The authors wish to thank Miss Emma Wise and Miss Megan Golding for assistance in completing the experiments. The development of this protocol was financially supported by the UK Department for Environment, Food and Rural Affairs (Defra), Scottish Government and Welsh Government by grants [SV3500, and SE0431] and by European Virus Archive global (EVAg) project that has received funding from the European Union&#8217;s Horizon 2020 research and innovation programme under grant agreement No 653316.

Samples from diagnostic material received at APHA after natural infection, or obtained by inoculating mice using protocols assessed by the APHA ethics and statistical committee under UK Home Office regulations under licence 70/7394.
1. Quantification of RNA Using a Micro-volume Spectrophotometer
<ol>
	<li>Ensure the settings of the spectrophotometer are set to RNA.</li>
	<li>Use 1-2 &#181;L of molecular grade water to initialize the machine and set a baseline.</li>
	<li>Use 1-2 &#181;L of each test RNA sample to assess the RNA quantity.</li>
	<li>Save the readings and document.</li>
	<li>Adjust the RNA to 1 &#181;g/&#181;L, if required. 
	NOTE: RNA must be kept on ice (or in a cool block) at all times. If RNA is obtained using a column, or bead-based method the RNA is usually less than 1 &#181;g/&#181;L. In this situation use the RNA neat.</li>
</ol>
2. Preparation of RNA Dilution Series to Determine End Point Sensitivity
<ol>
	<li>Make a 10-fold serial dilution of the RNA.
	<ol>
		<li>Label tubes with the dilution series (e.g., 10-1, 10-2, etc.) and the RNA details.</li>
		<li>Add 45 &#181;L of molecular grade water to each tube.</li>
		<li>Add 5 &#181;L of the RNA (previously diluted to 1 &#181;g/&#181;L) and mix well.</li>
		<li>Dispose of pipette tip in appropriate disinfectant and replace with a fresh tip.</li>
		<li>Take 5 &#181;L from the 10-1 and add to 10-2 tube and mix well.</li>
		<li>Repeat 2.1.3-2.1.5 with the remaining dilutions. 
		NOTE: RNA must be kept on ice (or in a cool block) at all times.</li>
	</ol>
	</li>
</ol>
3. Preparation of Real-time RT-PCR Reactions
<ol>
	<li>Using a spreadsheet, plan the plate layout according to the number of test samples and control samples, for both the lyssavirus and &#223;-actin assays. 
	NOTE: If 4 samples are to be tested in duplicate with a positive and negative control, this equates to 10 reactions for both assays.</li>
	<li>In a &#8216;clean room&#8217; or area separate from the RNA template, wipe down the surfaces with an appropriate disinfectant prior to use or prepare a PCR workstation (if using). To prepare the workstation, wipe the cabinet surface with an appropriate disinfect and place the items required into the workstation and close the doors. Switch on the UV light for 10 minutes</li>
	<li>Remove regents and primers from the freezer and thaw (reagents listed in Table 1 and primers in Table 2). 
	NOTE: The enzyme mix is stored in glycerol so does not require thawing and must be kept on ice (or in a cool block) at all times. All other reagents can be thawed at room temperature.</li>
	<li>Once thawed, mix the reagents and centrifuge briefly to collect liquid. 
	NOTE: Do not vortex the enzyme mix, just centrifuge briefly.</li>
	<li>Prepare separate master mixes for lyssavirus and &#223;-actin. For each reaction, add 7.55 &#181;L of molecular grade water, 10 &#181;L of 2x Universal SYBR green reaction mix, 0.6 &#181;L of forward primer, 0.6 &#181;L of reverse primer, 0.25 &#181;L of iTaq RT enzyme mix.
	<ol>
		<li>Use a spreadsheet to calculate correct volumes to avoid errors in manual calculating. Ensure enough master-mix is prepared to compensate for pipetting error. Therefore if 10 reactions are required (see NOTE in 3.1), prepare 12 reactions</li>
		<li>Prepare the master-mix on ice (or in a cool block) and remain on ice until placed into the real-time machine.</li>
	</ol>
	</li>
	<li>Mix the prepared master-mixes, centrifuge briefly and dispense 19 &#181;L into the relevant wells of strip tubes or a 96 well plate compatible with the real-time machine in use. 
