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In order to verify the incidence of re-infection during this study, including RT-PCRs, CS, faeces and winter/summer results were compared (Table 3). For TAstV-2 search, 10% of CS and 20% of faeces were positive at winter, and 36% of CS and 20% of faeces showed an increase in the same period for TCoV, when individual RT-PCR was evaluated. In addition, the TAstV-2 was less detectable, CS 50% and faeces 72%, in the winter when multiplex RT-PCR was used (Table 3). Otherwise, the TCoV was equal detected from CS and 28% more detectable in winter for the same analysis. The results showed on Table 3, RT-PCR assayed for both virus in a single tube have 3.98 (p=0.89982) more chance to present positive results (faeces) than CS, at dry season, and faeces have 0.67 of chance to give false negative results for the same statistical analysis (p=0.67851).
South Eastern of Brazil is classified as being tropical wet and dry or savannah (Aw) type according to Köpper climate classification system, since the area presents an extend dry season during winter (end of May to September) temperature winter month > 20ºC and rainy season during summer (October to March approximately) precipitation in the driest summer month < 30 mm (19). The meteorological data (rainfall and relative humidity) for this region during the period of study presented monthly average temperature ranging from 39.3 and 31.2ºC, the precipitation in rainy season varied from 40 to 270.3 mm/month, and there was no rain at all during dry season (results not shown).
Statistical analysis was performed using GraphPad Prism 6.0. Correlation of C. psittaci bacterial load and adenovirus viral load was determined using linear regression analysis. For statistical calculation, 50% of the lower detection limit (6,000 copies/ml for C. psittaci and 600 copies/ml for adenovirus) was assigned to specimens that tested negative. Log-transformed viral loads were used for statistical analysis.
At present, the emergence of new pathogens and the continuous circulation of common etiological agents in dogs have made canine diseases more complex and difficult to diagnose. Dog infectious diseases mainly include respiratory and intestinal viral diseases, including CRV (CAV-2, CDV, CIV and CPIV) and CEV (CAV-2, CanineCV, CCoV and CPV). However, the traditional methods of virus identification and isolation are time consuming, causing delays in treatment initiation. A few methods for detecting virus-induced respiratory or enteric disease have been developed [4, 27, 28, 34], but no previous study had developed a systematic way to detect both CRV and CEV in dogs. Here, we developed two mPCR methods for detection of the most frequently coinfected viruses; these methods could be performed to diagnose dogs according to their clinical symptoms.
Primer design is the first and most important step in the process of establishing a detection method, and the following conditions must be satisfied: primers were designed to bind to conserved sequence regions, to have similar annealing temperatures, and to lack dimers or hairpin structures. In these novel mPCR methods, the primer combination produced amplicons that were easy to distinguish from each other, the primer annealing temperatures were similar, and degenerate bases were required only infrequently. The specificity, sensitivity and reproducibility tests all showed good results.
The mPCR methods were tested on 20 NS and 20 AS samples collected from dogs with symptoms of respiratory disease or enteric disease. The ratio of positive samples to total samples was 80% (16/20) for CRV detection and 85% (17/20) for CEV detection. Because the sample number was insufficient, these results were not statistically significant. However, CPV and CDV clearly remain two of the more serious and epidemic diseases in dogs in worldwide at present [35–38]. Epidemiological monitoring of CPV is particularly important because CPV evolves at a rapid rate, similar to that of Porcine Circovirus 3 [39, 40]. Because a small number of dogs were negative for the viruses tested by the CRV or CEV detection assays, although they suffered respiratory illness or intestinal problems, we suggest that some viruses with low prevalence and pathogenic bacteria may also cause disease in dogs [2, 41]. A variety of pathogenic bacteria are often present along with viruses in canine infections [42, 43], and thus, it is essential to expand the coverage of mPCR detection in the future. For example, CIRD also include CHV-1, canine reovirus, and Bordetella bronchiseptica and so on. At the same time, other pathogens causing serious zoonotic diseases, such as pseudorabies virus, should also be monitored in future [44, 45].
