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 0.5-1000 &#181;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 &#181;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

Research

JoVE Journal

Immunology and Infection

21.3K Views.  Animal and Plant Health Agency. 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.

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 ...

Real-time Live Imaging of T-cell Signaling Complex Formation

Protection against infectious diseases is mediated by the immune system 1,2. T lymphocytes are the master coordinators of the immune system, regulating the activation and responses of multiple immune cells 3,4. T-cell activation is dependent on the recognition of specific antigens displayed by antigen presenting cells (APCs). The T-cell antigen receptor (TCR) is specific to each T-cell clone and determines antigen specificity 5. The binding of the TCR to the antigen induces the phosphorylation of components of the TCR complex. In order to promote T-cell activation, this signal must be transduced from the membrane to the cytoplasm and the nucleus, initiating various crucial responses such as recruitment of signaling proteins to the TCR;APC site (the immune synapse), their molecular activation, cytoskeletal rearrangement, elevation of intracellular calcium concentration, and changes in gene expression 6,7. The correct initiation and termination of activating signals is crucial for appropriate T-cell responses. The activity of signaling proteins is dependent on the formation and termination of protein-protein interactions, post translational modifications such as protein phosphorylation, formation of protein complexes, protein ubiquitylation and the recruitment of proteins to various cellular sites 8. Understanding the inner workings of the T-cell activation process is crucial for both immunological research and clinical applications.
Various assays have been developed in order to investigate protein-protein interactions; however, biochemical assays, such as the widely used co-immunoprecipitation method, do not allow protein location to be discerned, thus precluding the observation of valuable insights into the dynamics of cellular mechanisms. Additionally, these bulk assays usually combine proteins from many different cells that might be at different stages of the investigated cellular process. This can have a detrimental effect on temporal resolution. The use of real-time imaging of live cells allows both the spatial tracking of proteins and the ability to temporally distinguish between signaling events, thus shedding light on the dynamics of the process 9,10. We present a method of real-time imaging of signaling-complex formation during T-cell activation. Primary T-cells or T-cell lines, such as Jurkat, are transfected with plasmids encoding for proteins of interest fused to monomeric fluorescent proteins, preventing non-physiological oligomerization 11. Live T cells are dropped over a coverslip pre-coated with T-cell activating antibody 8,9, which binds to the CD3/TCR complex, inducing T-cell activation while overcoming the need for specific activating antigens. Activated cells are constantly imaged with the use of confocal microscopy. Imaging data are analyzed to yield quantitative results, such as the colocalization coefficient of the signaling proteins.

We describe a live-cell imaging method that provides insight into protein dynamics during the T-cell activation process. We demonstrate the combined usage of the T-cell spreading assay, confocal microscopy and imaging analysis to yield quantitative results to follow signaling complex formation throughout T-cell activation.

Protection against infectious diseases is mediated by the immune system 1,2. T lymphocytes are the master coordinators of the immune system, regulating ...

High-throughput Quantitative Real-time RT-PCR Assay for Determining Expression Profiles of Types I and III Interferon Subtypes

Described in this report is a qRT-PCR assay for the analysis of seventeen human IFN subtypes in a 384-well plate format that incorporates highly specific locked nucleic acid (LNA) and molecular beacon (MB) probes, transcript standards, automated multichannel pipetting, and plate drying. Determining expression among the type I interferons (IFN), especially the twelve IFN-&#945; subtypes, is limited by their shared sequence identity; likewise, the sequences of the type III IFN, especially IFN-&#955;2 and -&#955;3, are highly similar. This assay provides a reliable, reproducible, and relatively inexpensive means to analyze the expression of the seventeen interferon subtype transcripts.

This protocol describes a high-throughput qRT-PCR assay for the analysis of type I and III IFN expression signatures. The assay discriminates single base pair differences between the highly similar transcripts of these genes. Through batch assembly and robotic pipetting, the assays are consistent and reproducible.

Described in this report is a qRT-PCR assay for the analysis of seventeen human IFN subtypes in a 384-well plate format that incorporates highly specific ...