	NOTE: Minimize the production of bubbles in the wells whilst pipetting.</li>
	<li>In a separate room, or in a UV cabinet prepared as described in 3.2, carefully add 1 &#181;L of the RNA previously adjusted to 1 &#181;g/&#181;L (see step 1.5) below the surface of the appropriate master-mix well and mix gently. Discard pipette tip into disinfectant directly after use (under the surface).
	<ol>
		<li>Add the controls after the test samples, with the positive control added next and the no template control (NTC &#8211; molecular grade water) added last. The amount of RNA used can be altered depending on the sample type, and RNA extraction used. The amount used must be validated to ensure the reaction is optimized.</li>
	</ol>
	</li>
	<li>Seal plate using strip lids or sealer taking care to ensure all the lids are firmly closed and labelled sufficiently to orientate the samples. Label the edge of the plate/strip tubes.</li>
	<li>Spin down the samples using a centrifuge to collect all the liquid at the bottom of the wells.</li>
	<li>Transfer plate to the real-time PCR machine, open the door and place in the holder ensuring the correct location/orientation of the samples according to the plate layout. 
	NOTE: If tube strips are used, ensure the holder is in place.</li>
	<li>Open up the real-time PCR machine program and choose the option for SYBR real-time experiment with dissociation curve. Program the real-time PCR machine using the thermal cycling conditions specified in Table 3, including the data collection points.</li>
	<li>Select SYBR as the fluorescent dye and select unknown as the sample type and insert a name into the correct sample name box. 
	NOTE: Differentiate between the replicates and also between the lyssavirus and &#223;-actin wells.</li>
	<li>Choose a file location to save the experimental data, ensure the lamp will be switched off at the end of the run, then start the run. 
	NOTE: As the first step is an RT stage, no data is collected during this time, therefore, if the lamp requires a warm up period this can occur during the RT stage. The real-time machine and software will display the amplification curves in real-time, while the melting curve will be generated at the end of the cycle.</li>
</ol>
4. Data Analysis
<ol>
	<li>Once the run has been completed perform the data analysis as follows.
	<ol>
		<li>First analyze the amplification plot results of the test samples alongside the control samples. Positive samples display exponential ramps, usually followed by plateau and a Ct value. Negative samples display flat amplification plots with no Ct values (Figure 1A). The Ct value is automatically calculated by the software, although this should be checked and manually altered if required.</li>
		<li>Second, analyze the dissociation curve results of the test samples alongside the control samples. A positive sample will have a melting temperature (Tm) 77 &#8211; 80 &#176;C, and overlap with the positive control Figure 1B).</li>
		<li>Obtain the overall diagnostic result by ensuring the controls are valid. Use Table 4 to interpret the results in relation to the internal &#223;-actin control. If the positive control samples are negative and/or the negative samples are positive, the run should be disregarded.</li>
		<li>Record the Ct and Tm values obtained for the control RNA in a &#8216;control card&#8217; to enable trend analysis and help identify drifts in the assay sensitivity.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=OIE.">OIE.</a> OIE Terrestrial Manual 2018. , 8-14 (2018).</li><li>Panning, M., 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=Comparative+analysis+of+rabies+virus+reverse+transcription-PCR+and+virus+isolation+using+samples+from+a+patient+infected+with+rabies+virus.">Comparative analysis of rabies virus reverse transcription-PCR and virus isolation using samples from a patient infected with rabies virus.</a> Journal of Clinical Microbiology. 48 (8), 2960-2962 (2010).</li><li>Wakeley, P. 