and STRATV. There were more examples of such instances (Table S1).

4. Discussion

WNV has been reported in crocodilians, including alligators (Alligator mississippiensis) in the Americas, Nile crocodiles (Crocodylus niloticus) in Africa, Israel, and Mexico [51,52], and saltwater crocodiles (Crocodylus porosus) in Australia [3,23]. This is the first study to provide serological evidence of infections with zoonotic flaviviruses, other than WNV, in saltwater crocodiles in Australia. Apart from WNV, none of the other flaviviruses tested for have ever been reported in crocodilian species.
Most serological studies, so far, have focused on mammalian and avian species; thus, this is the first study of the seroepidemiology of flaviviruses infection in reptilians, and it included a substantial sample size. Antibodies to most of the flaviviruses tested in this study have been detected in other hosts, mostly avian and mammalian species. However, while data from this study provides general information on zoonotic flaviviruses circulating in farmed saltwater crocodiles in the Northern Territory, it does not necessarily represent the true prevalence of these viruses. Considering that flavivirus infections are seasonally- and locality-dependent, understanding the true prevalence, dynamism, and risk factors of the tested viruses would require sampling crocodiles of different ages and sex in different seasons, with varying rainfall and more locations, as well as sampling wild-caught crocodiles. Since crocodiles are farmed for skin and hides and harvested when they are three to four years old, they would have been through at least three wet seasons.
Most of the studied flaviviruses have been associated with clinical and subclinical infections in various hosts. These hosts include humans, birds, horses, and European rabbits [53]. Antibodies to ALFV, FRV, and STRATV have been detected in humans, horses, and cattle [30,54]. In contrast, the detection of neutralizing antibodies for NMV is puzzling, given that no vertebrate host has been identified since its isolation from the C. annulirostris mosquitoes collected in 1998 on the Cape York Peninsula, Queensland [55]. It would be important to investigate the main host(s) for this virus.
Intriguingly, some samples had neutralising antibodies to SEPV, a virus that has never been reported in Australia. The virus was first isolated from the Mansonia septempunctata collected from the Sepik district of Papua New Guinea (PNG). The virus was then isolated in 1998 from Balimo in Western Province of PNG, a place relatively closer to Australian mainland [56]. Since then, the virus has been thought to be restricted to PNG [57]. While the spread of flaviviruses from one location to another can result from international travel, there would always need to be a vector of the virus that would sustain the infection locally. Mansonia septempunctata, the mosquito vector for SEPV, is not known to occur in the Northern Territory [58]. Most SEPV-positive samples had a neutralising antibody titre varying between 20 and 40. Additionally, three-quarters of the samples positive for SEPV, which had a neutralising titre of 40 and above, were positive for FRV, another flavivirus belonging to the YFV serogroup. Moreover, another virus of the YFV serogroup, the Bamaga virus (BgV), has been reported in Australia [59]. While this virus was not tested for in this study, the possibility of it infecting crocodiles cannot be ruled out, since infection in vertebrates was reported earlier [59,60]. Whether the detection of neutralising antibodies to SEPV is linked to an infection or result of cross-reactivity with another flavivirus of the YFV serogroup, such as FRV, BgV, or any other unknown flavivirus, would require further investigation.
It is not unexpected that a relatively high number of samples tested positive for two or more flaviviruses. This scenario can be explained by the fact that most of the studied flaviviruses are transmitted by the same vectors [61]. Both MVEV and WNV, the most clinically significant flaviviruses in Australia, are mainly transmitted by Culex annulirostris. Similarly, KOKV and NMV have also been isolated from Cx. annulirostris, which is abundant in the Northern Territory and, therefore, likely to have transmitted these viruses to the crocodiles. STRV has been isolated from, and is transmitted by, Aedes spp. (Aedes aculeatus, Ae. alternans, Ae. notoscriptus, Ae. procax, and Ae. vigilax) and Anopheles annulipes [61,62,63]. EHV has also been isolated from two mosquito species, including Cx. annulirostris and Ae. vigilax [64]. Neutralising antibodies to FRV were the most frequently detected in analysed samples. This finding is not surprising, considering the ecology of FRV and its principal vector Ae. normanensis, a mosquito species that is abundantly prevalent in northern Australia. The maintenance of the Ae. normanensis population requires dry areas, and its activity can significantly increase after rainfall. Thus, these environmental conditions are fulfilled in northern Australia [30,65].
Another possibility for the occurrence of antibodies to a variety of flaviviruses is the cross-reactive immunity among flaviviruses from the same serogroup [66]. Some samples that were positive for WNV were also positive for MVEV and ALFV. The same phenomenon was observed for flaviviruses from the KOKV and YFV groups (Table S1). Since the criteria of a four-fold greater neutralising antibody titre was used to take the potential cross-reactivities into account [50], co-infection and superinfection with various flaviviruses are likely possible in saltwater crocodiles.
The high occurrence of samples with neutralising antibodies to FRV and occurrence of those with dual neutralising antibodies to FRV and WNV are interesting findings that should be investigated further, in the context of the development of diagnostic tools for WNV [67]. The relatively high occurrence of samples positive in the pan-flavivirus ELISA using mAb 6B6C-1, but with no neutralising antibodies to any of the tested flaviviruses, suggests the circulation of unknown vertebrate infecting flaviviruses, which also infect saltwater crocodiles. This finding could be further explored via elucidation of the virome of the crocodiles and mosquitoes abundantly found on these farms, using deep sequencing and transcriptomic analysis [68].
The main limitations of this study were the similar age of all the animals and predominance of males. Crocodiles are harvested for their hides when they reach a particular size, primarily between three to four years of age. Due to the lack of sex chromosomes, crocodile sex is determined by egg incubation temperature, which happen to favour males. Additionally, most crocodile farmers prefer rearing faster-growing males, which was probably the case with the cohort we sampled in this study [69,70]; therefore, it was impossible to investigate the additional risk factors associated with the seroconversion to the studied flaviviruses. Hence, while this study gives a snapshot of the flaviviruses circulating in crocodiles in the Northern Territory of Australia, it is not necessarily a complete representation of the prevalence of the studied mosquito-borne flaviviruses. To better investigate the dynamics, it would be necessary to also study ecological factors, such as season, rainfall (including draught events), and humidity [71], as well as wild versus farmed crocodiles. A longitudinal study sampling crocodiles hatched and or slaughtered in different seasons and additional regions, such as northern Queensland, might shed further light on the various risk factors.