In this study, the detection of CanineCV was added to an mPCR method for the first time, because coinfection of this pathogen with other pathogens is common. Though the pathogenic mechanism of CanineCV is unclear, epidemiological testing is important for future research. CanineCV was not detected from the AS clinical samples; perhaps the limited source of these clinical samples was responsible for this result. We didn’t get a lot of clinical samples because it was not easy to get disease samples. CAV-2 mostly replicates in the lower respiratory tract and was detected in the NS samples; however, the CAV-2 primer pair used in this study was probably able to amplify the CAV-1 DNA virus despite the optimization performed. Notably, the live vaccine strains used may have an unavoidable impact on disease detection using the methods developed in this study. Additionally, discriminating between wild-type infections and vaccines is important, and therefore, a trend exists toward later development of broad-spectrum and accurate mPCR detection methods. Sometimes, cross contamination may lead to experimental failure. It is worth noting that PCR pretreatment and post-treatmen performed in different isolation zones can effectively avoid pollution. Besides, regular air spray cleaning will also play a role.
In conclusion, these newly established mPCR methods provide an efficient, sensitive, specific and low-cost testing tool for the detection of CRV (CAV-2, CDV, CIV and CPIV) and CEV (CAV-2, CanineCV, CCoV and CPV). The use of Taq Master Mix makes the detection process more convenient and reduces the chance of contamination during the process of sample addition; PCRs can be initiated by simply adding enzyme, ddH2O, premixed primers, and template, and thus, this method is superior to other mPCR detection methods. Here, detection of CanineCV was added to mPCR for the first time, making this method suitable for the further study of coinfection by CanineCV and other pathogens. This study provides a novel tool for systematic clinical diagnosis and laboratory epidemiological surveillance of CRV and CEV among dogs.
The patient was a 55-year-old male artisan working at the NTNAMC. He presented after four days of dyspnea and two days of hemoptysis. Chest radiograph showed bilateral patchy consolidation. Blood test showed neutrophilia and elevated liver enzymes with alanine transaminase of 51 U/L. He was put on non-invasive positive pressure ventilation for respiratory failure. Bronchoalveolar lavage tested positive for C. psittaci and rhinovirus by PCR and reverse transcriptase-PCR respectively. Direct fluorescent antigen detection and viral culture did not reveal other viral co-pathogens. Paired serology, collected 18 days apart (on days 2 and 20 after hospitalization), showed a rise of C. psittaci IgG titer from <32 to 128 by microimmunofluorescence assay, but there was no increase in adenovirus or other respiratory virus antibody titer. The patient recovered with oral doxycycline. He had contact with birds, monkeys, iguanas and snakes at the NTNAMC within one month of symptom onset.
To evaluate the reproducibility of the assay, the detection mPCRs for both CRV and CEV were performed as three independent mPCR assays by using three different PCR instruments at different times. Three premixed plasmids for CRV (Fig 5A) and CEV (Fig 5B), with different dilutions, could be amplified under different conditions and showed similar results among the assays.
A throat swab was taken from the patient at the time of his first clinical examination and transported in viral transport medium (VTM) to the National Public Health Laboratory for virus isolation. The sample was treated with antibiotics (C. penicillin 100,000 I.U./ml and streptomycin 100 µg/ml) for an hour before being inoculated in duplicate (100 µl and 200 µl, respectively) into freshly confluent monolayers of MDCK (ATCC, CCL-34), Vero (ATCC, CCL-81) and Hep-2 (ATCC, CCL-23) cells cultured in a 24-well tissue culture plate. The plate was incubated at 37°C in 5% CO2 and examined daily for the presence of CPE in cultured cells. Supernatant from cultures with visible syncytial cytopathic effect (CPE) after 3 days was taken for further analysis by serial passage in different cell lines available in the laboratory.
The investigation conducted in this study was approved by the ethics committee of the Malaysian National Public Health Laboratory. All patients (subjects) in this manuscript have given written informed consent (as outlined in the PLoS consent form) to publication of their case details. No identification of the subjects is to be revealed in any publication.
Sequence alignment was performed by ClustalW Multiple alignment using BioEdit software (version 188.8.131.52). Phylogenetic analysis was done using MEGA software (version 6.06), and phylogenetic trees were constructed by the neighbor-joining method based on MEGA software. The evolutionary distances were calculated using the Kimura 2-parameter method. Bootstrap value was computed on 1000 replicates and the significance of branch length was estimated by maximum likelihood.