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=Development+of+a+real-time,+TaqMan+reverse+transcription-PCR+assay+for+detection+and+differentiation+of+lyssavirus+genotypes+1,+5,+and+6.">Development of a real-time, TaqMan reverse transcription-PCR assay for detection and differentiation of lyssavirus genotypes 1, 5, and 6.</a> Journal of Clinical Microbiology. 43 (6), 2786-2792 (2005).</li><li>Wang, L., 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=A+SYBR-green+I+quantitative+real-time+reverse+transcription-PCR+assay+for+rabies+viruses+with+different+virulence.">A SYBR-green I quantitative real-time reverse transcription-PCR assay for rabies viruses with different virulence.</a> Virologica Sinica. 29 (2), 131-132 (2014).</li><li>Wadhwa, A., 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=A+Pan-Lyssavirus+Taqman+Real-Time+RT-PCR+Assay+for+the+Detection+of+Highly+Variable+Rabies+virus+and+Other+Lyssaviruses.">A Pan-Lyssavirus Taqman Real-Time RT-PCR Assay for the Detection of Highly Variable Rabies virus and Other Lyssaviruses.</a> PLOS Neglected Tropical Diseases. 11 (1), e0005258 (2017).</li><li>Nadin-Davis, S. A., Sheen, M., Wandeler, A. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+real-time+reverse+transcriptase+polymerase+chain+reaction+methods+for+human+rabies+diagnosis.">Development of real-time reverse transcriptase polymerase chain reaction methods for human rabies diagnosis.</a> Journal of Medical Virology. 81 (8), 1484-1497 (2009).</li><li>Hoffmann, B., 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=Improved+safety+for+molecular+diagnosis+of+classical+rabies+viruses+by+use+of+a+TaqMan+real-time+reverse+transcription-PCR+&#34;double+check&#34;+strategy.">Improved safety for molecular diagnosis of classical rabies viruses by use of a TaqMan real-time reverse transcription-PCR &#34;double check&#34; strategy.</a> Journal of Clinical Microbiology. 48 (11), 3970-3978 (2010).</li><li>Faye, M., 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=Development+and+validation+of+sensitive+real-time+RT-PCR+assay+for+broad+detection+of+rabies+virus.">Development and validation of sensitive real-time RT-PCR assay for broad detection of rabies virus.</a> Journal of Virological Methods. 243, 120-130 (2017).</li><li>Dacheux, L., 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=Dual+Combined+Real-Time+Reverse+Transcription+Polymerase+Chain+Reaction+Assay+for+the+Diagnosis+of+Lyssavirus+Infection.">Dual Combined Real-Time Reverse Transcription Polymerase Chain Reaction Assay for the Diagnosis of Lyssavirus Infection.</a> PLOS Neglected Tropical Diseases. 10 (7), e0004812 (2016).</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=Rabies.">Rabies.</a> Nature Reviews Disease Primers. 3, 17091 (2017).</li><li>Marston, D. A., 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=Ikoma+lyssavirus,+highly+divergent+novel+lyssavirus+in+an+african+civet.">Ikoma lyssavirus, highly divergent novel lyssavirus in an african civet.</a> Emerging Infectious Diseases. 18 (4), 664-667 (2012).</li><li>. Virus Taxonomy: 2018 Release: Email ratification October 2018 (MSL #33) Available from: <a target="_blank" href="https://talk.ictvonline.org/taxonomy/">https://talk.ictvonline.org/taxonomy/</a> (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 Disease. 24 (4), 782-785 (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>Hayman, D. 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=A+universal+real-time+assay+for+the+detection+of+Lyssaviruses.">A universal real-time assay for the detection of Lyssaviruses.</a> Journal of Virological Methods. 177 (1), 87-93 (2011).</li><li>Calisher, C. H., 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=Antigenic+relationships+among+rhabdoviruses+from+vertebrates+and+hematophagous+arthropods.">Antigenic relationships among rhabdoviruses from vertebrates and hematophagous arthropods.</a> Intervirology. 30 (5), 241-257 (1989).</li><li>Aznar-Lopez, 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=Detection+of+rhabdovirus+viral+RNA+in+oropharyngeal+swabs+and+ectoparasites+of+Spanish+bats.">Detection of rhabdovirus viral RNA in oropharyngeal swabs and ectoparasites of Spanish bats.</a> Journal of General Virology. 94 (1), 69-75 (2013).</li></ol>