5. Conclusions

The present study shows that farmed crocodiles from the Northern Territory are exposed to a variety of zoonotic flaviviruses, other than WNV. The detection of neutralising antibodies to more than one virus in one host suggests co-infection with different viruses at a given time point or animals that were consecutively exposed during their lifetime. Considering the level of neutralising titres for some viruses, it would be worth investigating the pathogenesis of infection in crocodiles with the viruses of “One Health” importance, notably MVEV. Studying the pathogenesis will help our understanding of the role of crocodiles in the transmission cycle, i.e., whether they can develop a viremia high enough to sustain the transmission cycle or are dead-end hosts.
Most of the viruses used in this study have been linked to clinical and subclinical infections in humans in Australia and other Pacific regions; thus, they all are clinically significant in the ”One Health” context. Therefore, saltwater crocodile can be used as a sentinel species to monitor arboviral infection dynamics and predict flaviviruses outbreaks in humans and animals using the “One Health” approach.

Supplementary Materials

The following supporting information can be downloaded at: https://0-www-mdpi-com.brum.beds.ac.uk/article/10.3390/v14051106/s1. Table S1: Summary data of 6B6C-1 blocking ELISA and virus neutralisation test. Table S2: Frequency of co-infection or cross-reactivity among tested samples.

Author Contributions

Conceptualisation, H.B.-O., R.A.H. and G.H.; methodology, H.B.-O., R.A.H., G.H. and S.R.I.; validation, H.B.-O., G.H., S.R.I. and R.A.H.; formal analysis, G.H., J.H.-P., R.A.H. and H.B.-O.; investigation, G.H., J.J.H., J.M., S.R.I. and H.B.-O.; resources, G.H., S.R.I., J.H.-P., R.A.H. and H.B.-O.; data curation, G.H. and H.B.-O.; writing—original draft preparation, G.H. and H.B.-O.; writing—review and editing, all authors; visualisation, G.H. and H.B.-O.; supervision, J.H.-P., R.A.H., S.R.I. and H.B.-O.; project administration, H.B.-O. and S.R.I.; funding acquisition, S.R.I., R.A.H., J.H.-P., G.H. and H.B.-O. All authors have read and agreed to the published version of the manuscript.

Funding

This research has been funded by the Cooperative Research Centre for Developing Northern Australia (CRC-DNA; S.R.I., R.A.H., J.H.-P. and H.B.-O.), National Health and Medical Research Council (NHMRC) of Australia (grant no. 1146237. R.A.H., H.B.-O. and J.H.-P), and SVS Bequest Fund (H.B-O. and G.H.), as well as a donation from BIOHMPATHOLOGY. G.H. is a recipient of an Australian Postgraduate Scholarship Award (2018–2022), Adam Gillespie Memorial Scholarship for Animal Husbandry, the IUCN-SSC Crocodile Specialist Group Fritz Huchzermeyer Veterinary Science Student Research Assistance Scheme (FHVS-SRAS), and Fred Z. Eager Award. J.H.-P. is supported by an Advance Queensland Industry Research Fellowship.