The nucleotide sequences of the MRV obtained from environmental sewage described here have been deposited in the GenBank database under the following accession numbers: KR296769 to KR296785.
Culture supernatants of Cangyuan virus was analyzed by VIDISCR. Virus particles were harvested from cells by three freeze-thaw cycles and the resulting suspension purified from cell debris by low-speed centrifugation. Nucleic acids were extracted using the AxyPrep Body Fluid Viral DNA/RNA Miniprep Kit (Axygen, Inc.). Reverse transcription of the viral RNA was performed by using the RevertAid™ First Strand cDNASynthesis Kit (Fermentas, Inc). The VIDISCR assay was performed as previously described. To further characterize the virus and its phylogeny primers were designed with Primer Premier 5.0 based on published sequences selective for the 10 genome segments of Melaka virus and other orthoreoviruses (Additional file 4: Table S4). Each of the 10 genome segments were amplified using the pfu PCR Polymerase Kit (Fermentas, Inc) with the primers as listed in Additional file 4: Table S4. Reverse transcription PCR was performed as described. The amplicons were visualized using 2% agarose gel electrophoresis. PCR products were sequenced after cloning by using CloneJET™ PCR cloning kit (Fermentas, Inc). Sequence alignment was conducted using DNAMAN5.0 and phylogenetic analysis of the whole Cangyuan virus genome sequences of all L, M and S segments were performed by the neighbor-joining method using MEGA6 software (www.megasoftware.net). The Phylogenetic data have been deposited in TreeBase (Study Accession URL: http://purl.org/phylo/treebase/phylows/study/TB2:S16635). RT-QPCRs testing were repeated on the 50 fruit bats original samples including the Kidney, heart, lung, liver, spleen, intestine, rectal swab sample, and brain samples with the L2 segment primers (CY-L2QF1: 5′ GCA ATG CCG AAT ATC TAA AGC 3′, CY-L2QR1:5′ AGA GCA AGA GCC CAA ATG AA 3′). The reaction was performed using the One Step SYBR® PrimeScript™ PLUS RT-PCR Kit (TAKARA BIOTECHNOLOGY (Dalian) CO., LTD) by lightcycler2.0 (Roche). To test the viral growth curve of the Cangyuan virus, the virus samples were harvested at 2, 4, 8, 12, 16, 20, 24 and 28 hours after infection and quantified with the already established RT-QPCR. To test whether the Cangyuan virus is the virus (or one of the viruses) replicating in the cells and responsible for the observed CPE, the culture supernatants 0.1 mL (after the second passage,105.5 TCID50/0.1 mL titer) was serially diluted until 10−3 and infected Vero E6 cells. After the 24 hours, the culture supernatants were analyzed by RT-QPCRs with the L2 segment primers.
All procedures using animals were approved by the Animal Care and Use Committees of Centre for Disease Control and Prevention, Chengdu Military Region and were in compliance with the China Animal Welfare Act. We state clearly that no specific permissions were required for these locations/activities and confirm that the field studies did not involve endangered or protected species.
Cell viability, based on entry of fluorescent dye into cells with compromised cell membranes, was quantified using a LIVE/DEAD Fixable far red fluorescent kit (L10120; Life Technologies) in a BD FACS CANTO II flow cytometer (BD Biosciences). Data analysis was carried out using the Kaluza Analysis software (Beckman Coulter). A heat-killed control, subjected to 60 °C for 20 min, and uninfected control were used to determine the fluorescence threshold between viable and dead cells.
Radioimmunoprecipitation assay (RIPA) buffer (Santa Cruz) supplemented with 1% phenylmethylsulfonyl fluoride (PMSF) (Santa Cruz), 1% cocktail inhibitor and 1% sodium orthovanadate (Santa Cruz) was used to lyse cells. Bio-RAD protein assay was used to determine protein concentration (Bio-Rad). Primary antibodies used were mouse anti-viral NP (at 1:3000 dilution; PA5–32242, Pierce), goat anti-viral PB1 (at 1:10000 dilution; 17,601, Santa Cruz), goat anti-viral M1 (at 1:2000 dilution; ab20910, Abcam), mouse anti-β-actin (at 1:10000 dilution; A5316, Sigma) and secondary antibodies used were donkey anti-goat IgG (at 1:10000 dilution; sc-2020, Santa Cruz) and goat anti-mouse IgG (at 1:1000 dilution; HAF007, R&D Systems).
