The study was supported by the National Key Research and Development Plan (Grant no. 2016YFD0500401) and the National Natural Science Foundation of China (Grant no. 31302043).

The use of mice in this protocol was approved by the Administrative Committee on Animal Welfare of the Institute of Military Veterinary Medicine, the Academy of Military Medical Sciences, China (Laboratory Animal Care and Use Committee Authorization, permit number JSY-DW-2010-02). All institutional and national guidelines for the care and use of laboratory animals were followed.
1. RNA Extraction
<ol>
	<li>Extract RNA from rabies-suspected brain tissue, skin biopsies, saliva, or CSF or from RABV-infected cell culture, using guanidinium isothiocyanate-phenol-chloroform-based extraction methods or commercially available viral RNA extraction kits. Use the prepared RNA immediately or store it at -80 &#176;C until required.</li>
</ol>
2. Reverse Transcription of the Viral RNA
<ol>
	<li>Remove the RT reagents listed in Table 1 from the freezer, keep them on ice, and thaw and vortex them before use.</li>
	<li>Prepare 12 &#181;L of RT reaction mix in a 0.2 mL PCR tube with the reagents listed in Table 1. Allow for pipetting variations by preparing a volume of master mix at least one reaction size greater than required.</li>
	<li>Add 8 &#181;L sample, positive control RNA or negative control to the RT reaction mix within a PCR workstation in a template room. The RT positive control is RNA extracted from the cell culture infected with fixed RABV strain CVS-11 (challenge virus standard-11) and stored at -80 &#176;C. The negative control contains RNase-free ddH2O.</li>
	<li>Mix the contents of the RT tubes by vortexing; then, centrifuge briefly.</li>
	<li>Load the reaction tubes into a thermal cycler. Set up the cDNA synthesis program with the following conditions: 42 &#176;C for 90 min, 95 &#176;C for 5 min, and 4 &#176;C on hold. Set the reaction volume to 20 &#181;L. Start the RT run.</li>
</ol>
3. First-round PCR
<ol>
	<li>Keep the PCR reagents listed in Table 2 on ice in a clean room until use; then, thaw and vortex them.</li>
	<li>Prepare the first-round PCR mix in a 0.2 mL PCR tube with the reagents listed in Table 2.</li>
	<li>Add a 2 &#181;L sample of cDNA or plasmid into the first-round PCR mix within a PCR workstation in a template room. The PCR positive control is CVS-11 cDNA prepared as mentioned in step 2.3 for the above RT method. The PCR negative control is ddH2O.</li>
	<li>Transfer the sealed tubes to a PCR thermal cycler and cycle using the parameters listed in Table 3.</li>
</ol>
4. Second-round PCR
<ol>
	<li>Prepare the second-round PCR mix in a 0.2 mL PCR tube using the reagents listed in Table 4.</li>
	<li>Add 2 &#181;L of first-round PCR product into the second-round PCR mix. In addition, include ddH2O as a negative control of the second-round PCR.</li>
	<li>Perform PCR thermal cycling using the same parameters as given in step 3.4.</li>
</ol>
5. Analysis of the PCR Products by Electrophoresis on Agarose Gels
<ol>
	<li>Prepare a 1.5% agarose gel by adding 1.5 g of agarose to 100 mL of Tris-acetate-EDTA (TAE) and dissolving it thoroughly by heating it in a microwave oven.</li>
	<li>Add ethidium bromide (EB) (at a final concentration of 0.01%) or another commercial EB substitution. Pour the gel into the mold and leave it to solidify at ambient temperature for at least 30 min.</li>
	<li>Prepare the loading samples by mixing 5 &#181;L of each PCR product with 1 &#181;L of 6x loading buffer.</li>
	<li>Load the samples and suitable DNA marker separately into the wells and run the gel for approximately 30-45 min at 120 V until the dye line is approximately 75%-80% down the gel.</li>
	<li>Turn off the power, disconnect the electrodes from the power source, and then, carefully remove the gel from the gel box.</li>
	<li>Use a UV gel documenting device to visualize and photograph the DNA fragments.</li>
</ol>
6. Characterization of Nested RT-PCR
<ol>
	<li>6.1. Specificity and sensitivity for the detection of 18 lyssaviruses plasmids 
	<ol>
		<li>Order 18 commercial plasmids containing the full N gene of each lyssavirus (16 ICTV species and two novel species) for the PCR.</li>
		<li>Calculate the copy number of the plasmid using Avogadro&#8217;s number (NA) and the following formula. 
		[(g/&#181;L plasmid DNA)/(plasmid length in bp x 660)] x 6.022 x 1023 = number of molecules/&#181;L</li>
		<li>Prepare stock solutions (2.24 x 109 molecules/&#181;L) of all 18 plasmids in ddH2O.</li>
		<li>Perform nine 10-fold serial dilutions of all 18 plasmids in ddH2O. Dilute 10 &#181;L of each plasmid stock with 90 &#181;L of ddH2O. Vortex and centrifuge briefly.</li>
		<li>Perform PCR amplification as described in sections 3-5.</li>
		<li>Analyze the specificity and sensitivity of the nested PCR by detecting a series of lyssavirus plasmids.</li>
	</ol>
	</li>
	<li>Determination of the detection limit
	<ol>
		<li>Adjust the titer of the rabies virus strain CVS-11 cell culture to 105.5 TCID50/mL (virus titer determined according to the OIE Manual)5.</li>
		<li>Perform five 10-fold serial dilutions of CVS-11 (stock solution is 105.5 TCID50/mL) as described in step 6.1.4.</li>
		<li>Perform the RNA extraction and nested RT-PCR amplification procedures of all viral dilutions as described in sections 1-5.</li>
	</ol>
	</li>
	<li>Comparison of the nested RT-PCR with the &#34;gold standard&#34;&#160;FAT
	<ol>
		<li>Test all clinical samples by nested RT-PCR, and then, confirm the results with the FAT5 and N gene sequencing9.</li>
		<li>Use normalized data for a statistical analysis with SAS 9.1. Use the kappa test and McNemar&#8217;s chi-squared test for a statistical comparison of the diagnostic tests (SAS command: proc freq; table/agree). Calculate confidence intervals assuming abinomial distribution.</li>
	</ol>
	</li>
	<li>Evaluation of the efficacy in testing degraded samples
	<ol>
		<li>Expose two confirmed clinical brain tissue samples of rabid dogs at 37 &#176;C.</li>
		<li>Assay the two samples on each day of exposure at 37 &#176;C by nested RT-PCR, the FAT, and the MIT5.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Hemachudha, T., Laothamatas, J., Rupprecht, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+rabies:+a+disease+of+complex+neuropathogenetic+mechanisms+and+diagnostic+challenges.">Human rabies: a disease of complex neuropathogenetic mechanisms and diagnostic challenges.</a> Lancet Neurology. 1 (2), 101-109 (2002).</li><li>Amarasinghe, G. K., Arechiga Ceballos, N. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Taxonomy+of+the+order+Mononegavirales:+update+2018.">Taxonomy of the order Mononegavirales: update 2018.</a> Archives of Virology. 163 (8), 2283-2294 (2018).</li><li>Nokireki, T., Tammiranta, N., Kokkonen, U. M., Kantala, T., Gadd, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tentative+novel+lyssavirus+in+a+bat+in+Finland.">Tentative novel lyssavirus in a bat in Finland.</a> Transboundary and Emerging Diseases. 65 (3), 593-596 (2018).</li><li>Hu, S. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lyssavirus+in+Japanese+Pipistrelle,+Taiwan.">Lyssavirus in Japanese Pipistrelle, Taiwan.</a> Emerging Infectious Diseases. 24 (4), 782-785 (2018).</li><li>Rupprecht, C. E., Fooks, A. R., Abela-Ridder, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laboratory+techniques+in+rabies.">Laboratory techniques in rabies.</a> World Health Organization. , 74-195 (2018).</li><li>Mani, R. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Utility+of+real-time+Taqman+PCR+for+antemortem+and+postmortem+diagnosis+of+human+rabies.">Utility of real-time Taqman PCR for antemortem and postmortem diagnosis of human rabies.</a> Journal of Medical Virology. 86 (10), 1804-1812 (2014).</li><li>Fooks, A. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Emerging+technologies+for+the+detection+of+rabies+virus:+challenges+and+hopes+in+the+21st+century.">Emerging technologies for the detection of rabies virus: challenges and hopes in the 21st century.</a> PLOS Neglected Tropical Diseases. 3 (9), 530 (2009).</li><li>Jiang, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+outbreak+of+pig+rabies+in+Hunan+province,+China.">An outbreak of pig rabies in Hunan province, China.</a> Epidemiology and Infection. 136 (4), 504-508 (2008).</li><li>Liu, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+the+complete+genome+of+the+first+Irkut+virus+isolate+from+China:+comparison+across+the+Lyssavirus+genus.">Analysis of the complete genome of the first Irkut virus isolate from China: comparison across the Lyssavirus genus.</a> Molecular Phylogenetics and Evolution. 69 (3), 687-693 (2013).</li><li>Chen, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Possible+Transmission+of+Irkut+Virus+from+Dogs+to+Humans.">Possible Transmission of Irkut Virus from Dogs to Humans.</a> Biomedical and Environmental Sciences. 31 (2), 146-148 (2018).</li><li>Feng, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disease+outbreaks+caused+by+steppe-type+rabies+viruses+in+China.">Disease outbreaks caused by steppe-type rabies viruses in China.</a> Epidemiology and Infection. 143 (6), 1287-1291 (2015).</li><li>Feng, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Livestock+rabies+outbreaks+in+Shanxi+province,+China.">Livestock rabies outbreaks in Shanxi province, China.</a> Archives of Virology. 161 (10), 2851-2854 (2016).</li></ol>