The authors have nothing to disclose.

The pan-lyssavirus real-time RT-PCR assay described is a closed-tube, one-step assay which detects lyssaviruses across all three phylogroups. The assay has been validated for both animal and human rabies diagnosis, including post-mortem brain tissue (optimally brainstem), and ante-mortem samples such as skin biopsy, serially collected saliva, or cerebral spinal fluid (CSF). The primers utilized in this assay were first designed and utilized for a probe-based assay to differentiate between RABV, EBLV-1 and EBLV-23, which has been used in many OIE rabies laboratories and performs consistently in the EURL proficiency schemes. Subsequently the &#8216;pan-lyssavirus&#8217; nature of these primers has been confirmed using a 2-step real-time assay15. The assay described here has utilized the primers to further optimize the RT-PCR in a one-step SYBR assay enabling a closed-tube, rapid system. Furthermore, training in rabies endemic countries using this assay has confirmed suitability to implement in any laboratory with basic PPE and quality systems to reduce cross contamination and trace samples, facilities to store the reagents and a real-time machine with SYBR detection. The assay is extremely robust and has 100% correlation with the FAT, with improved sensitivity for decomposed samples3. One of the main advantages for a DNA intercalating dye based assay in comparison to a probe based assay is the relative cost. An additional benefit is that the assay only utilizes two primers, therefore there is less risk of failed detection due to sequence divergence, which has been a weakness in previously published probe-based assays. Indeed, representatives of all lyssavirus species (apart from TWBLV and KBLV) are detected using this assay, and sequence analysis of TWBLV and KBLV across the primer sites reveals no significant divergence strongly suggesting that they will also be detected using this method. The range of limit of detection observed across the viruses analyzed, can be considered to be due to two main factors. The first is that the RNA was isolated from clinical brain material, therefore the amount of genome copies in each undiluted sample is not directly comparable. The RNA was &#8216;normalized&#8217; by adjusting the total RNA to 1 &#181;g/&#181;L; however, the proportion of viral genome RNA within that sample will vary. The second is the diversity of lyssavirus sequences, despite the primer sites being located in conserved regions, there remain positions of variation. Therefore, it is not surprising that the majority of lyssaviruses with lower sensitivity for the assay are phylogroup II and III viruses. The dissociation curve analysis represents an essential parameter, minimizing a false positive result which could otherwise occur due to the formation of primer dimers, or amplification of a non-specific region in the host genome. In reality, this is a rare occurrence and the dissociation curve analysis is equivalent to running an agarose gel to visualize correctly sized conventional RT-PCR amplicons. The range of acceptable Tm values has been provided (77-80 &#176;C), based on the data collected in our laboratory. It is strongly recommended that individual laboratories collate in-house data to ensure the range is transferrable and amend accordingly. Interpretation of the results from both the amplification and dissociation plots, alongside the positive and negative controls and the &#223;-actin results, enables robust and reproducible diagnostic outcomes.
Outside the scope of this protocol is the RNA extraction method used to obtain high quality RNA. All RNA analyzed in this protocol was prepared using TRIzol; however, there are many suitable guanidium-based extractions RNA extraction protocols available, including column and bead-based extraction kits. Handling of lyssavirus positive (or suspected positive) samples must be within licenced biocontainment facilities approved within country. However, the total RNA extracted is non-infectious, therefore handled within low containment laboratories. Depending on the extraction method used, the requirement to quantify and dilute the RNA would need to be assessed. For phenol-based extractions, including TRIzol, this step is required and prevents inhibition of the assay from contaminating gDNA; however, column and bead-based extractions (particularly those with a DNA depletion stage) do not require dilution prior to testing.
Throughout the protocol, it is essential that care and diligence are used to prevent cross-contamination and accurate addition of sample to the correct wells. A spreadsheet with the reagent calculations and plate layout is available for download as a supplemental file. Good laboratory practice, including clean work surfaces, regular changes of gloves, use of barrier tips and different rooms/UV cabinets to separate each stage will minimize the chance of contamination. To ensure the test is performing with expected parameters, positive and negative controls must be included and all test samples run in duplicate (or triplicate).
The inclusion of controls is an essential feature of any PCR, particularly for diagnostics. Positive control RNA was prepared from CVS (challenge virus standard) infected mouse brains in batches and validated and calibrated to ensure consistency between batches. The control RNA was quantified and diluted to 1 &#181;g/&#181;L. RNA for which a positive result was obtained in a serial dilution down to at least 10-4 (equal to 100 pg/&#181;L) was considered fit for purpose. The positive control RNA was stored at -80 &#176;C in 10-1 aliquots. When required, an aliquot was diluted 1:100 to provide a working stock at 1 ng/&#181;L and stored at -80 &#176;C in 5 &#181;L single use aliquots. The diluted positive control RNA was used to represent low level positive samples, and to ensure that any reduction in sensitivity of the assay was detected. A &#8216;control card&#8217; was kept for each control to monitor the Ct values and identify trends (Table 5). Table 5 demonstrated good inter-run comparability for the CVS positive control sample Ct and Tm values across multiple days and operators, providing reassurance that the assay is robust and reproducible. Results that deviate from these measurements should be investigated and test samples repeated if necessary. Molecular grade water was included in every run as an NTC to confirm the reagents were free from contamination with lyssavirus RNA and confirm a negative sample. Furthermore, to ensure RNA extraction efficacy, &#223;-actin was tested alongside the test samples in a separate tube. The lyssavirus positive control RNA was also used for the &#223;-actin positive control. Other endogenous genes or heterologous internal control systems can be utilized. Use of these controls ensured that all steps were analyzed under the same conditions as the test samples. Occasionally, if the sample was highly degraded or did not contain sufficient host RNA (such as saliva or CSF), the endogenous gene PCR can fail. In this instance, where the lyssavirus real-time RT-PCR result was positive, confirmation on an independent RNA extraction - to rule out contamination during RNA extraction on the original test or by a secondary test (either molecular, such as conventional RT-PCR), or FAT. Use of Table 4 during analysis of a diagnostic sample ensured the correct interpretation.
Regardless of the molecular assay used to confirm rabies infection, follow up investigations using Sanger sequencing to determine the lyssavirus species and classical techniques in virology such as FAT or virus isolation should also be undertaken to allow for further virus characterisation and support the notification of positive cases to OIE and WHO.