Immunofluorescence antibody testing (IFAT) and virus neutralization assay were conducted as previously described. Briefly, for IFAT a freshly confluent monolayer of MDCK was infected with KamV and at full CPE, the infected cells were harvested, washed four times and suspended in sterile PBS at a cell concentration of approximately 3000 cells per millilitre. An aliquot of the infected cell suspension was carefully spotted onto each well of Teflon coated slides, followed by air-drying over a warm plate and subsequent fixation in cold acetone for 10 min. Serial 2-fold dilutions of serum samples were then added to detect specific reactivity. For detection of IgM, IgG was removed by absorption with protein A prior to serum dilution. Bound antibodies were detected using fluorescein conjugated rabbit anti-human IgM or IgG (Dako, USA). Specific reactivity/labelling were read under a UV fluorescence microscope (Olympus BX50, Japan). For VNT, serial 2-fold dilutions of control and test sera were prepared in duplicate starting at 1∶10. An equal volume of virus working stock containing 150 TCID50 was added to the diluted sera and incubated for 30 min. The pre-incubated virus/serum mix was added to confluent cell monolayers and incubated for 1 h. The inoculum was removed, monolayers washed three times with PBS and cell media replaced. Ability of sera to neutralize virus was determined by scoring the extent of CPE observed in duplicate wells three days later.
Evaluation of immunosuppression is based on field criteria and laboratory investigations (Fig. 1). Yet, practical and evaluable methods are restricted for an accurate evaluation.
Field criteria consist on global evaluation of the flock health statute. However, these criteria are nonspecific and allow for only first orientation. In general, immunosuppression leads to degradation of performances, with poor feed conversion, decrease in growth rate, heterogeneity, low weight, and increased mortality. Due to the influence of viruses on host immune system, vaccination failures are declared. Depressed animals are more susceptible to develop bacterial and parasitic secondary infections, which are accompanied by increased mortality (Fig. 2). The various symptoms and lesions induced by immunosuppressive viral diseases can help to establish clinical suspicion, which need laboratory investigations to confirm it.
Many laboratory tests are useful in order to evaluate immunosuppression in turkeys. The global approach can be summarized in four main criteria (Dohm and Saif, 1984):
Morphometric changes in central and peripheral lymphoid organsChanges in concentration or ratio of immunoglobulin classes within serum and secretions, and changes in serum complement levelChanges in functional activity of the immune responseDemonstration that the suspected immunosuppressive agent will interfere with vaccination and/or exacerbate the course of a disease induced by another agent.
Pathomorphological examination of lymphoid organs is easily practicable. The use of quantitative indices may largely contribute to more rapid and correct diagnosis. Lymphoid organs masses (bursa, thymus, and spleen) contribute to objective evaluation (Halouzka and Jurajda, 1991; Sellaoui et al., 2012). Histopathological investigation is an important tool for evaluating severity of immunosuppression and discriminating between several diseases (Pope, 1991).
Detection of specific causal agents by viral isolation and molecular detection is a practical approach. Serology may be an easy test for the diagnosis. Evaluation of cellular immune competency is used in vitro as well as in vivo (Fadly et al., 1982). Hematological investigations may provide heterophil:lymphocyte ration, as significant indicator of stress (Huff et al., 2005; Cotter, 2015) and immunocompetency (Hocking et al., 2002).
In vitro lymphocyte proliferation response to mitogen is widely used to evaluate the integrity of cell-mediated immunity. Lymphocyte proliferation responses of spleen cells are higher in turkeys infected by TCoV than in non-infected animals, with increase of CD4+ subpopulation of T lymphocytes (Loa et al., 2001). The lymphoproliferative response to phytohemagglutinin phosphate is performed as an indicator of a T-cell-induced delayed-type hypersensitivity reaction. A mononuclear phagocytic system function assessment is used to study the degree of clearance from the blood circulation in commercial turkeys (Cheema et al., 2007).
Determining cytokines levels to evaluate cellular immune response is well developed in mammals due to available commercial systems, especially ELISA tests and RT-PCR (Wigley and Kaiser, 2003). For avian species, IFN-gamma can be quantified by currently available ELISA test in chickens (Lambrecht et al., 2000) and in turkeys (Lawson et al., 2001).