Real-time Quaking-induced Conversion Assay for Detection of CWD Prions in Fecal Material

The RT-QuIC technique is a sensitive in vitro cell-free prion amplification assay based mainly on the seeded misfolding and aggregation of recombinant prion protein (PrP) substrate using prion seeds as a template for the conversion. RT-QuIC is a novel high-throughput technique which is analogous to real-time polymerase chain reaction (PCR). Detection of amyloid fibril growth is based on the dye Thioflavin T, which fluoresces upon specific interaction with &#7526;-sheet rich proteins. Thus, amyloid formation can be detected in real time. We attempted to develop a reliable non-invasive screening test to detect chronic wasting disease (CWD) prions in fecal extract. Here, we have specifically adapted the RT-QuIC technique to reveal PrPSc seeding activity in feces of CWD infected cervids. Initially, the seeding activity of the fecal extracts we prepared was relatively low in RT-QuIC, possibly due to potential assay inhibitors in the fecal material. To improve seeding activity of feces extracts and remove potential assay inhibitors, we homogenized the fecal samples in a buffer containing detergents and protease inhibitors. We also submitted the samples to different methodologies to concentrate PrPSc on the basis of protein precipitation using sodium phosphotungstic acid, and centrifugal force. Finally, the feces extracts were tested by optimized RT-QuIC which included substrate replacement in the protocol to improve the sensitivity of detection. Thus, we established a protocol for sensitive detection of CWD prion seeding activity in feces of pre-clinical and clinical cervids by RT-QuIC, which can be a practical tool for non-invasive CWD diagnosis.

Here, we present a protocol to describe a simple, fast and efficient prion amplification technique, the real-time quaking-induced conversion (RT-QuIC) method.

The RT-QuIC technique is a sensitive in vitro cell-free prion amplification assay based mainly on the seeded misfolding and aggregation of recombinant ...

In Vitro ELISA Test to Evaluate Rabies Vaccine Potency

The growing global concern for the animal welfare is encouraging manufacturers and the National Control Laboratories (OMCLs) to follow the 3Rs strategy for the Replacement, Reduction, and Refinement of the laboratory animal testing. The development of in vitro approaches is recommended at the WHO and European levels as alternatives to the NIH test for evaluating the rabies vaccine potency. At the surface of the rabies virus (RABV) particle, trimers of glycoprotein constitute the major immunogen to induce Viral Neutralizing Antibodies (VNAbs). An ELISA test, where Neutralizing Monoclonal Antibodies (mAb-D1) recognize the trimeric form of the glycoprotein, has been developed to determine the contents of the native folded trimeric glycoprotein along with the production of the vaccine batches. This in vitro potency test demonstrated a good concordance with the NIH test and has been found suitable in collaborative trials by RABV vaccine manufacturers and OMCLs. Avoidance of animal use is an achievable objective in the near future.
The method presented is based on an indirect ELISA sandwich immunocapture using the mAb-D1 which recognizes the antigenic sites III (aa 330 to 338) of the trimeric RABV glycoprotein, i.e., the immunogenic RABV antigen. mAb-D1 is used for both coating and detection of glycoprotein trimers present in the vaccine batch. Since the epitope is recognized because of its conformational properties, the potentially denatured glycoprotein (less immunogenic) cannot be captured and detected by the mAb-D1. The vaccine to be tested is incubated in a plate sensitized with the mAb-D1. Bound trimeric RABV glycoproteins are identified by adding the mAb-D1 again, labeled with peroxidase and then revealed in the presence of substrate and chromogen. Comparison of the absorbance measured for the tested vaccine and the reference vaccine allows for the determination of the immunogenic glycoprotein content.

Here we describe an indirect ELISA sandwich immunocapture to determine the immunogenic glycoprotein contents in rabies vaccines. This test uses a neutralizing Monoclonal Antibody (mAb-D1) recognizing glycoprotein trimers. It is an alternative to the in vivo NIH test to follow the consistency of vaccine potency during production.

The growing global concern for the animal welfare is encouraging manufacturers and the National Control Laboratories (OMCLs) to follow the 3Rs strategy ...

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. ...

Monitoring Influenza Virus Survival Outside the Host Using Real-Time Cell Analysis

Methods for virus particle quantification represent a critical aspect of many virology studies. Although several reliable techniques exist, they are either time-consuming or unable to detect small variations. Presented here is a protocol for the precise quantification of viral titer by analyzing electrical impedance variations of infected cells in real-time. Cellular impedance is measured through gold microelectrode biosensors located under the cells in microplates, in which magnitude depends on the number of cells as well as their size and shape. This protocol allows real-time analysis of cell proliferation, viability, morphology and migration with enhanced sensitivity. Also provided is an example of a practical application by quantifying the decay of influenza A virus (IAV) submitted to various physicochemical parameters affecting viral infectivity over time (i.e., temperature, salinity, and pH). For such applications, the protocol reduces the workload needed while also generating precise quantification data of infectious virus particles. It allows the comparison of inactivation slopes among different IAV, which reflects their capacity to persist in given environment. This protocol is easy to perform, is highly reproducible, and can be applied to any virus producing cytopathic effects in cell culture.

Reported here is a protocol for the quantification of infectious viral particles using real-time monitoring of electrical impedance of infected cells. A practical application of this method is presented by quantifying influenza A virus decay under different physicochemical parameters mimicking environmental conditions.

Methods for virus particle quantification represent a critical aspect of many virology studies. Although several reliable techniques exist, they are ...

Tools for the Real-Time Assessment of a Pseudomonas aeruginosa Infection Model

Pseudomonas aeruginosa (Pa) is one of the most common opportunistic pathogens associated with cystic fibrosis (CF). Once Pa colonization is established, a large proportion of the infecting bacteria form biofilms within airway sputum. Pa biofilms isolated from CF sputum have been shown to grow in small, dense aggregates of ~10-1,000 cells that are spatially organized and exhibit clinically relevant phenotypes such as antimicrobial tolerance. One of the biggest challenges to studying how Pa aggregates respond to the changing sputum environment is the lack of nutritionally relevant and robust systems that promote aggregate formation. Using a synthetic CF sputum medium (SCFM2), the life history of Pa aggregates can be observed using confocal laser scanning microscopy (CLSM) and image analysis at the resolution of a single cell. This in vitro system allows the observation of thousands of aggregates of varying size in real time, three dimensions, and at the micron scale. At the individual and population levels, having the ability to group aggregates by phenotype and position facilitates the observation of aggregates at different developmental stages and their response to changes in the microenvironment, such as antibiotic treatment, to be differentiated with precision.

Tools for the Real-Time Assessment of a Pseudomonas aeruginosa Infection Model

Synthetic cystic fibrosis sputum medium (SCFM2) can be utilized in combination with both confocal laser scanning microscopy and fluorescence-activated cell sorting to observe bacterial aggregates at high resolution. This paper details methods to assess aggregate populations during antimicrobial treatment as a platform for future studies.

Pseudomonas aeruginosa (Pa) is one of the most common opportunistic pathogens associated with cystic fibrosis (CF). Once Pa colonization is established, a ...

Real-time Monitoring of Mitochondrial Respiration in Cytokine-differentiated Human Primary T Cells

During activation, the metabolism of T cells adapts to changes that impact their fate. An increase in mitochondrial oxidative phosphorylation is indispensable for T cell activation, and the survival of memory T cells is dependent on mitochondrial remodeling. Consequently, this affects the long-term clinical outcome of cancer immunotherapies. Changes in T cell quality are often studied by flow cytometry using well-known surface markers and not directly by their metabolic state. This is an optimized protocol for measuring real-time mitochondrial respiration of primary human T cells using an Extracellular Flux Analyzer and the cytokines IL-2 and IL-15, which differently affect T cell metabolism. It is shown that the metabolic state of T cells can clearly be distinguished by measuring the oxygen consumption when inhibiting key complexes in the metabolic pathway and that the accuracy of these measurements is highly dependent on optimal inhibitor concentration and inhibitor injection strategy. This standardized protocol will help implement mitochondrial respiration as a standard for T cell fitness in monitoring and studying cancer immunotherapies.

Metabolic adaptation is fundamental for T cells as it dictates differentiation, persistence, and cytotoxicity. Here, an optimized protocol for monitoring mitochondrial respiration in ex vivo cytokine-differentiated human primary T cells is presented.

Pan-lyssavirus Real Time RT-PCR for Rabies Diagnosis

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Rozdziały w tym wideo

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

Real-time Live Imaging of T-cell Signaling Complex Formation

High-throughput Quantitative Real-time RT-PCR Assay for Determining Expression Profiles of Types I and III Interferon Subtypes

Real-time Quaking-induced Conversion Assay for Detection of CWD Prions in Fecal Material

In Vitro ELISA Test to Evaluate Rabies Vaccine Potency

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

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

Monitoring Influenza Virus Survival Outside the Host Using Real-Time Cell Analysis

Tools for the Real-Time Assessment of a Pseudomonas aeruginosa Infection Model

Real-time Monitoring of Mitochondrial Respiration in Cytokine-differentiated Human Primary T Cells