Institutional Review Board Statement

This study was conducted in accordance with the University of Queensland research ethical guidelines. Ethical approval for this study was obtained from the University of Queensland’s Native and Exotic Wildlife and Marine Animal Ethics Committee (SVS/354/20/NT). The Centre for Crocodile Research has a license to conduct research on animals with the animal welfare unit of the Northern Territory government (permit no. 061). During sample collection, animal care and use protocols adhered to the Animal Welfare Regulations 2000 of the Northern Territory of Australia and Code of Practice on the Humane Treatment of Wild and Farmed Australian Crocodiles.

Informed Consent Statement

Not applicable.

Data Availability Statement

All relevant data are included in the report and supplementary data.

Acknowledgments

The authors would like to thank the management and staff on crocodile farms for providing access to and help with the sampling of crocodile blood at scheduled slaughter.

Conflicts of Interest

All authors declare no conflict of interest. The funders had no role in the study design, collection, analysis, or interpretation of data; in the writing of the manuscript, or in the decision to publish this article.

References

  1. Simmonds, P.; Becher, P.; Bukh, J.; Gould, E.A.; Meyers, G.; Monath, T.; Muerhoff, S.; Pletnev, A.; Rico-Hesse, R.; Smith, D.B.; et al. ICTV Virus Taxonomy Profile: Flaviviridae. J. Gen. Virol. 2017, 98, 2–3. [Google Scholar] [CrossRef] [PubMed]
  2. Chen, S.; Wu, Z.; Wang, M.; Cheng, A. Innate immune evasion mediated by flaviviridae non-structural proteins. Viruses 2017, 9, 291. [Google Scholar] [CrossRef] [PubMed]
  3. Habarugira, G.; Moran, J.; Colmant, A.M.G.; Davis, S.S.; O’Brien, C.A.; Hall-Mendelin, S.; McMahon, J.; Hewitson, G.; Nair, N.; Barcelon, J.; et al. Mosquito-Independent Transmission of West Nile virus in Farmed Saltwater Crocodiles (Crocodylus porosus). Viruses 2020, 12, 198. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  4. Mackenzie, J.; Hall, R.; Lindsay, M.; Bielefeldt-Ohmann, H.; Poidinger, M. Molecular epidemiology and phylogeny of Australian arboviruses. In Molecular Epidemiology of Infectious Diseases; Hodder Education: Cambridge, UK, 2000; pp. 297–315. [Google Scholar]
  5. Broom, A.K. Sentinel Chicken Surveillance Program in Australia, July 2002 to June 2003. Commun. Dis. Intell. Q. Rep. 2003, 27, 367–369. [Google Scholar] [PubMed]
  6. Harapan, H.; Michie, A.; Sasmono, R.T.; Imrie, A. Dengue: A Minireview. Viruses 2020, 12, 829. [Google Scholar] [CrossRef] [PubMed]
  7. Pyke, A.T.; Smith, I.L.; Van Den Hurk, A.F.; Northill, J.A.; Chuan, T.F.; Westacott, A.J.; Smith, G.A. Detection of Australasian Flavivirus encephalitic viruses using rapid fluorogenic TaqMan RT-PCR assays. J. Virol. Methods 2004, 117, 161–167. [Google Scholar] [CrossRef]
  8. Webb, C.E.; Porigneaux, P.G.; Durrheim, D.N. Assessing the Risk of Exotic Mosquito Incursion through an International Seaport, Newcastle, NSW, Australia. Trop. Med. Infect. Dis. 2021, 6, 25. [Google Scholar] [CrossRef]
  9. Hanna, J.N.; Ritchie, S.A.; Phillips, D.A.; Lee, J.M.; Hills, S.L.; van den Hurk, A.F.; Pyke, A.T.; Johansen, C.A.; Mackenzie, J.S. Japanese encephalitis in north Queensland, Australia, 1998. Med. J. Aust. 1999, 170, 533–536. [Google Scholar] [CrossRef]
  10. Johansen, C.A.; van den Hurk, A.F.; Pyke, A.T.; Zborowski, P.; Phillips, D.A.; Mackenzie, J.S.; Ritchie, S.A. Entomological investigations of an outbreak of Japanese encephalitis virus in the Torres Strait, Australia, in 1998. J. Med. Entomol. 2001, 38, 581–588. [Google Scholar] [CrossRef] [Green Version]