The authors have nothing to disclose.

Currently, RABV is a major lyssavirus responsible for nearly all human and animal rabies in China, as well as in other countries. In addition, an IRKV variant was first identified from a Murina leucogaster bat in the Jilin province in Northeast China in 201210, and it has been reported to cause a dog&#8217;s death in the Liaoning Province in 201711. Most recently, a novel lyssavirus, TWBLV, was also identified from a Japanese pipistrelle bat in Taiwan, China in 2017. These results suggest that the effective detection of other lyssaviruses is also important to prevent the spill-over of the bat-borne lyssaviruses. In this regard, the pan-lyssavirus nested RT-PCR targeting the most conserved N gene region is a very useful tool, and the results have shown that it can effectively detect all 16 ICTV-approved and two novel lyssavirus species identified so far, including genetically divergent RABVs and IRKV in China (data on IRKV strain not shown). However, it is also interesting to note that ARAV was detected only by the second-round PCR primers, while IKOV was detected only by first-round PCR primers (Figure 1). To investigate the cause of this discrepancy, sequence comparison of all 18 lyssaviruses within the four primer regions was conducted, with the results showing that the 3&#39; end nucleotide T of outer primer N829 in the first-round PCR and the second nucleotide T at the 3&#39; end of inner primer N371F in the second-round PCR were not identical to the corresponding positions of ARAV (G) and IKOV (A) (Figure 2). Once the T of primer N829 was changed to the G of ARAV and the T of primer N371F changed to the A of IKOV, both viruses were successfully detected in PCR (data not shown). This result demonstrates the critical role of the 3&#39; end or near-end nucleotides of primers in the successful amplification of the target region.
To evaluate the sensitivity and specificity of the method in the detection of rabies virus for clinical diagnosis and surveillance, 9,624 animal brain tissue samples were tested in the last 10 years by the FAT and nested RT-PCR in parallel. Of the 165 samples that tested rabies positive by nested RT-PCR, 162 were detected as positive by the FAT; therefore, the two methods show 99.07% accordance. The RABVs detected in these 165 samples could be classified into different lineages in Asian, Arctic-related, and Cosmopolitan clades12,13, indicating that nested RT-PCR can cover the various genetic clades of RABVs. Three highly decayed clinical specimens were tested positive by nested RT-PCR but negative by the FAT. This result is consistent with the evaluation of the efficacy in testing two degraded samples as shown in the representative results section, indicating that nested RT-PCR is more sensitive than the FAT in the detection of viruses in highly decayed specimens.
The performance of nested RT-PCR was further confirmed by participation in the International Laboratory Comparison Test (ILCT) organized by the EU Reference Laboratory for Rabies at the French Agency for Food, Environmental and Occupational Health &#38; Safety (ANSES) in 2010, using a set of 12 samples (one each of RABV, EBLV-1, EBLV-2, and ABLV as positive controls, one negative control, and seven blinded samples). In the test, nested RT-PCR successfully identified all the lyssaviruses with a 100% consistency. In 2018, the method was approved as a National Standard of Rabies Diagnoses (GB/T36789-2018) by the National Technical Standardization Committee of Animal Health of China.

