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Our literature review suggests that community-acquired adenoviral pneumonia in immunocompetent adult civilians presents as a non-specific febrile respiratory illness that progresses rapidly to respiratory failure and often requires mechanical ventilation. The laboratory and radiological findings are typical of viral infection but are also non-specific. Novel respiratory virus real-time RT-PCR testing enabled us to rapidly detect adenovirus as the cause of severe community-acquired pneumonia in our patient.
The HAdV detection rates for the different populations and different seasons were compared by a χ2 test, and the relationship between vomiting and HAdV types was statistically analyzed by Fisher’s exact probability, with statistical significance set at P < 0.05. Vector NTI 11.0 software was used for sequence alignment and Mega 5.0 software for phylogenetic analysis. Epidemiological data were analyzed using statistical product and service solutions (SPSS) 21.0 software.
All the hexon gene sequences obtained by nested PCR together with the 26 HAdV strain sequences available in GenBank were used for the phylogenetic analyses. The phylogenetic tree was constructed using molecular evolutionary genetics analysis (Mega) 5.0 and evaluated using 100 bootstrap replicates to verify the HAdV genotypes. Positive PCR products were named according to the corresponding serial numbers of the specimens.
Regarding HAdV infection comparisons between age groups were performed using the Fisher’s exact test. P value < 0.05 was considered statistically significant and the 0–5 year age group was used as reference group. HAdV mono-infections were also compared to HAdV co-infections. The R.3.0.1 tool was used to perform the analyses.
In consideration with low Ct-values, 80 HAdV positives samples (20 per year) were selected using a random number generator on MS Excel for further molecular characterization using classical PCR and sequencing. Viral DNA was extracted as previously described and eluted with 50 μl water nuclease-free. DNAs were stored at −20°C until PCR reactions. For HAdV molecular characterization the last 300 base pairs (bp) of the hexon gene were amplified with the following specific primers: Adeno3 (5’-CCTTTGGCGCATCCCATTCT-3’) and Adeno4 (5’-TGGGCACCTATGACAAGCGC-3’) previously used by Garcia et al,. The Phusion High-Fidelity PCR Master Mix with HF Buffer (New England Biolabs, Ipswich MA, USA) was used for amplifications. For each sample, PCR was carried out in a total reaction volume of 50 μl consisting of 15 μl H2O RNase free, 2.5 μl of each primer (diluted at 10μM), 25 μl of 2X Phusion Master Mix and 5 μl of DNA template. Cycling conditions were as follows: denaturation step of 15 min at 95°C, 40 PCR cycles including 30 s at 95°C, 60 s at 55°C, 60 s at 72°C followed by an extension step of 10 min of 72°C.
Five microliters of the PCR product was then mixed with 1 μl of 10X 5PRIME loading dye and loaded on to a 1% agarose gel along with an appropriated molecular weight markers (100 bp ladder, New England Biolabs), and gels were stained with ethidium bromide (0.5 μg/ml) before visualization under UV.
For positive samples (380 bp size band), amplicons were cut and purified using the GeneJET Gel Extraction Kit (Thermo Scientific). Purified products are then sent for sequencing to Beckman Coulter Services. Sequencing was performed in both directions with the same PCR primers (Adeno3 and Adeno4) on an ABI PRISM BigDye Terminator v3.1 Ready Reaction Cycle Sequencing kit (Applied Biosystems) on a 96-capillary ABI PRISM 3730-XL (Applied Biosystems). Data in FASTA format were then sent to the laboratory for analysis.
ALT: alanine aminotransferase; ANCA: anti-neutrophil cytoplasmic antibodies; BAL: bronchoscopic alveolar lavage; CPK: creatinine phosphokinase; Ct: cycle threshold; RSV: respiratory syncytial virus; RT-PCR: reverse transcriptase polymerase chain reaction.
The representation of patient’s data and detected pathogens were analyzed using the SPSS 20.0 statistical package. The relation and differences between pathogens infection rates, clinical manifestations and the statistical calculations were performed using the Chi-square (X2) test or the Fisher’s exact test (where cell counts below 5 were encountered in the statistical table) on SPSS. Additionally, to compare total positive, single, and multiple viral infections with patient’s demographic information and the manifestation of ARTIs, the binary logistic regression test was performed when more than 2 categories in the variable were counted. It evaluates the association between the exposure of 2 properties and their outcome (odds ratio: OR) and estimates the associated 95% confidence interval (95% CI) which indicates the degree of uncertainty associated with the OR. A value of p< = 0.05 was considered as significant.
Virus isolation for the adenovirus-positive specimens was performed by using HEp-2 cell lines (from American Type Culture Collection, ATCC Number CCL-23) following the standard protocol. Cells inoculated with clinical samples were incubated at 37°C for 7 days. If no cytopathic effect was observed, the culture was used to inoculate fresh cells for up to 2 additional passages; the cultures with adenovirus-like cytopathic effects were passaged again to confirm the presence of the virus.
Samples enrolled for this study were collected in the context of routine laboratory testing of RSV infection using rapid diagnostic assays (data and results not shown). The study was approved with a formal authorization by the Scientific and Ethical Committee of Farhat Hached University-hospital of Sousse, Tunisia (approval no. IRB 00008931 provided by OHRP).
Seropositivity ratios were evaluated by χ
2 test (Minitab 12.0). P < 0.05 value was taken to indicate statistical significance.
Among the 800 selected samples, viral detection was negative in 400, 338 were positive for 1 viral target, 58 were positive for 2 targets and 4 were positive for 3 targets. The average cellularity was 4.76 (+/- 1.41) Log/ml, 4.95 (+/- 1.26) Log/ml, 5.30 (+/- 1.17) Log/ml, and 6.19 (+/- 0.21) Log/ml for these 4 groups respectively. The average cellularity in Negative samples was significantly lower than in cases of mono (p = 0.049), bi (p = 0.004) or tri-detection (p = 0.032). A significant tendency was observed between positive samples for one viral target and those positive for 2 or 3 virus (p = 0.064), this trend was confirmed by a Spearman correlation (ρ = 1) indicating a strong correlation between sample cellularity and the number of viruses detected.
The average cellularity was determined for each viral species detected in the positive samples for a single virus (n = 338/400). The 62 viral co-detection samples were excluded. The results were as follow: RSV = 4.56 (+/- 1.27) Log/ml (n = 40); HCoV = 4.73 (+/- 1.45) Log/ml (n = 49); PIV 1-4 = 4.77 (+/- 1.37) Log/ml (n = 19); Flu A-B = 4.89 (+/- 1.29) Log/ml (n = 79); AdV = 5.04 (+/- 0.94) Log/ml (n = 25); RhV/EV = 5.15 (+/- 1.20) Log/ml (n = 106); hMPV = 5.47 (+/- 0.85) Log/ml (n = 16) (Fig. 3). There is a significant difference of cellularity between RSV and RhV/EV positive samples (p = 0.012), between RSV and hMPV positive samples (p = 0.015), and between HCoV and hMPV positive samples (p = 0.041).
Blood serum samples belonging to 188 dogs, which had either been admitted to the Internal Medicine Clinic of Selcuk University, Faculty of Veterinary Medicine, with clinical symptoms or had been sampled at the dog shelters they were cared after in Isparta and Burdur provinces, were examined using the ELISA method. Of these samples, 103 (54.7%) were found to be positive for antibodies against CAV infection (Table 3).
Of the 108 female animals sampled in the study, 55 (50.9%) were determined to be positive for CAV antibodies, while 48 (60%) of the sampled 80 male animals were confirmed to be positive (Table 3). Of the 7 animals below 1 year of age, only 1 (14.2%; 2-month-old female puppy) was positive, and the remaining ones were found to be negative for CAV antibodies. Of the 53 animals aged 1-2 years, 22 (41.5%); of the 58 animals aged 2 years, 31 (53.4%); of the 64 animals aged 3 years, 44 (68.7%); and of the 6 animals aged >4 years, 5 (83.33%) were found to be positive (Table 2).
Blood leukocyte samples from dogs were processed and inoculated onto confluent monolayers of MDCK cells using standard virological techniques. The inoculated cells were incubated at 37°C and observed daily for the appearance of cytopathic effect (CPE). After third passage, cells were examined by immunofluorescence test for virus isolation. No morphological changes were observed in cell cultures, and a positive result was not detected by immunofluorescence test.
Clinical Findings. Blood samples were taken from 111 dogs showing clinical symptoms which were brought to the Internal Medicine Clinic of Selcuk University, Faculty of Veterinary Medicine. Seventy-seven dogs were sampled from Isparta and Burdur dog shelters by random sampling, regardless of the clinical findings. Dogs showed a systemic disease, characterized by fever, diarrhea, vomiting, mucopurulent oculonasal discharge, mucopurulent conjunctivitis, severe moist cough, signs of pulmonary disease, and dehydration. Corneal opacity and photophobia were determined for two dogs.
Given the observation that picornavirus sequences generated high number of reads in all 3 pools examined by deep sequencing, we asked whether type-specific enteroviruses could be identified among the individual samples comprising each pool. Towards this, we tested each sample for the virus identified from the most frequent virus reads found in each pool using RT-PCR. For HFMD-lib01, we found one sample positive for CV-A16 after conventional sequencing and BLAST analysis, while the rest of the samples in this pool tested negative. In pool HFMD-lib02, which showed high number of reads for rhinovirus-C, we also found that only one sample tested positive for rhinovirus. Finally, we identified one sample positive for CV-A21 in HFMD-lib03 pool. Taken together, these results verified that the presence of enterovirus originated from a unique sample and contributed to the high number of reads observed in each pool.
A direct immunofluorescence assay for broad detection of adenoviruses was used to screen for TMAdV in necropsy tissues from experimentally infected marmosets in the current study and naturally infected titi monkeys from the previously reported pneumonia outbreak. Areas of bright apple-green fluorescence, predominantly cytoplasmic, were observed in lung tissues from titi monkeys with fatal TMAdV pneumonia (Fig. 4A–D), but not in lung or intestinal tissues from experimentally TMAdV-infected marmosets (Fig. 4E–H). To confirm these negative findings, necropsy marmoset tissues were then tested for the presence of TMAdV by real-time qPCR and nested PCR. TMAdV was not detected in necropsy tissues from any of the marmosets by these two PCR assays.
This study did not involve human experimentation; the only human material used in this study was throat swab specimens collected from cases with respiratory tract infection during the implementation of the surveillance project on viral aetiology of acute respiratory infection. This study was approved by the second session of the Ethics Review Committee of the National Institute for Viral Disease Control and Prevention in China CDC. Written informed consent for the use of the clinical samples has been obtained from all patients involved in this study.
Pharynx and tonsil secretions of the patients were wiped with disinfection long cotton swabs with gently action, and after samples collection, all samples were transported under a cold chain and preserved at −80°C for further identification. A multiplex one-step reverse transcription-polymerase chain reaction (PCR) was performed to screen for 15 different respiratory viruses (respiratory syncytial virus A and B, influenza virus A and B, parainfluenza virus 1–4, human adenovirus, human enterovirus, human rhinovirus, human metapneumovirus, human bocavirus, and human coronavirus NL63-229E and OC43-HKU1) simultaneously by using a commercial kit (Seeplex RV 15 ACE Detection kit; Seegene, Inc., Seoul, Korea). Adenovirus-positive specimens were cultured and further analysed.
Nasal swabs, serum, and tissues from experimentally infected and control marmosets were screened for TMAdV using a qPCR assay from the adenoviral IVα2 gene with standard curve analysis as previously described. Negative results were then screened further using a more sensitive nested PCR assay from the hexon gene. PCR amplicons were confirmed to be TMAdV by Sanger sequencing. Primers used for the PCR assays are listed in Table S3.
To evaluate the clinical performance, a total of 152 clinical samples previously confirmed HAdV-positive by a GeXP-based multiplex RT-PCR assay were nucleic acid extracted again, then detected in duplicate by the duplex RAA assays. A reference tq-PCR assay for the detection of HAdV 3 and HAdV 7, respectively, was performed in parallel on these samples. A tq-PCR threshold cycle (CT) value of < 38 was determined as a positive result. A slope of the RAA amplification curve > 20 set by the detection device was determined as positive.
Given the clinical presentation of a severe acute viral respiratory illness and the appearance of intranuclear inclusion bodies on histological examination, we strongly suspected that a virus that had eluded detection by conventional assays was the cause of the titi monkey outbreak. Nasal, lung, and liver swab samples collected during necropsy were analyzed using the Virochip,. Microarrays were analyzed using ranked Z-scores to assess the highest-intensity viral probes. From a lung swab sample from an affected monkey, 4 of the top 80 probes on the Virochip corresponded to adenoviruses. Other viruses or viral families with ≥4 probes among the top 80, including chimpanzee herpesvirus (Herpesviridae), bovine viral diarrhea virus (Flaviviridae), and endogenous retroviruses (Retroviridae), were regarded as less likely to cause fulminant pneumonia and hepatitis, so were not pursued any further. The 4 adenovirus probes mapped to 2 different gene regions corresponding to the DNA polymerase and penton base (Fig. 2A). Interestingly, the 4 viral probes were derived from 2 different Adenoviridae genera (SAdV-23, simian adenovirus 23, PAdV-A, porcine adenovirus A, and HAdV-5, human adenovirus 5, in the Mastadenovirus genus; FAdV-D, fowl adenovirus D, in the Aviadenovirus genus), suggesting the presence of a divergent adenovirus that was not a member of any previously known species.