  11. van den Hurk, A.F.; Pyke, A.T.; Mackenzie, J.S.; Hall-Mendelin, S.; Ritchie, S.A. Japanese Encephalitis Virus in Australia: From Known Known to Known Unknown. Trop. Med. Infect. Dis. 2019, 4, 38. [Google Scholar] [CrossRef] [Green Version]
  12. Anonymous. Japanese Encephalitis Update. Available online: https://www.ava.com.au/news/japanese-encephalitis-update-23rd-march-2022/ (accessed on 25 April 2022).
  13. Friedman, D. Japanese Encephalitis Virus Detected in Victoria. Available online: https://www.health.vic.gov.au/health-advisories/japanese-encephalitis-virus-detected-in-victoria (accessed on 25 April 2022).
  14. Anonymous. Japanese Encephalitis. Available online: https://www.outbreak.gov.au/current-responses-to-outbreaks/japanese-encephalitis#:~:text=In%20February%202022%2C%20Japanese%20encephalitis,piggeries%20across%20the%20four%20states (accessed on 25 April 2022).
  15. Boughton, C.R.; Hawkes, R.A.; Naim, H.M. Illness caused by a Kokobera-like virus in south-eastern Australia. Med. J. Aust. 1986, 145, 90–92. [Google Scholar] [CrossRef] [PubMed]
  16. Hawkes, R.A.; Pamplin, J.; Boughton, C.R.; Naim, H.M. Arbovirus infections of humans in high-risk areas of south-eastern Australia: A continuing study. Med. J. Aust. 1993, 159, 159–162. [Google Scholar] [CrossRef] [PubMed]
  17. Roche, S.; Wicks, R.; Garner, M.; East, I.; Paskin, R.; Moloney, B.; Carr, M.; Kirkland, P. Descriptive overview of the 2011 epidemic of arboviral disease in horses in Australia. Aust. Vet. J. 2013, 91, 5–13. [Google Scholar] [CrossRef] [PubMed]
  18. Selvey, L.A.; Dailey, L.; Lindsay, M.; Armstrong, P.; Tobin, S.; Koehler, A.P.; Markey, P.G.; Smith, D.W. The changing epidemiology of Murray Valley encephalitis in Australia: The 2011 outbreak and a review of the literature. PLoS Negl. Trop. Dis. 2014, 8, e2656. [Google Scholar] [CrossRef] [Green Version]
  19. Selvey, L.A.; Johansen, C.A.; Broom, A.K.; Antão, C.; Lindsay, M.D.; Mackenzie, J.S.; Smith, D.W. Rainfall and sentinel chicken seroconversions predict human cases of Murray Valley encephalitis in the north of Western Australia. BMC Infect. Dis. 2014, 14, 672. [Google Scholar] [CrossRef] [Green Version]
  20. Brown, A.; Bolisetty, S.; Whelan, P.; Smith, D.; Wheaton, G. Reappearance of human cases due to Murray Valley encephalitis virus and Kunjin virus in central Australia after an absence of 26 years. Commun. Dis. Intell. Q. Rep. 2002, 26, 39–44. [Google Scholar]
  21. Frost, M.J.; Zhang, J.; Edmonds, J.H.; Prow, N.A.; Gu, X.; Davis, R.; Hornitzky, C.; Arzey, K.E.; Finlaison, D.; Hick, P. Characterization of virulent west nile virus kunjin strain, australia, 2011. Emerg. Infect. Dis. 2012, 18, 792–800. [Google Scholar] [CrossRef]
  22. Habarugira, G.; Suen, W.W.; Hobson-Peters, J.; Hall, R.A.; Bielefeldt-Ohmann, H. West Nile Virus: An Update on Pathobiology, Epidemiology, Diagnostics, Control and “One Health” Implications. Pathogens 2020, 9, 589. [Google Scholar] [CrossRef]
  23. Isberg, S.; Moran, J.; De Araujo, R.; Elliott, N.; Davis, S.; Melville, L. First evidence of Kunjin strain of West Nile virus associated with saltwater crocodile (Crocodylus porosus) skin lesions. Aust. Vet. J. 2019, 97, 390–393. [Google Scholar] [CrossRef]
  24. Hadfield, J.; Brito, A.F.; Swetnam, D.M.; Vogels, C.B.F.; Tokarz, R.E.; Andersen, K.G.; Smith, R.C.; Bedford, T.; Grubaugh, N.D. Twenty years of West Nile virus spread and evolution in the Americas visualized by Nextstrain. PLoS Pathog. 2019, 15, e1008042. [Google Scholar] [CrossRef] [Green Version]
  25. Petersen, L.R.; Hayes, E.B. West Nile virus in the Americas. Med. Clin. N. Am. 2008, 92, 1307–1322. [Google Scholar] [CrossRef] [PubMed]