Microarray technology is performed to evaluate cellular immune response in poultry, by detecting genes involved in antiviral and pro-inflammatory cytokine responses (Kapczynski et al., 2013). Moreover, innate and adaptive immune responses can be explored by real-time RT-PCR to investigate changes in the gene expression of cytokines interleukin (IL) and chemokines (Gadde et al., 2011).
Evaluation of immunosuppression represents a delicate approach, which is not routinely applied in poultry pathology. Exploration of the different components of the immune system is based on several in vivo and in vitro methods. Due to the complexity of the immunosuppression etiology, the consequence of several intrinsic and extrinsic factor interactions, the use of combination of many techniques would help interpreting the data.
The infected cells were fixed with 4% paraformaldehyde (Sangon Biotech) for 10 min, followed by rinsing with PBS. 0.2% v/v triton X-100 (Sigma) diluted in PBS was applied to each well for 10 min. After washing three times with sterile PBS, the cells were overlaid with PBS containing 5% w/v bovine serum albumin (Sangon Biotech) and incubated for 30 min at 37° C. The mouse antiserum anti-capsid P2 was used as the primary antibody with a 1:200 dilution in PBS. After 45min incubation at 37 °C, the cell monolayers were rehydrated by rinsing three times with PBS. Coverslips were stained using goat anti-mouse IgG conjugated with FITC (KPL) and incubated for a further 45 min at 37° C. The stained cells were rinsed with PBS, then dyed with Hoechst33342 (Solarbio) and rinsed again. The stained cells were then examined under a confocal microscope.
Blood was drawn from the metatarsal or brachial vein and aliquoted into additive-free, EDTA, and lithium heparin blood collection tubes (Becton Dickinson, Franklin Lakes, NJ). Samples were kept cool on ice for transport back to the University of California, Davis where they were aliquoted and stored at -80°C until shipment to the diagnostic laboratories. Serological testing for avian adenovirus, Chlamydophila psittaci, infectious bronchitis virus (IBV) (Arkansas (Ark), Connecticut (Conn) and Massachusetts (Mass) strains), Mycoplasma gallisepticum, Mycoplasma synoviae, avian paramyxovirus-1 (Newcastle Disease virus), avian paramyxovirus-2, avian paramyxovirus-3, and avian reovirus was performed at Texas Veterinary Medical Diagnostic Laboratory (College Station, TX) using assays optimized for poultry species. Because these assays have not been validated in condors, vultures, or eagles, we relied on cut-off titers established for poultry. Exposure to avian adenovirus was evaluated using an agar gel immunodiffusion (AGID) test. Chlamydophila psittaci exposure status was determined by a direct complement fixation (DCF) assay. Exposure status for the three strains of IBV was determined by hemagglutination inhibition (HI) assay. A titer of 1:16 or greater was considered positive for exposure to IBV. Mycoplasma gallisepticum exposure status was determined using serial tests. Samples were initially screened using a plate agglutination assay. Any samples that were positive on the first assay were then tested by HI. A titer of 1:80 or above on the HI assay was considered positive. A sample testing positive on both the plate agglutination test and HI assay was considered positive for exposure to M. gallisepticum. Mycoplasma synoviae exposure status was determined in the same manner as M. gallisepticum. Avian paramyxovirus 1, 2 and 3 exposure status was determined by HI. A titer of 1:16 or greater on the HI assay was considered positive for each of the three paramyxoviruses. Avian reovirus exposure status was determined using AGID. The number of individuals tested varied between pathogens due to limitations in sample volume.
Serological testing for Toxoplasma gondii was performed at University of California, Davis (Department of Pathology, Microbiology, and Immunology, University of California-Davis) using a T. gondii agglutination test kit (Eiken Chemical Co., LTD. Tokyo, Japan, distributed by Tanabe USA, Inc., San Diego, CA). Manufacturer’s recommendations were followed, and a titer of 1:32 or greater was considered positive. Testing for arboviruses common to California was performed at the Center for Vectorborne Diseases (University of California-Davis). West Nile virus titers were determined using an indirect enzyme immunoassay (EIA) as previously described [75–78]. West Nile Virus cross reacts with St. Louis encephalitis virus (SLEV) and Western equine encephalitis virus on EIA, so EIA positive samples were tested using end-point plaque neutralization (PRNT) assays using the NY99 strain of WNV and the Kern 217 strain of SLEV. The PRNT assays were performed using >75 plaque forming units of virus grown on Vero cell culture. To be considered positive, sera had to neutralize >90% of the virus in at least a 1:4 dilution. Sera from free-flying California condors were also evaluated for active WNV infection by real time-polymerase chain reaction (rt-PCR) using primers as previously described [79, 80]. Additionally, free-flying condors that had a high WNV titer (≥ 1:256) were also screened for active WNV infection by PCR on whole blood (n = 9).