Rabies is a worldwide zoonotic disease caused by viruses within the genus Lyssavirus1. Lyssaviruses (family&#160;Rhabdoviridae) are single-negative-stranded RNA viruses with an approximately 12 kb genome that encodes five proteins: N, phosphoprotein (P), matrix protein (M), glycoprotein (G), and the large protein or polymerase (L).&#160;Based on nucleotide sequences of the N gene, genetic distance, and antigenic patterns, the lyssaviruses have been divided into 16 species, comprising classical rabies virus (RABV) and the rabies-related viruses (RRV): Lagos bat virus (LBV), Duvenhage virus (DUVV), Mokola virus (MOKV), European bat lyssavirus 1 (EBLV-1), European bat lyssavirus 2 (EBLV-2), Australian bat lyssavirus (ABLV), Aravan virus (ARAV), Ikoma virus (IKOV), Bokeloh bat lyssavirus (BBLV), Gannoruwa bat lyssavirus (GBLV), Irkut virus (IRKV), Khujand virus (KHUV), West Caucasian bat virus (WCBV), Shimoni bat virus (SHIBV), and Lleida bat lyssavirus (LLEBV)2. Recently, two additional lyssaviruses have been identified: Kotalahti bat lyssavirus (KBLV) isolated from a Brandt&#8217;s bat (Myotis brandtii) in Finland in 20173 and Taiwan bat lyssavirus (TWBLV) isolated from a Japanese pipistrelle (Pipistrellus abramus) in Taiwan, China in 2016&#8211;20174.
All mammals are susceptible to rabies; however, no gross pathognomonic lesions or specific clinical signs permit its identification, and diagnosis can only be made in the laboratory5. The most widely used method for rabies diagnosis is the FAT, which is considered as the gold standard by both the WHO and the OIE5,6. Nevertheless, the FAT can produce unreliable results on degraded/autolyzed brain tissue samples. Additionally, it cannot be used to assay biological fluid specimens such as cerebrospinal fluid (CSF), saliva, and urine, thereby largely precluding its employment in antemortem diagnosis7. Alternative conventional diagnostic tests, such as the rabies tissue culture infection test (RTCIT) and the mouse inoculation test (MIT), require several days6, a major drawback when a rapid diagnosis is essential.
Various molecular diagnostic tests (e.g., the detection of viral RNA by RT-PCR, the PCR&#8211;enzyme-linked immunosorbent assay [PCR-ELISA], in situ hybridization, and real-time PCR) are used as rapid and sensitive techniques for rabies diagnosis8. RT-PCR is now recommended by OIE for routine rabies diagnosis, and a heminested (hn) PCR is described in the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals to detect all lyssaviruses5. Here we describe a pan-lyssavirus nested RT-PCR, which allows the specific and sensitive detection of all 18 lyssavirus species comparable to or exceeding that obtained by the FAT. The principle of the method is an RT of the target RNA (conserved region of the lyssavirus N gene) into cDNA, followed by the amplification of the cDNA by two rounds of PCR. The cDNA undergoes the first-round PCR with outer primers to amplify an 845 bp fragment; then, the second-round PCR uses the first-round PCR product as a template to amplify a 371 bp fragment with inner primers. The two rounds of PCR significantly increase the sensitivity of the assay.

<table><tbody><tr><td>50 &amp;times; TAE</td><td>Various</td><td>Various</td><td></td></tr><tr><td>6 &amp;times; loading buffer</td><td>TakaRa</td><td>9156</td><td></td></tr><tr><td>Agarose</td><td>US Everbright&amp;reg; Inc</td><td>A-2015-100g</td><td></td></tr><tr><td>ddH&lt;sub&gt;2&lt;/sub&gt;O</td><td>Various</td><td>Various</td><td></td></tr><tr><td>DL 2,000 Marker</td><td>Takara</td><td>3427A</td><td></td></tr><tr><td>dNTPs (10 mM)</td><td>TakaRa</td><td>4019</td><td></td></tr><tr><td>dNTPs (2.5 mM)</td><td>TakaRa</td><td>4030</td><td></td></tr><tr><td>Electrophoresis System</td><td>Tanon</td><td>EPS300</td><td></td></tr><tr><td>Ex-Taq (5 U/&amp;mu;L)</td><td>TakaRa</td><td>RR001</td><td></td></tr><tr><td>Gel Imaging System</td><td>UVITEC</td><td>Fire Reader</td><td></td></tr><tr><td>Microcentifuge tubes</td><td>Various</td><td>Various</td><td></td></tr><tr><td>M-MLV reverse transcriptase (200 IU/&amp;micro;L)</td><td>TakaRa</td><td>2641A&amp;nbsp;</td><td></td></tr><tr><td>NanoDrop 1000 Spectrophotometer</td><td>Thermoscientific</td><td>ND1000</td><td></td></tr><tr><td>Oligo (dT)&lt;sub&gt;15&lt;/sub&gt;</td><td>TakaRa</td><td>3805</td><td></td></tr><tr><td>PCR Machine</td><td>BIO-RAD</td><td>T100</td><td></td></tr><tr><td>PCR Tubes</td><td>Various</td><td>Various</td><td></td></tr><tr><td>Phusion High-Fidelity DNA Polymearase</td><td>NEW ENGLAND BioLabs</td><td>M0530S</td><td></td></tr><tr><td>Pipettors</td><td>Various</td><td>Various</td><td></td></tr><tr><td>Random Primer</td><td>TakaRa</td><td>3801</td><td></td></tr><tr><td>RNase Inhibitor (40 IU/&amp;micro;L)</td><td>TakaRa</td><td>2313A</td><td></td></tr><tr><td>RNase-free ddH&lt;sub&gt;2&lt;/sub&gt;O</td><td>TakaRa</td><td>9102</td><td></td></tr><tr><td>Taq Buffer (10&amp;times;)</td><td>TakaRa</td><td>9152A</td><td></td></tr><tr><td>Tips</td><td>Various</td><td>Various</td><td></td></tr><tr><td>Vortex mixer</td><td>Various</td><td>Various</td><td></td></tr></tbody></table>

evaluation of a universal nested reverse transcription polymerase chain reaction for the detection of lyssaviruses