  26. Hawkes, R.A.; Boughton, C.R.; Naim, H.M.; Wild, J.; Chapman, B. Arbovirus infections of humans in New South Wales. Seroepidemiology of the flavivirus group of togaviruses. Med. J. Aust. 1985, 143, 555–561. [Google Scholar] [CrossRef] [PubMed]
  27. Mein, J.; O’Grady, K.-A.; Whelan, P.; Merianos, A. Dengue or Kokobera? A case report from the Top End of the Northern Territory. Commun. Dis. Intell. 1998, 22, 105–107. [Google Scholar] [PubMed]
  28. Aaskov, J.; Phillips, D.; Wiemers, M. Possible clinical infection with Edge Hill virus. Trans. R. Soc. Trop. Med. Hyg. 1993, 87, 452–453. [Google Scholar] [CrossRef]
  29. Doherty, R.; Carley, J.; Gorman, B.; Buchanan, P.; Welch, J.; Whitehead, R. Studies of Arthropod-Born Virus Infections in Queensland. Aust. J. Exp. Biol. Med. Sci. 1964, 42, 149–164. [Google Scholar] [CrossRef]
  30. Johansen, C.A.; Williams, S.H.; Melville, L.F.; Nicholson, J.; Hall, R.A.; Bielefeldt-Ohmann, H.; Prow, N.A.; Chidlow, G.R.; Wong, S.; Sinha, R.; et al. Characterization of Fitzroy River Virus and Serologic Evidence of Human and Animal Infection. Emerg. Infect. Dis. 2017, 23, 1289–1299. [Google Scholar] [CrossRef] [Green Version]
  31. Dunstan, R.A.; Seed, C.R.; Keller, A.J. Emerging viral threats to the Australian blood supply. Aust. N. Z. J. Public Health 2008, 32, 354–360. [Google Scholar] [CrossRef]
  32. Cleton, N.B.; Godeke, G.J.; Reimerink, J.; Beersma, M.F.; Doorn, H.R.; Franco, L.; Goeijenbier, M.; Jimenez-Clavero, M.A.; Johnson, B.W.; Niedrig, M.; et al. Spot the difference-development of a syndrome based protein microarray for specific serological detection of multiple flavivirus infections in travelers. PLoS Negl. Trop. Dis. 2015, 9, e0003580. [Google Scholar] [CrossRef]
  33. Prow, N.; Tan, C.; Wang, W.; Hobson-Peters, J.; Kidd, L.; Barton, A.; Wright, J.; Hall, R.; Bielefeldt-Ohmann, H. Natural exposure of horses to mosquito-borne flaviviruses in south-east Queensland, Australia. Int. J. Environ. Res. Public Health 2013, 10, 4432–4443. [Google Scholar] [CrossRef] [Green Version]
  34. Franklin, C.E.; Davis, B.M.; Peucker, S.K.; Stephenson, H.; Mayer, R.; Whittier, J.; Lever, J.; Grigg, G.C. Comparison of stress induced by manual restraint and immobilisation in the estuarine crocodile, Crocodylus porosus. J. Exp. Zool. A Comp. Exp. Biol. 2003, 298, 86–92. [Google Scholar] [CrossRef]
  35. Pfitzer, S.; Ganswindt, A.; Fosgate, G.T.; Botha, P.J.; Myburgh, J.G. Capture of farmed Nile crocodiles (Crocodylus niloticus): Comparison of physiological parameters after manual capture and after capture with electrical stunning. Vet. Rec. 2014, 175, 304. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  36. Hall, R.; Broom, A.; Hartnett, A.; Howard, M.; Mackenzie, J. Immunodominant epitopes on the NS1 protein of MVE and KUN viruses serve as targets for a blocking ELISA to detect virus-specific antibodies in sentinel animal serum. J. Virol. Methods 1995, 51, 201–210. [Google Scholar] [CrossRef]
  37. Roehrig, J.T.; Mathews, J.H.; Trent, D.W. Identification of epitopes on the E glycoprotein of Saint Louis encephalitis virus using monoclonal antibodies. Virology 1983, 128, 118–126. [Google Scholar] [CrossRef]
  38. May, F.J.; Lobigs, M.; Lee, E.; Gendle, D.J.; Mackenzie, J.S.; Broom, A.K.; Conlan, J.V.; Hall, R.A. Biological, antigenic and phylogenetic characterization of the flavivirus Alfuy. J. Gen. Virol. 2006, 87, 329–337. [Google Scholar] [CrossRef]
  39. Vet, L.J.; Setoh, Y.X.; Amarilla, A.A.; Habarugira, G.; Suen, W.W.; Newton, N.D.; Harrison, J.J.; Hobson-Peters, J.; Hall, R.A.; Bielefeldt-Ohmann, H. Protective Efficacy of a Chimeric Insect-Specific Flavivirus Vaccine against West Nile Virus. Vaccines 2020, 8, 258. [Google Scholar] [CrossRef]