Diagnosis of rabies using molecular methodologies was accepted by the OIE in 20181, recognizing the advantages of these techniques in confirming rabies cases, particularly in situations when the samples are sub-optimal, or for ante-mortem diagnosis, as there is no requirement for live virus or fresh samples. PCR assays for lyssaviruses require a reverse transcription (RT) before PCR can commence as the genome is RNA. RT-PCR assays that detect the 3&#8217; proximal region of the genome are considered the most sensitive, as transcriptional gradients occur during lyssavirus replication. Commonly used RT-PCR assays can be divided broadly into two categories, end-point (or gel-based) and real-time. Both approaches are sensitive and specific; however, the real-time assay has some additional benefits such as obtaining results in &#8216;real-time&#8217; and being performed in an entirely closed tube system, thereby reducing the potential for operator contamination. There are two main approaches to detect lyssavirus-specific amplicons obtained using real-time assays. The first utilizes hydrolysis probes (such as TaqMan probes) that contain a fluorophore and a quencher. When the probe binds to the target region during amplification, the exonuclease activity of the polymerase results in dissociation of fluorophore and quencher, enabling the resulting fluorescence to be measured. The second utilizes a DNA intercalating dye (fluorochrome such as SYBR Green) that binds to double stranded DNA during amplification. The bound fluorochromes emit fluorescence that is detected at each cycle, allowing real time detection and quantification of the product. Due to the non-specific nature of binding to any dsDNA, a dissociation curve analysis is undertaken to confirm the specificity of the reaction. Real-time RT-PCRs are rapid due to the small amplicon sizes, typically less than 200 bp in length; however, identifying suitable regions to design primers and probes in conserved regions, can prove challenging, therefore removing the requirement for a probe is a distinct advantage.
A number of real-time RT-PCRs have been designed for specifically detecting individual strains or lineages of RABV2 and also to detect lyssaviruses across the genus3,4,5,6,7,8,9. All assays will have a limit of detection dependent on how conserved the primer (and if necessary, the probe) sequences are across the genus. Indeed, emerging or novel virus strains may render the highly specific probe-based assays ineffective. The choice of the real-time detection (dye vs. probe) will depend on the intended application. For a laboratory conducting surveillance on locally sourced brain material and expecting high numbers of negative samples, the use of the cheaper intercalating dye is a sensible choice. The SYBR Green approach would also be optimal when conducting scanning surveillance where the presence of novel or divergent lyssaviruses would remain undetected by more restricted probe-based assays.
All members of the genus Lyssavirus cause the disease rabies, which is fatal once symptoms appear. The vast majority of human and animal rabies cases are due to rabies virus (RABV), the dominant reservoir for which is the domestic dog10. Bats are important host reservoirs for lyssaviruses and all but two lyssavirus species characterized have been identified directly in bats &#8212; Ikoma lyssavirus (IKOV) and Mokola virus (MOKV) &#8212; and of these two, IKOV has been speculated to have a bat host reservoir11. In addition to the 16 recognized lyssavirus species12, there are two lyssaviruses that have been recently described: Taiwan bat lyssavirus (TWBLV)13 and Kotalahti bat lyssavirus (KBLV)14. Lyssaviruses can be genetically divided into three phylogroups, with the majority of lyssaviruses, including RABV, belonging to phylogroup I. However, the most divergent lyssaviruses belong to phylogroup III and are unlikely to be detected by RT-PCRs designed to target RABV or phylogroup I virus sequences.
The assay described here utilizes the real-time primer pair JW12-N165, first described in 20053. The primers were designed to be pan-lyssavirus albeit the original application was as a TaqMan assay with probes to differentiate lyssavirus species. Subsequent confirmation that the primer pair was pan-lyssavirus in specificity was achieved utilizing a 2-step SYBR real-time assay on all lyssavirus species available including WCBV15. The primer pair is described here in a one-step RT-PCR real-time assay utilizing an intercalating fluorochrome, validated using representatives from all 16 recognized lyssavirus species. This one-step real-time assay is a rapid, sensitive, lyssavirus-specific assay and demonstrates that the robustness of the primer set to identify even highly divergent lyssavirus species.