To detect rabies virus and other member species of the genus Lyssavirus within the family Rhabdoviridae, the pan-lyssavirus nested reverse transcription polymerase chain reaction (nested RT-PCR) was developed to detect the conserved region of the nucleoprotein (N) gene of lyssaviruses. The method applies reverse transcription (RT) using viral RNA as template and oligo (dT)15 and random hexamers as primers to synthesize the viral complementary DNA (cDNA). Then, the viral cDNA is used as a template to amplify an 845 bp N gene fragment in first-round PCR using outer primers, followed by second-round nested PCR to amplify the final 371 bp fragment using inner primers. This method can detect different genetic clades of rabies viruses (RABV). The validation, using 9,624 brain specimens from eight domestic animal species in 10 years of clinical rabies diagnoses and surveillance in China, showed that the method has 100% sensitivity and 99.97% specificity in comparison with the direct fluorescent antibody test (FAT), the gold standard method recommended by the World Health Organization (WHO) and the World Organization for Animal Health (OIE). In addition, the method could also specifically amplify the targeted N gene fragment of 15 other approved and two novel lyssavirus species in the 10th Report of the International Committee on Taxonomy of Viruses (ICTV) as evaluated by a mimic detection of synthesized N gene plasmids of all lyssaviruses. The method provides a convenient alternative to FAT for rabies diagnosis and has been approved as a National Standard (GB/T36789-2018) of China.