To confirm the Virochip finding of an adenovirus, we used consensus primers to amplify a 301 bp fragment from the hexon gene by PCR. The fragment shared ∼86% nucleotide identity with its closest phylogenetic relatives in GenBank, SAdV-18, an Old World vervet monkey adenovirus, and the human species D adenoviruses. The newly identified adenovirus was designated TMAdV, or titi monkey adenovirus. Specific PCR for TMAdV was then used to screen body fluids and tissues from affected monkeys (Table 1). PCR results were positive from post-necropsy liver and lung tissues as well as from sera, conjunctival swabs, oral swabs, and nasal swabs collected at time of quarantine in 8 different affected monkeys, but were negative from a throat swab from an asymptomatic animal whose other 5 cage mates had become sick. In addition, nasal swabs were negative in 3 asymptomatic, minimal-risk titi monkeys housed in a separate building. To confirm the presence of virus in diseased tissues, we examined lung tissue from affected monkeys by transmission electron microscopy, revealing abundant icosahedral particles characteristic of adenovirus filling the alveoli (Fig. 1D-4).
Next, to assess persistent subclinical infection from TMAdV, we analyzed serum samples from at-risk asymptomatic or affected surviving monkeys 2 months after the outbreak (n = 41). All post-outbreak serum samples were negative for TMAdV by PCR (Table 1). To assess potential TMAdV shedding, stool samples collected from all cages housing titi monkeys 2 months post-outbreak were analyzed by PCR (n = 27), and were found to be negative. In addition, we checked for TMAdV in rectal swab samples from rhesus macaques housed in the same building as the titi monkeys (n = 26) and in pooled droppings from wild rodents (n = 2) living near the titi monkey cages. All macaque and rodent fecal samples were negative for TMAdV by PCR.
We also sought to determine whether PCR assays commonly used to detect human adenoviruses in clinical or public health settings could detect TMAdV. Adenovirus PCR was performed on a TMAdV-positive clinical sample, a TMAdV culture, and a human adenovirus B culture (as a positive control) using an additional 5 pairs of primers, according to previously published protocols,, Three of the 5 primer pairs, designed to detect human respiratory adenoviruses B, C, and E, failed to amplify TMAdV. The remaining 2 pairs of primers, both derived from highly conserved sequences in the hexon gene,, were able to detect TMAdV in culture as well as directly from clinical material.
RAA assays for the detection of HAdV 3 and HAdV 7 with high sensitivity and specificity were demonstrated here. The method is rapid and does not require a thermal cycler. By introducing an IC, the duplex RAA assays successfully eliminated false negative results, thereby producing much more reliable tests. This method might have great potential for clinical use, especially in resource-poor settings.
Using specifically designed RT-PCR primers, we detected viral RNA in 223 of the 285 acute serum samples tested (Table 5). The specificity of the RT-PCR was confirmed by sequencing selected PCR products. None of the 80 sera from patients with respiratory diseases or the 50 sera from healthy subjects was positive using the novel virus-specific RT-PCR.
Sequence alignment was performed by ClustalW Multiple alignment using BioEdit software (version 184.108.40.206). 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.
With the DNase-free VIDISCA-NGS, only eight samples contained sequences of an RNA virus (six HIV-1 and two enterovirus) (Table 2), indicating that background DNA seriously hampered detection of RNA viruses. On the other hand, detection of herpesviruses greatly increased. Without a DNase treatment, 11 samples became VIDISCA-NGS positive: four for HSV-1/2, five for VZV, and two for CMV (Figure 3). The viral load of the nuclease-free VIDISCA-NGS herpesvirus positive samples was higher (median: 1.04 × 105) than the negative samples (median: 4.42 × 103, p = 0.00009, Mann Whitney U test). This association between the virus load and VIDISCA-detection became more visible when 104 DNA copies/mL was taken as a threshold; 11 of 18 samples positive by qPCR with >104 DNA copies/mL were also positive by VIDISCA-NGS, but none below.
We identified several co-infecting DNA viruses (torque teno virus (TTV), n = 5; human papillomavirus (HPVs), n = 5; and hepatitis B virus (HBV), n = 1), which were not included in the routine diagnostics of the CSF samples, but were identified by VIDISCA-NGS (n = 11). Similar to the effects we observed for herpesvirus detection, we hypothesized that more non-herpes DNA viruses would be detected under the DNase-free condition. Surprisingly, no additional non-herpes DNA viruses were identified using the DNase-free method. On the contrary, of the 11 samples containing non-herpes DNA viruses detected by regular VIDISCA-NGS, only four samples were positive when excluding a DNase treatment (Figure 4).
To assess the overall effect of a DNase treatment, we determined the ratio of viral reads, adjusted for sequencing depth, between the two treatment arms for all viruses identified by VIDISCA-NGS in this study (Figure 5). All herpesviruses had substantially more, or a roughly equal number of viral reads in the DNase-free condition. In contrast, the opposite was true for non-herpes DNA and RNA viruses.