<table><tbody><tr><td>Art Barrier pipette tips (various sizes)</td><td>Thermofisher</td><td>various</td><td></td></tr><tr><td>Centrifuge</td><td>Beckman</td><td>Allegra 21R</td><td>Rotor capable of holding 96 well plates required. Step 3.8.</td></tr><tr><td>Centrifuge (mico)</td><td>Sigma</td><td></td><td></td></tr><tr><td>Finnpipettes (to dispense 0.5-1,000 &amp;micro;L)</td><td>Thermofisher</td><td>various</td><td></td></tr><tr><td>iTaq Universal SYBR Green One-Step RT-PCR kit</td><td>Bio-Rad</td><td>172-5150</td><td>Equivalent kits can be used if validated</td></tr><tr><td>MX3000P or MX3005P real-tme PCR system</td><td>Stratagene</td><td>N/A</td><td>Eqivalent machines can be used if validated</td></tr><tr><td>MicroAmp reaction plate base</td><td>Any suitable</td><td></td><td>Used to hold tube strip and plates securely.</td></tr><tr><td>Optically clear flat clear strips (8)</td><td>ABgene</td><td>AB-0866</td><td></td></tr><tr><td>Perfect fit frame (if using tube strips)</td><td>Stratagene</td><td>N/A</td><td>Specific to machine</td></tr><tr><td>Primers: for primer details see Table 2.</td><td></td><td></td><td>Ordered at 0.05 &amp;micro;mole scale HPLC purified.</td></tr><tr><td>Thermo-Fast 96 well plares, non skirted</td><td>ABgene</td><td>AB-600</td><td></td></tr><tr><td>Thermo-Fast strips (8) Thermo-tubes</td><td>ABgene</td><td>AB-0452</td><td></td></tr><tr><td>Vortex machine / Whirlimixer</td><td>Fisons Scientific equipment</td><td>SGP-202-010J</td><td></td></tr><tr><td>Unless stated, alternative equipment can be used</td><td></td><td></td><td></td></tr></tbody></table>

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.

Watch this Scientific Journal Video about Pan-lyssavirus Real Time RT-PCR for Rabies Diagnosis at JoVE.com

Pan-lyssavirus Real Time RT-PCR for Rabies Diagnosis

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 ...

pan-lyssavirus-real-time-rt-pcr-for-rabies-diagnosis

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Immunology and Infection

21.5K Views.  Animal and Plant Health Agency. Deze real-time RT-PCR is geschikt om snel rabiës te diagnosticeren in ante-mortem- en postmortemmonsters. De Pan-Lyssavirus primers zijn geoptimaliseerd om alle leden van het Lyssavirus geslacht te identificeren. Dit is een snelle, gevoelige en gesloten buistest die virussen uit het hele geslacht Lyssavirus detecteert, waaronder zeer uiteenlopende soorten uit klinische specimens.Het OIE heeft onlangs aanvaard moleculaire tests om rabiës infectie te melden. Dit is vooral belangrijk vo...