Results of nested RT-PCR to detect 18 lyssavirus species are shown in Figure 1. All PCR positive controls showed the expected 845 bp in the first- and 371 bp in the second-round amplifications with no band in the negative control. All 18 lyssaviruses produced the expected 845 and/or 371 bp bands, indicating that the nested RT-PCR detected all 18 lyssaviruses. Sixteen lyssaviruses plasmids had efficient amplification in two rounds of PCR, but two, namely ARAV and IKOV, had amplification in either the first- or second-round PCR. The sensitivity of the method varied in the detection of different lyssavirus plasmids, with limits ranging from 2.24 x 100 to 2.24 x 105 molecules/&#181;L, as shown in Table 6. These differences can be attributed to the mismatches between the primers and templates due to viral sequence diversity. Furthermore, the sensitivity of detecting rabies virus CVS-11 in cell culture was 102.5 TCID50/mL.
A total of 9,624 brain tissues from clinical specimens were tested by nested RT-PCR in comparison with the FAT and the results are summarized in Table 7, which shows that nested RT-PCR had a 100% sensitivity (CI, 97.75% to 100%) and a 99.97% specificity (CI, 99.91% to 99.99%). The accordance between the two methods was 99.07%. Three tests that were positive by nested RT-PCR but negative by the FAT were of highly decayed clinical material. These three specimens were confirmed as RABV positive by N gene sequencing.
Comparison of the test performance in the detection of the two brain specimens incubated at 37 &#176;C (step 6.4.1 of the protocol) indicated that the nested RT-PCR could effectively detect virus in decayed brain tissues for least 17 days postdegradation, which is for a longer period of time when compared with only 7 days by the FAT and not even 1 day by the MIT. This result shows that nested RT-PCR is more sensitive in the detection of degraded samples than the FAT and the MIT.
To further validate the nested RT-PCR, 10 rabies laboratories in China were invited to conduct tests on a set of specimens. Of these, eight laboratories were provided a set of 10 blinded animal brain tissues from our laboratory, including RABV positive and negative specimens. The other two laboratories used their own specimens. All specimens had been archived and confirmed by the FAT previously. All 10 laboratories obtained results by nested RT-PCR in 100% accordance with the FAT, with no false-negatives or false-positives (Table 8), indicating that the nested RT-PCR had a high specificity and reproducibility.
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Components</td>
			<td>Volume per reaction (&#181;L)</td>
		</tr>
		<tr>
			<td>dNTPs (2.5 mM )</td>
			<td>4</td>
		</tr>
		<tr>
			<td>Random Primer (50 &#956;M)</td>
			<td>1.5</td>
		</tr>
		<tr>
			<td>Oligo(dT) 15 (50 &#956;M)</td>
			<td>0.5</td>
		</tr>
		<tr>
			<td>M-MLV buffer (5x)</td>
			<td>4</td>
		</tr>
		<tr>
			<td>M-MLV reverse transcriptase (200 IU/&#181;L)</td>
			<td>1</td>
		</tr>
		<tr>
			<td>RNasin (40 IU/&#181;L)</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Total volume</td>
			<td>12</td>
		</tr>
	</tbody>
</table>
Table 1: Reagents of reverse transcription for cDNA synthesis.
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Components</td>
			<td>Volume per reaction (&#181;L)</td>
		</tr>
		<tr>
			<td>dNTPs (10 mM)</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Ex-Taq (5 U/&#956;L)</td>
			<td>0.3</td>
		</tr>
		<tr>
			<td>Taq Buffer (10x)</td>
			<td>5</td>
		</tr>
		<tr>
			<td>N127 (20 &#956;M)</td>
			<td>1</td>
		</tr>
		<tr>
			<td>N829 (20 &#956;M)</td>
			<td>1</td>
		</tr>
		<tr>
			<td>dd H2O</td>
			<td>39.7</td>
		</tr>
		<tr>
			<td>Total volume</td>
			<td>48</td>
		</tr>
	</tbody>
</table>
Table 2: Reagents of the first-round PCR.
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Temperature</td>
			<td>Time</td>
			<td>Cycles</td>
		</tr>
		<tr>
			<td>94 &#176;C</td>
			<td>2 min</td>
			<td>1</td>
		</tr>
		<tr>
			<td>94&#160;&#176;C</td>
			<td>30 s</td>
			<td>35</td>
		</tr>
		<tr>
			<td>56&#160;&#176;C</td>
			<td>30 s</td>
			<td>35</td>
		</tr>
		<tr>
			<td>72&#160;&#176;C</td>
			<td>40 s</td>
			<td>35</td>
		</tr>
		<tr>
			<td>72&#160;&#176;C</td>
			<td>10 min</td>
			<td>1</td>
		</tr>
		<tr>
			<td>4&#160;&#176;C</td>
			<td>&#8734;</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 3: Cycling parameters of the first- and second-round PCR.
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Components</td>
			<td>Volume per reaction (&#181;L)</td>
		</tr>
		<tr>
			<td>dNTPs (10 mM)</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Ex-Taq (5 U/&#956;L)</td>
			<td>0.3</td>
		</tr>
		<tr>
			<td>Taq Buffer (10x)</td>
			<td>5</td>
		</tr>
		<tr>
			<td>N371F (20 &#956;M)</td>
			<td>1</td>
		</tr>
		<tr>
			<td>N371R (20 &#956;M)</td>
			<td>1</td>
		</tr>
		<tr>
			<td>dd H2O</td>
			<td>39.7</td>
		</tr>
		<tr>
			<td>Total volume</td>
			<td>48</td>
		</tr>
	</tbody>
</table>
Table 4: Reagents of the second-round PCR.
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Primer Name</td>
			<td>Details</td>
			<td>Direction</td>
			<td colspan="2">Sequence (5&#8217;-3&#8217;)</td>
			<td>Nucleotide position</td>
			<td>Product Size</td>
		</tr>
		<tr>
			<td>N127</td>
			<td>The first round PCR&#160;</td>
			<td>Forward</td>
			<td colspan="2">ATGTAACNCCTCTACAATGG</td>
			<td>-19~0</td>
			<td>845bp</td>
		</tr>
		<tr>
			<td>N829</td>
			<td>The first round PCR&#160;</td>
			<td>Reverse</td>
			<td colspan="2">GCCCTGGTTCGAACATTCT</td>
			<td>807~825</td>
			<td>845bp</td>
		</tr>
		<tr>
			<td>N371F</td>
			<td>The second round PCR&#160;</td>
			<td>Forward</td>
			<td colspan="2">ACAATGGAKKCTGACAARATTG</td>
			<td>-6~15</td>
			<td>371bp</td>
		</tr>
		<tr>
			<td>N371R</td>
			<td>The second round PCR</td>
			<td>Reverse</td>
			<td colspan="2">CCTGYYWGAGCCCAGTTVCCYTC</td>
			<td>345~367</td>
			<td>371bp</td>
		</tr>
	</tbody>
</table>
Table 5: Primer sequences of the first- and second-round PCR. Degenerate bases: N (A/T/C/G), K (G/T), R (A/G), Y (C/T), WV (G/A/C).
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Lyssavirus species</td>
			<td>Strain</td>
			<td>Template plasmid (molecules /&#181;L)</td>
		</tr>
		<tr>
			<td>RABV</td>
			<td>GQ918139.1</td>
			<td>2.24 x 101</td>
		</tr>
		<tr>
			<td>LBV</td>
			<td>EU293110.1</td>
			<td>2.24&#160;x&#160;102</td>
		</tr>
		<tr>
			<td>MOKV</td>
			<td>KF155005.1</td>
			<td>2.24&#160;x&#160;101</td>
		</tr>
		<tr>
			<td>DUVV</td>
			<td>EU293119.1</td>
			<td>2.24&#160;x&#160;101</td>
		</tr>
		<tr>
			<td>EBLV-1</td>
			<td>EF157976.1</td>
			<td>2.24&#160;x&#160;101</td>
		</tr>
		<tr>
			<td>EBLV-2</td>
			<td>KF155004.1</td>
			<td>2.24&#160;x&#160;101</td>
		</tr>
		<tr>
			<td>ABLV</td>
			<td>GU992312.1</td>
			<td>2.24&#160;x&#160;100</td>
		</tr>
		<tr>
			<td>IRKV</td>
			<td>NC_020809.1</td>
			<td>2.24&#160;x&#160;101</td>
		</tr>
		<tr>
			<td>WCBV</td>
			<td>NC_025377.1</td>
			<td>2.24&#160;x&#160;101</td>
		</tr>
		<tr>
			<td>KHUV</td>
			<td>NC_025385.1</td>
			<td>2.24&#160;x&#160;103</td>
		</tr>
		<tr>
			<td>ARAV</td>
			<td>NC_020808.1</td>
			<td>2.24&#160;x&#160;102</td>
		</tr>
		<tr>
			<td>SHIBV</td>
			<td>NC_025365.1</td>
			<td>2.24&#160;x&#160;101</td>
		</tr>
		<tr>
			<td>BBLV</td>
			<td>NC_025251.1</td>
			<td>2.24&#160;x&#160;101</td>
		</tr>
		<tr>
			<td>IKOV</td>
			<td>NC_018629.1</td>
			<td>2.24&#160;x&#160;105</td>
		</tr>
		<tr>
			<td>LLEBV</td>
			<td>NC_031955.1</td>
			<td>2.24&#160;x&#160;103</td>
		</tr>
		<tr>
			<td>GBLV</td>
			<td>NC_031988.1</td>
			<td>2.24&#160;x&#160;105</td>
		</tr>
		<tr>
			<td>KBLV</td>
			<td>MF960865.1</td>
			<td>2.24&#160;x&#160;101</td>
		</tr>
		<tr>
			<td>TWBLV</td>
			<td>MF472710.1</td>
			<td>2.24&#160;x&#160;101</td>
		</tr>
	</tbody>
</table>
Table 6: Detection limit of nested RT-PCR of 18 lyssaviruses.
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Standard method and result</td>
			<td></td>
			<td>RT-nPCR 
			Positive</td>
			<td>RT-nPCR 
			Negative</td>
			<td>Correlation (%)</td>
		</tr>
		<tr>
			<td>FAT</td>
			<td>Positive 
			Negative</td>
			<td>162 
			3</td>
			<td>0 
			9459</td>
			<td>99.07</td>
		</tr>
	</tbody>
</table>
Table 7: Correlation between nested RT-PCR and the FAT in the detection of RABVs in clinical specimens.
<img alt="Table 8" src="/files/ftp_upload/59428/59428table8.jpg" /> 
Table 8: Validation results of nested RT-PCR by 10 laboratories.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59428/59428fig1.jpg" /> 
Figure 1: Detection of 18 lyssaviruses by nested RT-PCR. (A) The result of the first-round PCR. (B) The result of the second-round PCR. M = DL 2000 DNA marker. Lanes 1&#8211;18 = RABV, LBV, MOKV, DUVV, EBLV-1, EBLV-2, ABLV, ARAV, IKOV, BBLV, GBLV, IRKV, KHUV, LLEBV, SHIBV, WCBV, KBLV, and TWBLV, respectively. Lane 19 = PCR positive control; lane 20 = negative control for the first-round PCR; lane 21 = negative control for the second-round PCR. A positive PCR result shows a band at 845 bp in the first round and 371 bp in the second-round PCR. The amplicon of ARAV is not visible in the first round, but visible in the second-round PCR (lane 8), while the amplicon of IKOV is visible in the first round, but not visible in the second-round PCR (lane 9). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59428/59428fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59428/59428fig2.jpg" /> 
Figure 2: Comparison with primer sequences shows the differential nucleotides in primer regions of 18 lyssavirus species. N127 and N829 were outer primers, N371F and N371R were inner primers. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59428/59428fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Evaluation of a Universal Nested Reverse Transcription Polymerase Chain Reaction for the Detection of Lyssaviruses at JoVE.com