Liver and lung tissue samples collected from 14 condors that died of various causes between 1997 and 2009 were also selected for detection of a subset of potential pathogens. These 14 condors had all been in captivity and in the wild at different points in their lives. Representative cases were selected for analysis based on availability of frozen tissues at San Diego Zoo Safari Park and degree of autolysis. Tissue samples were screened for presence of avian adenovirus, coronavirus (including infectious bronchitis virus), paramyxovirus and Mycoplasma spp. DNA from tissues was extracted using the DNeasy Blood and Tissue kit (Qiagen, Valencia, CA, USA) following the manufacturer’s protocol. RNA was also extracted from the lung tissue samples using the QIAamp cador Pathogen Mini kit (Qiagen, Valencia, CA, USA) following the manufacturer’s guidelines. PCR primers were synthesized by Integrated DNA Technologies (San Diego, CA, USA) and were utilized in nine assays using the following primer pairs: adenovirus primers AdenokissF/AdenokissR, coronavirus primers Corona8pF/Corona7mR, IN-2F/IN-4R, Cor-p-F2/ Cor-p-R1 and Cor-p-F3/ Cor-p-R1, paramyxovirus primers NCD-3/NCD-4, ParamyxoP1/ParamyxoPR and ParamyxoP2 /ParamyxoPR and Mycoplasma spp. primers MycogenusF/MycogenusR and MycaldP/CapaldM. Positive controls were available for adenovirus and Mycoplasma spp.
For bacteriological diagnosis, liver, and kidney samples from dead goslings were first inoculated onto tryptic soy agar plates (BD Science, MD, USA) containing 2% fetal calf serum, and incubated at 37 °C under an atmosphere with 5% CO2 for 48 h. Then the spleen, liver, and kidney tissue were pooled and tested for the presence of goose parvovirus28, goose hemorrhagic polyomavirus29, AstV21, reovirus30, and Tembusu virus31, respectively.
Clinical manifestations of AvRV infection include mild to severe diarrhea, a varied degree of dehydration, and stunted growth; it also may remain asymptomatic. These variations may be due to the differences in the severity of the particular strain or the interaction between different environmental and management factors. These manifestations alone are not sufficient for confirmatory diagnosis. Therefore, the identification of virus in fecal content or antibody detection in serum should be used to confirm the RV infection. To date, there is no assay for RVD antibody detection. The major methods used for diagnosis of RVs in poultry include:
Electron microscopy: The distinct wheel-like morphology of RV was initially used for detecting RV infection by direct visualization of the virus in feces or intestinal content. Using this technique, RVs of different groups cannot be distinguished. Immune electron microscopy can be used to distinguish serogroups, although it requires the availability of specific antisera. It is a sensitive diagnostic approach, but is also costly and cumbersome.
Virus isolation: AvRV can be isolated in embryonated chicken eggs (via yolk sac route), primary in cell culture (chicken embryo liver cells/chicken embryo kidney cells) or in continuous cell lines (MA104/Rhesus monkey kidney cell line). The isolation is useful only for AvRVAs, but it is not commonly used for diagnosis. It is very difficult to propagate other RV serogroups in cell cultures, and it has been reported that RVD cannot be propagated in MA104 cell culture systems.