  40. Reed, L.J.; Muench, H. A simple method of estimating fifty per cent endpoints. Am. J. Epidemiol. 1938, 27, 493–497. [Google Scholar] [CrossRef]
  41. French, E.L. Murray Valley encephalitis isolation and characterization of the aetiological agent. Med. J. Aust. 1952, 1, 100–103. [Google Scholar] [CrossRef]
  42. Doherty, R.L.; Carley, J.G.; Kay, B.H.; Filippich, C.; Marks, E.N.; Frazier, C.L. Isolation of virus strains from mosquitoes collected in Queensland, 1972-1976. Aust. J. Exp. Biol. Med. Sci. 1979, 57, 509–520. [Google Scholar] [CrossRef]
  43. Warrilow, D.; Hall-Mendelin, S.; Hobson-Peters, J.; Prow, N.A.; Allcock, R.; Hall, R.A. Complete coding sequences of three members of the kokobera group of flaviviruses. Genome Announc. 2014, 2. [Google Scholar] [CrossRef] [Green Version]
  44. Doherty, R.L.; Whitehead, R.H.; Wetters, E.J.; Gorman, B.M. Studies of the epidemiology of arthropod-borne virus infections at Mitchell River Mission, Cape York Peninsula, North Queensland: II. Arbovirus infections of mosquitoes, man and domestic fowls, 1963–1966. Trans. R. Soc. Trop. Med. Hyg. 1968, 62, 430–438. [Google Scholar] [CrossRef]
  45. Westlake, D.; Bielefeldt-Ohmann, H.; Prow, N.A.A.; Hall, R.A.A. Novel Flavivirus Attenuation Markers Identified in the Envelope Protein of Alfuy Virus. Viruses 2021, 13, 147. [Google Scholar] [CrossRef] [PubMed]
  46. Prow, N.A.; Edmonds, J.H.; Williams, D.T.; Setoh, Y.X.; Bielefeldt-Ohmann, H.; Suen, W.W.; Hobson-Peters, J.; van den Hurk, A.F.; Pyke, A.T.; Hall-Mendelin, S. Virulence and evolution of West Nile virus, Australia, 1960–2012. Emerg. Infect. Dis. 2016, 22, 1353. [Google Scholar] [CrossRef] [PubMed]
  47. May, F.J.; Clark, D.C.; Pham, K.; Diviney, S.M.; Williams, D.T.; Field, E.J.; Kuno, G.; Chang, G.J.; Cheah, W.Y.; Setoh, Y.X.; et al. Genetic divergence among members of the Kokobera group of flaviviruses supports their separation into distinct species. J. Gen. Virol. 2013, 94, 1462–1467. [Google Scholar] [CrossRef] [PubMed]
  48. Doherty, R.L.; Carley, J.G.; Mackerras, M.J.; Marks, E.N. Studies of arthropod-borne virus infections in Queensland. III. Isolation and characterization of virus strains from wild-caught mosquitoes in North Queensland. Aust. J. Exp. Biol. Med. Sci. 1963, 41, 17–39. [Google Scholar] [CrossRef]
  49. Kuno, G.; Chang, G.J. Characterization of Sepik and Entebbe bat viruses closely related to yellow fever virus. Am. J. Trop. Med. Hyg. 2006, 75, 1165–1170. [Google Scholar] [CrossRef] [PubMed]
  50. Pauvolid-Corrêa, A.; Campos, Z.; Juliano, R.; Velez, J.; Nogueira, R.M.; Komar, N. Serological evidence of widespread circulation of West Nile virus and other flaviviruses in equines of the Pantanal, Brazil. PLoS Negl. Trop. Dis. 2014, 8, e2706. [Google Scholar] [CrossRef] [Green Version]
  51. Ariel, E. Viruses in reptiles. Vet. Res. 2011, 42, 100. [Google Scholar] [CrossRef] [Green Version]
  52. Steinman, A.; Banet-Noach, C.; Tal, S.; Levi, O.; Simanov, L.; Perk, S.; Malkinson, M.; Shpigel, N. West Nile virus infection in crocodiles. Emerg. Infect. Dis. 2003, 9, 887–889. [Google Scholar] [CrossRef] [Green Version]
  53. Prow, N.A.; Hewlett, E.K.; Faddy, H.M.; Coiacetto, F.; Wang, W.; Cox, T.; Hall, R.A.; Bielefeldt-Ohmann, H. The Australian Public is Still Vulnerable to Emerging Virulent Strains of West Nile Virus. Front. Public Health 2014, 2, 146. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Gyawali, N.; Bradbury, R.S.; Aaskov, J.G.; Taylor-Robinson, A.W. Neglected Australian Arboviruses and Undifferentiated Febrile Illness: Addressing Public Health Challenges Arising From the ‘Developing Northern Australia’ Government Policy. Front. Microbiol. 2017, 8, 2150. [Google Scholar] [CrossRef]