Evaluation of a Universal Nested Reverse Transcription Polymerase Chain Reaction for the Detection of Lyssaviruses

A pan-lyssavirus nested reverse transcription polymerase chain reaction has been developed to detect specifically all known lyssaviruses. Validation using rabies brain samples of different animal species showed that this method has a sensitivity and specificity equivalent to the gold standard fluorescent antibody test and could be used for routine rabies diagnosis.

To detect rabies virus and other member species of the genus Lyssavirus within the family Rhabdoviridae, the pan-lyssavirus nested reverse transcription ...

evaluation-universal-nested-reverse-transcription-polymerase-chain

Nanyang Technological University

Research

JoVE Journal

Genetics

8.3K Views.  Academy of Military Medical Sciences. De geneste RT-PCR is ontwikkeld om alle Lyssavirussen te detecteren, inclusief alle 16 ictv-goedgekeurde en twee nieuwe Lyssavirussoorten. De validatie van deze methode is bewezen door het gebruik van meer dan 9000 dierlijke hersenen specimens in 10 jaar van klinische rabiës diagnose en surveillance in China. De geneste RT-PCR kan niet alleen geloofwaardig resultaat opleveren op verse hersenweefselmonsters, maar ook op gedegradeerde monsters.De overstroming deze methode kan worden geb...

Video: Evaluation of a Universal Nested Reverse Transcription Polymerase Chain Reaction for the Detection of Lyssaviruses

Multiplexed Isothermal Amplification Based Diagnostic Platform to Detect Zika, Chikungunya, and Dengue 1

Zika, dengue, and chikungunya viruses are transmitted by mosquitoes, causing diseases with similar patient symptoms. However, they have different downstream patient-to-patient transmission potentials, and require very different patient treatments. Thus, recent Zika outbreaks make it urgent to develop tools that rapidly discriminate these viruses in patients and trapped mosquitoes, to select the correct patient treatment, and to understand and manage their epidemiology in real time.
Unfortunately, current diagnostic tests, including those receiving 2016 emergency use authorizations and fast-track status, detect viral RNA by reverse transcription polymerase chain reaction (RT-PCR), which requires instrumentation, trained users, and considerable sample preparation. Thus, they must be sent to &#34;approved&#34; reference laboratories, requiring time. Indeed, in August 2016, the Center for Disease Control (CDC) was asking pregnant women who had been bitten by a mosquito and developed a Zika-indicating rash to wait an unacceptable 2 to 4 weeks before learning whether they were infected. We very much need tests that can be done on site, with few resources, and by trained but not necessarily licensed personnel.
This video demonstrates an assay that meets these specifications, working with urine or serum (for patients) or crushed mosquito carcasses (for environmental surveillance), all without much sample preparation. Mosquito carcasses are captured on paper carrying quaternary ammonium groups (Q-paper) followed by ammonia treatment to manage biohazards. These are then directly, without RNA isolation, put into assay tubes containing freeze-dried reagents that need no chain of refrigeration. A modified form of reverse transcription loop-mediated isothermal amplification with target-specific fluorescently tagged displaceable probes produces readout, in 30 min, as a three-color fluorescence signal. This is visualized with a handheld, battery-powered device with an orange filter. Forward contamination is prevented with sealed tubes, and the use of thermolabile uracil DNA glycosylase (UDG) in the presence of dUTP in the amplification mixture.

Current multiplexed diagnostics to detect Zika, chikungunya, and dengue viruses require complex sample preparation and expensive instrumentation, and are difficult to use in low resource environments. We show a diagnostic that uses isothermal amplification with target-specific strand displaceable probes to detect and differentiate these viruses with high sensitivity and specificity.

Zika, dengue, and chikungunya viruses are transmitted by mosquitoes, causing diseases with similar patient symptoms. However, they have different ...

Biochemistry

Standard Operating Procedure for Lyssavirus Surveillance of the Bat Population in Taiwan

Viruses within the genus Lyssavirus are zoonotic pathogens, and at least seven lyssavirus species are associated with human cases. Because bats are natural reservoirs of most lyssaviruses, a lyssavirus surveillance program of bats has been conducted in Taiwan since 2008 to understand the ecology of these viruses in bats. In this program, non-governmental bat conservation organizations and local animal disease control centers cooperated to collect dead bats or bats dying of weakness or illness. Brain tissues of bats were obtained through necropsy and subjected to direct fluorescent antibody test (FAT) and reverse transcription polymerase chain reaction (RT-PCR) for detection of lyssavirus antigens and nucleic acids. For the FAT, at least two different rabies diagnosis conjugates are recommended. For the RT-PCR, two sets of primers (JW12/N165-146, N113F/N304R) are used to amplify a partial sequence of the lyssavirus nucleoprotein gene. This surveillance program monitors lyssaviruses and other zoonotic agents in bats. Taiwan bat lyssavirus is found in two cases of the Japanese pipistrelle (Pipistrellus abramus) in 2016&#8211;2017. These findings should inform the public, health professionals, and scientists of the potential risks of contacting bats and other wildlife.

This protocol introduces a standard laboratory operating procedure for diagnostic testing of lyssavirus antigens in bats in Taiwan.

Viruses within the genus Lyssavirus are zoonotic pathogens, and at least seven lyssavirus species are associated with human cases. Because bats are ...