RNA Polyacrylamide Gel Electrophoresis (RNA-PAGE): The detection of RV-RNA in feces or intestinal content provides an alternate means of diagnosis. Following RNA extraction, electrophoresis on polyacrylamide gels, and silver staining, RNA can be identified by the pattern of migration of genome segments. RNA-PAGE is a highly specific technique used for the detection of segmented viruses. It detects the electrophoretic migration pattern of all 11 segments of RV, which is different among different groups. According to the distribution of segments in each region, the AvRV-A has a pattern of 5:1:3:2, and the RV-D has a pattern of 5:2:2:2, while the mammalian RV-A shows a pattern of 4:2:3:2. AvRVs RVF and RVG, which shows sporadic shedding, have migration patterns of 4:1:2:2:2 and 4:2:2:2:1, respectively. However, these patterns can’t be totally relied upon, because substantial differences are observed in the electrophoretic pattern of RVs when conditions of gel electrophoresis are varied.
Reverse Transcription-Polymerase Chain Reaction (RT-PCR): This is the most sensitive molecular detection tool available for the diagnosis of RVs, and is mainly based on the VP6 gene segment. For the detection of AvRVs, only a few RT-PCR protocols are available, and most of them solely detect AvRV-A. In 2011, an RT-PCR was developed by Bezerra and co-workers specifically for the detection of the VP6 gene of RVD. Real-time PCR and RT-PCR are now available for the detection of RVD. The sensitivity of real-time RT-PCR was found to be similar to that of conventional RT-PCR when the same primer sets were used for both of the assays.
Serological methods: Serological methods used for the detection of RVs include counter immunoelectrophoresis, radioimmunoassay (RIA), the latex agglutination test (LAT), and enzyme-linked immunosorbent assay (ELISA). AvRVs can be detected using ELISA. Commercialized ELISA kits are available for detection of RVAs, such as the IDEIA RV assay (DAKO, Ely, UK), the RIDASCREEN® assay (r-biopharm, Darmstadt, Germany), etc. However, for the detection of other AvRVs such as RVD, RVF, or RVG, no ELISA is available.
The virus neutralization test was performed as reported31. Briefly, the convalescent sera both from the clinical survived goslings (n = 6) with gout disease and the experimental infected survived goslings (n = 9) were serially diluted with OPTI-MEM medium. Then, 50 µL of the diluted sera were mixed with an equal volume of 200 TCID50 virus. The mixture was incubated for 1 h at 37° C, then inoculated into LMH cells and the inoculated cells were washed once with PBS. At day 4 post inoculation, the inoculated cells was analyzed by indirect immunofluorescence assay to determine the neutralization titer of the sera.
Enteric infections are one of the principal causes of diarrhea, which results in poor feed conversion efficiency and reduced growth, and hence leads to heavy economic losses for the poultry industry. These infections are caused by different agents, including viruses (rotavirus, coronavirus, astrovirus, reovirus, adenovirus, parvovirus etc.), bacteria (Salmonella, Enterococcus, E. coli) and protozoans (Cryptosporidium and Eimeria). The clinical manifestations of these pathogens are almost similar. Hence, RVD must be differentiated from all of these enteric pathogens, as well as from the other groups of rotaviruses found in poultry.
The present work describes the isolation of the astrovirus AAstV/Goose/CHN/2017/SD01 from tissue samples of goslings dying from a disease characterized by visceral urate deposition. The successful reproduction of the disease by experimental infection demonstrates the etiological role of this AAstV. Based on the genetic analysis of the complete capsid region at amino acid level, the isolate should be assigned as a member within the genotype consisting of TAstV-2 and DAstV-1 strains. The high variability of the genomic sequence to other known astroviruses suggest more detailed antigenic investigations should be performed.
IFA was performed using MDCC-MSB1 cells (preserved by our lab) to detect CAV antigens, according to the method described by Yuasa et al.. Briefly, CAV-infected cells (PCR positive result) were smeared onto 35-mm culture dishes with 20-mm cover glass inserts (NEST, 801001, China) 3 days post infection, then dried, fixed with 4% paraformaldehyde for 30 min at room temperature (RT), and permeabilised with 0.1% Triton X-100 for 15 min at RT. After three washes with 0.05% Tween 20 PBS (PBST), cells were incubated with mouse monoclonal anti-VP1 primary antibodies (preserved by our lab that produced in mouse by prokaryotic expressed VP1) for 1.5 h at 37°C. After three washes with PBST, cells were incubated for 1 h with a 1:200 dilution of fluorescein isothiocyanate (FITC)-conjugated goat anti-mouse secondary antibody in the dark. After three washes with PBST, the infected MDCC-MSB1 cells were observed by fluorescence microscope (EVOS F1; Life, USA).
The session on hepatitis virus infection and liver cancer provided a research update on: the molecular virology and immunology of HBV; host factors related to hepatitis virus infection; the genetic landscape of virus-associated hepatocellular carcinoma (HCC); integrated genomics to identify drivers of human liver cancers; chemical-viral interaction between aflatoxin and HBV in induction of HCC; and antibody therapeutics targeting glypican-3 (GPC3) for the treatment of liver cancer. Through an extensive clinical network of patients in China, randomized clinical trials are being conducted to study new treatment strategies for chronic HBV infection involving nucleotide analog reverse transcriptase inhibitors and peginterferon. An ongoing effort to prevent mother–child transmission of HBV through a short-term antiviral therapy with tenofovir disoproxil fumarate during late pregnancy, reported a significantly lower HBV transmission rate compared to the control group. This session also included a discussion of host factors related to hepatitis virus infection using a genome-wide association study (GWAS). A genetic analysis using GWAS, identified host factors for various human multifactorial diseases, as well as interferon (IFN) lambda for drug response, and HLA II genes for susceptibility to chronic HBV infection (CHB). Frequencies of HLA-DP risk alleles are high in Asian populations, whereas frequencies of HLA-DP protective alleles are high in European populations. These findings could explain the high incidence of CHB in Asian countries and suggest that host genetic factors are important to viral infections. Another talk outlined the genomic and epigenomic associations in HBV-related liver cancer using data obtained from whole genome bisulfate sequencing and whole genome sequencing. Clonal HBV integrations preferentially occurred in inactive chromatin regions; massive rearrangements were detected in the integrated HBV genome, and a negative correlation exists between HBV rearrangement number and total somatic mutation number. These observations could be useful for understanding the progression of HBV-related liver cancer. It was noted that liver cancer, including HCC and cholangiocarcinoma, is the second leading cause of cancer death (about 9.1% of total cancer deaths) with a significant burden in low- and middle-income countries in Asia and sub-Saharan Africa. Etiological factors associated with HCC include infection with HBV and exposure to high levels of aflatoxin B1 (AFB1) in the diet. HCV-associated HCC is becoming the most rapidly rising solid tumor in the United States and Japan. The development of highly effective drugs that cure HCV infection is a major advance that, hopefully, will diminish the role of HCV in liver cancer. Ongoing clinical investigations are defining the utility of GPC3, a cell surface proteoglycan differentially expressed in HCC, and other promising antibody therapeutics to treat liver cancer.
Twenty nine commercial 1-day-old chickens were used to evaluate the protection provided by H120 vaccination against challenge with Egypt/F/03. Birds were divided into three groups; A (n = 12), B (n = 5), C (n = 12). Vaccination was performed at day 1 by eye drop application. Single dose of H120 vaccine (Nobilis, Intervet, The Netherlands BV) was used for each bird in groups A and C according to manufacturer's instructions while birds in group B were kept as unvaccinated control. Four weeks post vaccination, chickens in group A and B were challenged by eye drop with Egypt/F/03 (105 EID50 per bird) while birds in group C were not challenged and kept as vaccinated unchallenged control. Tracheae of all birds from all groups were collected four days post challenge for virus reisolation attempts and histopathological examination. Tracheal scrapings were emulsified in 2 ml of sterile PBS and centrifuged at 500 × g for 3 min. Virus reisolation attempts were performed by inoculating 2–3, 10-day-old SPF ECE by the supernatant fluid of each sample as described. Embryos were examined for typical lesions of IBV. For histopathological examination, tracheae were fixed in formalin, processed routinely for histopathology and stained with haematoxylin and eosin. The trachea from each bird was examined microscopically and assigned lesion scores of 0–3 with 0 = none, 1 = focal, 2 = multifocal, 3 = diffuse. Tracheae were scored for the amount of mucous, loss of cilia, epithelial hyperplasia, necrosis, lymphocyte and heterophil infiltrations as well as the extent of tissue reaction. The scores for each bird were added and the mean score for the birds in each group was calculated. Kidney samples were also taken 4 days post challenge and examined microscopically for tubular degeneration and inflammation consistent with interstitial nephritis. Focal, multifocal and diffuse were used to assign kidney histopathology. The presence of viral antigens in kidneys was screened by immunofluorescent antibody technique.