  55. Blasi, A.; Lo Presti, A.; Cella, E.; Angeletti, S.; Ciccozzi, M. The phylogenetic and evolutionary history of Kokobera virus. Asian Pac. J. Trop. Med. 2016, 9, 968–972. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  56. Nisbet, D.J.; Hall, R.A.; Johansen, C.A.; Macdonald, J.; Pam, R.; Ritchie, S.A.; Saleh, S.; van den Hurk, A.F.; Zborowski, P.; Mackenzie, J.S. Isolation of Sepik vims from Papua New Guinea. Arbovirus Res. Aust. 2001, 8, 263–266. [Google Scholar]
  57. Jonduo, M.H.; Bande, G.; Horwood, P.F. Arboviruses of human health significance in Papua New Guinea. PNG Med. J. 2012, 55, 35–44. [Google Scholar]
  58. Webb, C.E.; Doggett, S.L.; Ritchie, S.A.; Russell, R.C. Vector competence of three Australian mosquitoes, Verrallina carmenti, Verraullina lineata, and Mansonia septempunctata (Diptera: Culicidae), for Ross River virus. J. Med. Entomol. 2008, 45, 737–740. [Google Scholar] [CrossRef]
  59. Colmant, A.M.G.; Bielefeldt-Ohmann, H.; Hobson-Peters, J.; Suen, W.W.; O’Brien, C.A.; van den Hurk, A.F.; Hall, R.A. A newly discovered flavivirus in the yellow fever virus group displays restricted replication in vertebrates. J. Gen. Virol. 2016, 97, 1087–1093. [Google Scholar] [CrossRef]
  60. Colmant, A.M.G.; Bielefeldt-Ohmann, H.; Vet, L.J.; O’Brien, C.A.; Bowen, R.A.; Hartwig, A.E.; Davis, S.; Piyasena, T.B.H.; Habarugira, G.; Harrison, J.J.; et al. NS4/5 mutations enhance flavivirus Bamaga virus infectivity and pathogenicity in vitro and in vivo. PLoS Negl. Trop. Dis. 2020, 14, e0008166. [Google Scholar] [CrossRef] [Green Version]
  61. Thomas, S.J.; Martinez, L.J.; Endy, T.P. Flaviviruses: Yellow Fever, Japanese B, West Nile, and Others. In Viral Infections of Humans: Epidemiology and Control; Kaslow, R.A., Stanberry, L.R., Le Duc, J.W., Eds.; Springer: Boston, MA, USA, 2014; pp. 383–415. [Google Scholar]
  62. Russell, R.C. Vectors vs. humans in Australia--who is on top down under? An update on vector-borne disease and research on vectors in Australia. J. Soc. Vector Ecol. 1998, 23, 1–46. [Google Scholar]
  63. Russell, R.C.; Kay, B.H. Medical entomology: Changes in the spectrum of mosquito-borne disease in Australia and other vector threats and risks, 1972–2004. Aust. J. Entomol. 2004, 43, 271–282. [Google Scholar] [CrossRef]
  64. van den Hurk, A.F.; Nisbet, D.J.; Foley, P.N.; Ritchie, S.A.; Mackenzie, J.S.; Beebe, N.W. Isolation of arboviruses from mosquitoes (Diptera: Culicidae) collected from the Gulf Plains region of northwest Queensland, Australia. J. Med. Entomol. 2002, 39, 786–792. [Google Scholar] [CrossRef]
  65. Warchot, A. Mt Todd Mine Baseline Biting Insect Assessment, February 2012 to January 2013; Medical Entomology Department of Health: Casuarina, NT, Australia, 2013. [Google Scholar]
  66. Rathore, A.P.S.; St John, A.L. Cross-Reactive Immunity Among Flaviviruses. Front. Immunol. 2020, 11, 334. [Google Scholar] [CrossRef] [Green Version]
  67. Hobson-Peters, J.; Harrison, J.J.; Watterson, D.; Hazlewood, J.E.; Vet, L.J.; Newton, N.D.; Warrilow, D.; Colmant, A.M.G.; Taylor, C.; Huang, B.; et al. A recombinant platform for flavivirus vaccines and diagnostics using chimeras of a new insect-specific virus. Sci. Transl. Med. 2019, 11, eaax7888. [Google Scholar] [CrossRef] [PubMed]
  68. Harvey, E.; Rose, K.; Eden, J.S.; Lo, N.; Abeyasuriya, T.; Shi, M.; Doggett, S.L.; Holmes, E.C. Extensive Diversity of RNA Viruses in Australian Ticks. J. Virol. 2019, 93. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  69. Isberg, S.; Thomson, P.; Nicholas, F.; Barker, S.; Moran, C. Quantitative analysis of production traits in saltwater crocodiles (Crocodylus porosus): II. age at slaughter. J. Anim. Breed. Genet. 2005, 122, 370–377. [Google Scholar] [CrossRef] [PubMed]