Environment

Reverse Genetics to Engineer Positive-Sense RNA Virus Variants

The lack of a convenient method for the iterative generation of diverse full-length viral variants has impeded the study of directed evolution in RNA viruses. By integrating a full RNA genome error-prone PCR and reverse genetics, random genome-wide substitution mutagenesis can be induced. We have developed a method using this technique to synthesize diverse libraries to identify viral mutants with phenotypes of interest. This method, called full-length mutant RNA synthesis (FL-MRS), offers the following advantages: (i) the ability to create a large library via a highly efficient one-step error-prone PCR; (ii) the ability to create groups of libraries with varying levels of genetic diversity by manipulating the fidelity of DNA polymerase; (iii) the creation of a full-length PCR product that can directly serve as a template for mutant RNA synthesis; and (iv) the ability to create RNA that can be delivered into host cells as a non-selected input pool to screen for viral mutants of the desired phenotype. We have found, using a reverse genetics approach, that FL-MRS is a reliable tool to study viral-directed evolution at all stages in the life cycle of the hepatitis C virus, JFH1 isolate. This technique appears to be an invaluable tool to employ directed evolution to understand adaptation, replication, and the role of viral genes in pathogenesis and antiviral resistance in positive-sense RNA viruses.

The protocol describes a method for introducing controllable genetic diversity in the hepatitis C virus genome by combining full-length mutant RNA synthesis using error-prone PCR and reverse genetics. The method provides a model for phenotype selection and can be used for 10 kb long positive-sense RNA virus genomes.

The lack of a convenient method for the iterative generation of diverse full-length viral variants has impeded the study of directed evolution in RNA ...

Pan-lyssavirus Real Time RT-PCR for Rabies Diagnosis

Molecular assays are rapid, sensitive and specific, and have become central to diagnosing rabies. PCR based assays have been utilized for decades to confirm rabies diagnosis but have only recently been accepted by the OIE (World Organisation for Animal Health) as a primary method to detect rabies infection. Real-time RT-PCR assays provide real-time data, and are closed-tube systems, minimizing the risk of contamination during setup. DNA intercalating fluorochrome real-time RT-PCR assays do not require expensive probes, minimizing the cost per sample, and when the primers are designed in conserved regions, assays that are specific across virus genera rather than specific to just one virus species are possible. Here we describe a pan-lyssavirus SYBR real-time RT-PCR assay that detects lyssaviruses across the Lyssavirus genus, including the most divergent viruses IKOV, WCBV and LLEBV. In conjunction with dissociation curve analysis, this assay is sensitive and specific, with the advantage of detecting all lyssavirus species. The assay has been adopted in many diagnostic laboratories with quality assured environments, enabling robust, rapid, sensitive diagnosis of animal and human rabies cases.

This real-time RT-PCR using dsDNA intercalating dye is suitable to diagnose lyssavirus infections. The method begins with RNA extracted from rabies suspected ante-mortem or post-mortem samples, detailing master mix preparation, RNA addition, setup of the real-time machine and correct interpretation of results.

Molecular assays are rapid, sensitive and specific, and have become central to diagnosing rabies. PCR based assays have been utilized for decades to ...

Immunology and Infection

Field Postmortem Rabies Rapid Immunochromatographic Diagnostic Test for Resource-Limited Settings with Further Molecular Applications

Functional rabies surveillance systems are crucial to provide reliable data and increase the political commitment necessary for disease control. To date, animals suspected as rabies-positive must be submitted to a postmortem confirmation using classical or molecular laboratory methods. However, most endemic areas are in low- and middle-income countries where animal rabies diagnosis is restricted to central veterinary laboratories. Poor availability of surveillance infrastructure leads to serious disease underreporting from remote areas. Several diagnostic protocols requiring low technical expertise have been recently developed, providing opportunity to establish rabies diagnosis in decentralized laboratories. We present here a complete protocol for field postmortem diagnosis of animal rabies using a rapid immunochromatographic diagnostic test (RIDT), from brain biopsy sampling to the final interpretation. We complete the protocol by describing a further use of the device for molecular analysis and viral genotyping. RIDT easily detects rabies virus and other lyssaviruses in brain samples. The principle of such tests is simple: brain material is applied on a test strip where gold conjugated antibodies bind specifically to rabies antigens. The antigen-antibody complexes bind further to fixed antibodies on the test line, resulting in a clearly visible purple line. The virus is inactivated in the test strip, but viral RNA can be subsequently extracted. This allows the test strip, rather than the infectious brain sample, to be safely and easily sent to an equipped laboratory for confirmation and molecular typing. Based on a modification of the manufacturer&#39;s protocol, we found increased test sensitivity, reaching 98% compared to the gold standard reference method, the direct immunofluorescence antibody test. The advantages of the test are numerous: rapid, easy-to-use, low cost and no requirement for laboratory infrastructure, such as microscopy or cold-chain compliance. RIDTs represent a useful alternative for areas where reference diagnostic methods are not available.

We present a complete protocol for postmortem diagnosis of animal rabies under field conditions using a rapid immunochromatographic diagnostic test (RIDT), from brain biopsy sampling to final interpretation. We also describe further applications using the device for molecular analysis and viral genotyping.

Functional rabies surveillance systems are crucial to provide reliable data and increase the political commitment necessary for disease control. To date, ...

Using Colorimetric and RT-LAMP Analysis to Detect SARS-CoV-2 

Detecting SARS-CoV-2 Virus by Reverse Transcription-Loop-Mediated Isothermal Amplification

The severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) virus has dramatically impacted human health. It continues to be a threat to modern society because many people die as a result of infection. The disease is diagnosed using serologic and molecular tests, such as the gold standard real-time polymerase chain reaction (RT-PCR). The last has several disadvantages because it requires specialized infrastructure, costly equipment, and trained personnel. Here, we present a protocol outlining the steps required to detect the SARS-CoV-2 virus using reverse transcription-loop-mediated isothermal amplification (RT-LAMP) in human samples. The protocol includes instructions for designing primers in silico, preparing reagents, amplification, and visualization. Once standardized, this method can be easily implemented and adapted to any laboratory or point-of-care within 60 min at a low cost and using inexpensive equipment. It is adaptable to detecting different pathogens. Thus, it can potentially be used in the field and in health centers to carry out timely epidemiological surveillance.

Author Spotlight: Advancing Pathogen Diagnostics with Standardized LAMP

Here we provide a complete protocol to standardize and implement the method for detecting the SARS-CoV-2 virus in human samples by reverse transcription loop-mediated isothermal amplification (RT-LAMP). This method, done within 60 min, could be adapted to any laboratory or point-of-care at a low cost and using inexpensive equipment.

The severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) virus has dramatically impacted human health. It continues to be a threat to modern ...

Biology

Multiplex RT-qPCR Assay for Respiratory Virus Detection

Development of Multiplex Real-Time RT-qPCR Assays for the Detection of SARS-CoV-2, Influenza A/B, and MERS-CoV

The severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) that causes Coronavirus disease 2019 (COVID-19) is a serious threat to the general public&#39;s health. During influenza seasons, the spread of SARS-CoV-2 and other respiratory viruses may cause a population-wide burden of respiratory disease that is difficult to manage. For that, the respiratory viruses SARS-CoV-2, Influenza A, Influenza B, and Middle East respiratory syndrome (MERS-CoV) will need to be carefully watched over in the upcoming fall and winter seasons, particularly in the case of SARS-CoV-2, Influenza A, and Influenza B, which share similar epidemiological factors like susceptible populations, mode of transmission, and clinical syndromes. Without target-specific assays, it can be challenging to differentiate among cases of these viruses owing to their similarities. Accordingly, a sensitive and targeted multiplex assay that can easily differentiate between these viral targets will be useful for healthcare practitioners. In this study, we developed a real-time reverse transcriptase-PCR-based assay utilizing an in-house developed R3T one-step RT-qPCR kit for simultaneous detection of SARS-CoV-2, Influenza A, Influenza B, and SARS-CoV-2, MERS-CoV. With as few as 10 copies of their synthetic RNAs, we can successfully identify SARS-CoV-2, Influenza A, Influenza B, and MERS-CoV targets simultaneously with 100% specificity. This assay is found to be accurate, reliable, simple, sensitive, and specific. The developed method can be used as an optimized SARS-CoV-2, Influenza A, Influenza B, and SARS-CoV-2, MERS-CoV diagnostic assay in hospitals, medical centers, and diagnostic laboratories as well as for research purposes.

Author Spotlight: Advancements in Multiplex Detection of Respiratory Viruses 

We present two probe-based in-house one-step RT-qPCR kits for common respiratory viruses. The first assay is for SARS-CoV-2 (N), Influenza A (H1N1 and H3N2), and Influenza B. The second is for SARS-Cov-2 (N) and MERS (UpE and ORF1a). These assays can be successfully implemented in any specialized laboratory.

The severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) that causes Coronavirus disease 2019 (COVID-19) is a serious threat to the general ...

Postmortem Diagnosis of Rabies in Animals by the Updated, Multiplexed LN34 Real-Time Reverse Transcription-Polymerase Chain Reaction Assay

Rabies is a fatal zoonotic disease caused by Lyssavirus rabies (RABV) and related negative strand RNA viruses from the Lyssavirus genus (family Rhabdoviridae). The LN34 assay targets the highly conserved leader region and nucleoprotein gene of the lyssavirus genome and utilizes degenerate primers and a TaqMan probe containing locked nucleotides to detect RNA across the diverse Lyssavirus genus. A negative finding for rabies should be made only if a full cross-section of the brain stem and three lobes of the cerebellum are examined; however, identification of lyssavirus RNA in any tissue is diagnostic of rabies infection. Tissue is collected and homogenized in TRIzol reagent, which also inactivates the virus. RNA extraction is performed using a spin column-based commercial extraction kit. Master mixes are prepared in a clean space and aliquoted into a 96 well plate before adding sample RNA. In clinical settings, each sample is tested by real-time RT-PCR for the presence of lyssavirus RNA in triplicate and singly for host &#946; -actin mRNA. Positive and negative controls are included at extraction and real-time RT-PCR steps of the protocol. Data analysis involves manual adjustment of the thresholds to standardize Ct values across instrument runs. Positive results are determined by the presence of typical amplification in the pan-lyssavirus assay (Ct &#8804; 35). Negative results are determined by the absence of typical amplification in the pan-lyssavirus assay and detection of host &#946;-actin mRNA (Ct &#8804; 33). Observation of values outside of these ranges or failure of the assay controls can invalidate the run or result in inconclusive results for a specimen. The protocol should be closely followed to ensure high assay sensitivity and specificity. Procedural modifications can affect assay performance and lead to false positive, false negative, or uninterpretable results.

This protocol demonstrates the pan-lyssavirus LN34 real-time reverse transcription-polymerase chain reaction (RT-PCR) assay from tissue collection to result interpretation, including updates to primer sequences and formulations to improve assay performance for some non-rabies lyssaviruses and lagomorphs. We also demonstrate the assay setup for a single-well LN34 multiplexed (LN34M) format.

Rabies is a fatal zoonotic disease caused by Lyssavirus rabies (RABV) and related negative strand RNA viruses from the Lyssavirus genus (family ...

Real-Time Quantitative Reverse Transcription PCR for Diagnosing Viral Infections

Source: Marston, D. A. et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/59709">Pan-lyssavirus Real Time RT-PCR for Rabies Diagnosis.</a> J. Vis. Exp.&#160;(2019).
This video demonstrates a quantitative method for detecting viral infection using real-time reverse transcription PCR. The reverse transcription step converts the RNA to cDNA, which is then amplified via PCR cycles. The presence of viral RNA in the sample is confirmed by analyzing the amplification and dissociation curves of DNA obtained from the PCR.

Source: Marston, D. A. et al. Pan-lyssavirus Real Time RT-PCR for Rabies Diagnosis. J. Vis. Exp.&#160;(2019).
This video demonstrates a quantitative ...

JoVE Encyclopedia of Experiments

Biological Techniques

PCR Techniques

Nested-PCR to Detect a Specific Viral Genomic Sequence

Source: Feng, Y., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/59428">Evaluation of a Universal Nested Reverse Transcription Polymerase Chain Reaction for the Detection of Lyssaviruses.</a> J. Vis. Exp. (2019).
This video describes nested polymerase chain reaction, a technique that consists of two sequential PCR amplification processes using two primer sets. The first set of primers is intended to anneal to sequences upstream of the second set, resulting in selective amplification of specific gene sequences. This PCR is more sensitive and specific than a normal PCR and is widely used as a detection technique for various diseases.

Source: Feng, Y., et al. Evaluation of a Universal Nested Reverse Transcription Polymerase Chain Reaction for the Detection of Lyssaviruses. J. Vis. Exp. ...

Evaluation of a Universal Nested Reverse Transcription Polymerase Chain Reaction for the Detection of Lyssaviruses

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