Video: Pan-lyssavirus Real Time RT-PCR for Rabies Diagnosis

Detection of MicroRNAs in Microglia by Real-time PCR in Normal CNS and During Neuroinflammation

Microglia are cells of the myeloid lineage that reside in the central nervous system (CNS)1. These cells play an important role in pathologies of many diseases associated with neuroinflammation such as multiple sclerosis (MS)2. Microglia in a normal CNS express macrophage marker CD11b and exhibit a resting phenotype by expressing low levels of activation markers such as CD45. During pathological events in the CNS, microglia become activated as determined by upregulation of CD45 and other markers3. The factors that affect microglia phenotype and functions in the CNS are not well studied. MicroRNAs (miRNAs) are a growing family of conserved molecules (~22 nucleotides long) that are involved in many normal physiological processes such as cell growth and differentiation4 and pathologies such as inflammation5. MiRNAs downregulate the expression of certain target genes by binding complementary sequences of their mRNAs and play an important role in the activation of innate immune cells including macrophages6 and microglia7. In order to investigate miRNA-mediated pathways that define the microglial phenotype, biological function, and to distinguish microglia from other types of macrophages, it is important to quantitatively assess the expression of particular microRNAs in distinct subsets of CNS-resident microglia. Common methods for measuring the expression of miRNAs in the CNS include quantitative PCR from whole neuronal tissue and in situ hybridization. However, quantitative PCR from whole tissue homogenate does not allow the assessment of the expression of miRNA in microglia, which represent only 5-15% of the cells of neuronal tissue. Hybridization in situ allows the assessment of the expression of microRNA in specific cell types in the tissue sections, but this method is not entirely quantitative. In this report we describe a quantitative and sensitive method for the detection of miRNA by real-time PCR in microglia isolated from normal CNS or during neuroinflammation using experimental autoimmune encephalomyelitis (EAE), a mouse model for MS. The described method will be useful to measure the level of expression of microRNAs in microglia in normal CNS or during neuroinflammation associated with various pathologies including MS, stroke, traumatic injury, Alzheimer's disease and brain tumors.

Microglia are resident macrophages that provide the first line of defense and immune surveillance of the central nervous system. MicroRNAs are regulatory molecules that play an important role in many physiological processes including activation and differentiation of macrophages. In this article, we describe the method for measurement of microRNAs in microglia.

Microglia are cells of the myeloid lineage that reside in the central nervous system (CNS)1. These cells play an important role in pathologies of many ...

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.

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 ...

Genetics

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

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, ...

Immunohistochemistry Test for the Lyssavirus Antigen Detection from Formalin-Fixed Tissues

One of the primary diagnostic modalities for rabies is the detection of viral ribonucleoprotein (RNP) complex (antigen) in the infected tissue samples. While the direct fluorescent antibody (DFA) test or the direct rapid immunohistochemical test (DRIT) are most commonly utilized for the antigen detection, both tests require fresh and/or frozen tissues for impressions on slides prior to the antigen detection using antibodies. If samples are collected and fixed in formalin, neither test is optimal for the antigen detection, however, testing can be performed by conventional immunohistochemistry (IHC) after embedding in paraffin blocks and sectioning. With this IHC method, tissues are stained with anti-rabies antibodies, sections are deparaffinized, antigen retrieved by partial proteolysis or other methods, and incubated with primary and secondary antibodies. Antigens are stained using horseradish peroxidase / amino ethyl carbazole and counterstained with hematoxylin for the visualization using a light microscope. In addition to the specific antigen detection, formalin fixation offers other advantages like the determination of histological changes, relaxed conditions for specimen storage and transport (under ambient temperatures), ability to test retrospective cases and improved biological safety through the inactivation of infectious agents.

Here, we present an immunohistochemistry test protocol for the detection of rabies virus antigen as an alternative diagnostic test for formalin-fixed tissues.

One of the primary diagnostic modalities for rabies is the detection of viral ribonucleoprotein (RNP) complex (antigen) in the infected tissue samples. ...

Quantitation of Rabies Virus in Various Bovine Brain Structures

Bovine paralytic rabies (BPR) is a form of viral encephalitis that is of substantial economic importance throughout Latin America, where it poses a major zoonotic risk. Here, our objective was to utilize a laboratory protocol to determine the relative copy number of the rabies virus (RABV) genome in different bovine brain anatomical structures using quantitative real-time reverse transcription polymerase chain reaction (qRT-PCR). qRT-PCR quantifies the specific number of gene copies present in a sample based on fluorescence emitted after amplification that is directly proportional to the amount of target nucleic acid present in the sample. This method is advantageous owing to its short duration, reduced risk of contamination, and potential to detect viral nucleic acids in different samples more easily compared to other techniques. The brains of six rabid animals were divided into six anatomical structures, namely the Ammon&#8217;s horn, cerebellum, cortex, medulla, pons, and thalamus. All brains were identified as positive for RABV antigens based on a direct immunofluorescence test. The same anatomical structures from the brains of four RABV-negative bovines were also assessed. RNA was extracted from each structure and used for qRT-PCR. An assay was performed to determine the copy numbers of RABV genes using an in vitro transcribed nucleoprotein gene. The standard curve used to quantify viral RNA exhibited an efficiency of 100% and linearity of 0.99. Analysis revealed that the cortex, medulla, and thalamus were the ideal CNS portions for use in RABV detection, based on the observation that these structures possessed the highest levels of RABV. The test specificity was 100%. All samples were positive, no false positives were detected. This method can be used to detect RABV in samples that contain low levels of RABV during diagnosis of BPR.

This protocol presents a qRT-PCR-based approach for determining the rabies virus nucleoprotein (N) gene copy number within various bovine brain anatomical structures using in vitro transcription.

Bovine paralytic rabies (BPR) is a form of viral encephalitis that is of substantial economic importance throughout Latin America, where it poses a major ...

A Nanopore-Based Genomic Approach to Control Rabies in Low-Income Countries

Whole Genome Sequencing for Rapid Characterization of Rabies Virus Using Nanopore Technology

Genomic data can be used to track the transmission and geographic spread of infectious diseases. However, the sequencing capacity required for genomic surveillance remains limited in many low- and middle-income countries (LMICs), where dog-mediated rabies and/or rabies transmitted by wildlife such as vampire bats pose major public health and economic concerns. We present here a rapid and affordable sample-to-sequence-to-interpretation workflow using nanopore technology. Protocols for sample collection and the diagnosis of rabies are briefly described, followed by details of the optimized whole genome sequencing workflow, including primer design and optimization for multiplex polymerase chain reaction (PCR), a modified, low-cost sequencing library preparation, sequencing with live and offline base calling, genetic lineage designation, and phylogenetic analysis. Implementation of the workflow is demonstrated, and critical steps are highlighted for local deployment, such as pipeline validation, primer optimization, inclusion of negative controls, and the use of publicly available data and genomic tools (GLUE, MADDOG) for classification and placement within regional and global phylogenies. The turnaround time for the workflow is 2-3 days, and the cost ranges from $25 per sample for a 96 sample run to $80 per sample for a 12 sample run. We conclude that setting up rabies virus genomic surveillance in LMICs is feasible and can support progress toward the global goal of zero dog-mediated human rabies deaths by 2030, as well as enhanced monitoring of wildlife rabies spread. Moreover, the platform can be adapted for other pathogens, helping to build a versatile genomic capacity that contributes to epidemic and pandemic preparedness.

Author Spotlight: A Cost-Effective Genomic Workflow for Advancing Rabies Control in Resource-Limited Settings 

Here, we present a rapid and cost-effective workflow for characterizing rabies virus (RABV) genomes using nanopore technology. The workflow is intended to support genomics-informed surveillance at a local level, providing information on circulating RABV lineages and their placement within regional phylogenies to guide rabies control measures.

Genomic data can be used to track the transmission and geographic spread of infectious diseases. However, the sequencing capacity required for genomic ...

Biology

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/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 ...

Pan-lyssavirus Real Time RT-PCR for Rabies Diagnosis

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