  70. Hartley’s Creek Crocodile Adventures. Hartley’s Creek Crocodile Farming. Available online: https://www.crocodileadventures.com/sustainability/hartleys-creek-crocodile-farm (accessed on 1 January 2022).
  71. Sang, R.; Lutomiah, J.; Said, M.; Makio, A.; Koka, H.; Koskei, E.; Nyunja, A.; Owaka, S.; Matoke-Muhia, D.; Bukachi, S.; et al. Effects of Irrigation and Rainfall on the Population Dynamics of Rift Valley Fever and Other Arbovirus Mosquito Vectors in the Epidemic-Prone Tana River County, Kenya. J. Med. Entomol. 2017, 54, 460–470. [Google Scholar] [CrossRef] [Green Version]
Figure 1. Sero-prevalence of flavivirus antibodies in farmed crocodiles based on blocking ELISA using the flavivirus envelope protein-specific mAb, 6B6C-1. Positive samples are blue bars, and negative samples are black bars.
Figure 1. Sero-prevalence of flavivirus antibodies in farmed crocodiles based on blocking ELISA using the flavivirus envelope protein-specific mAb, 6B6C-1. Positive samples are blue bars, and negative samples are black bars.
Viruses 14 01106 g001
Figure 3. Proportion of samples positive in the 6B6C-1 blocking ELISA that have neutralising antibodies against tested flaviviruses. The overall frequency was determined based on n = 130. The frequency per farm was estimated based on n = 67 at Farm A, n = 41 at Farm B, and n = 22 at Farm C.
Figure 3. Proportion of samples positive in the 6B6C-1 blocking ELISA that have neutralising antibodies against tested flaviviruses. The overall frequency was determined based on n = 130. The frequency per farm was estimated based on n = 67 at Farm A, n = 41 at Farm B, and n = 22 at Farm C.
Viruses 14 01106 g003
Figure 4. Proportion (%) of samples positive in the 6B6C-1 blocking ELISA that have neutralising antibodies against two or more tested flaviviruses. The overall frequency was determined based on n = 130. The frequency per farm was estimated based on n = 67 at Farm A, n = 41 at Farm B, and n = 22 at Farm C (see Table S2 in Supplementary Materials).
Figure 4. Proportion (%) of samples positive in the 6B6C-1 blocking ELISA that have neutralising antibodies against two or more tested flaviviruses. The overall frequency was determined based on n = 130. The frequency per farm was estimated based on n = 67 at Farm A, n = 41 at Farm B, and n = 22 at Farm C (see Table S2 in Supplementary Materials).
Viruses 14 01106 g004
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Habarugira, G.; Moran, J.; Harrison, J.J.; Isberg, S.R.; Hobson-Peters, J.; Hall, R.A.; Bielefeldt-Ohmann, H. Evidence of Infection with Zoonotic Mosquito-Borne Flaviviruses in Saltwater Crocodiles (Crocodylus porosus) in Northern Australia. Viruses 2022, 14, 1106. https://0-doi-org.brum.beds.ac.uk/10.3390/v14051106

AMA Style

Habarugira G, Moran J, Harrison JJ, Isberg SR, Hobson-Peters J, Hall RA, Bielefeldt-Ohmann H. Evidence of Infection with Zoonotic Mosquito-Borne Flaviviruses in Saltwater Crocodiles (Crocodylus porosus) in Northern Australia. Viruses. 2022; 14(5):1106. https://0-doi-org.brum.beds.ac.uk/10.3390/v14051106

Chicago/Turabian Style

Habarugira, Gervais, Jasmin Moran, Jessica J. Harrison, Sally R. Isberg, Jody Hobson-Peters, Roy A. Hall, and Helle Bielefeldt-Ohmann. 2022. "Evidence of Infection with Zoonotic Mosquito-Borne Flaviviruses in Saltwater Crocodiles (Crocodylus porosus) in Northern Australia" Viruses 14, no. 5: 1106. https://0-doi-org.brum.beds.ac.uk/10.3390/v14051106

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop