This work was supported through the Global Alliance for Vaccines and Immunisation (GAVI), the Wolfermann N&#228;geli Foundation, the Swiss African Research Cooperation (SARECO), the SWF Stiftung f&#252;r wissenschaftliche Forschung, the Freiwillige Akademische Gesellschaft (FAG) Basel, the Bilateral Science and Technology Cooperation Programme of Switzerland with Asia and the Novartis Foundation for biomedical research.
We thank especially the dog owners, the veterinary personnel and the laboratory staff for their great commitment. We also want to acknowledge Lisa Crump for the language editing.

1. Sample collection via the foramen magnum (occipital route)25
NOTE: This technique can be implemented under laboratory conditions or in field settings. Samples should be processed as soon as possible after death of the suspected animal or kept at cool temperature (refrigerated or frozen, if possible) to avoid decomposition which could affect the results. Similar to other reference techniques based on lyssavirus antigens detection such as DFAT and DRIT, decomposed samples should not be tested because it can affect the result (risk of false negative result).
CAUTION: All samples should be considered as potentially infectious. Safety regulations and procedures should be strictly followed, even in field settings4. In particular, wear appropriate personal protective equipment including mask, glasses, gloves and a lab coat. Use appropriate disinfectant for material and sample decontaminations (e.g., sodium hypochlorite with recommended manufacturer dilutions, 70% alcohol - ethanol or isopropanol, 1% soap solution). All personnel handling samples should be vaccinated against rabies.
<ol>
	<li>Remove the animal head with a knife before the first cervical vertebra (atlas vertebra) to access the foramen magnum. 
	NOTE: To minimize infective aerosol, avoid using a manual saw or similar tool.</li>
	<li>Collect brainstem (medulla oblongata) sample using a disposable plastic pipette (Figure 2A), a drinking straw (Figure 2B), a clamp (Figure 2C) or a dropper (supplied with the RIDT) (Figure 2D). 
	NOTE: Special attention must be paid when collecting the sample, because it is an utmost important step for the reliability of the results. In addition to the associated video which shows in a simple way how to collect the part of the brainstem of interest, a training step is highly recommended to make sure to collect the correct anatomical section.</li>
	<li>Optionally and in addition of brain stem (medulla oblongata), collect other parts of the brainstem or the brain (cerebellum, hippocampus, thalamus and cortex) by the same occipital route by pushing and rotating the plastic pipette or straw towards the eye socket (Figure 3).</li>
	<li>If using a straw or pipette, gently squeeze it to deposit the brain sample (0.5-2 g) in a tube for subsequent analysis and/or biobanking. 
	NOTE: Sample storage in glycerol is not recommended, as it seems to affect capillary flow or the antibody binding step of the RIDT18.</li>
</ol>
2. Execution of the modified RIDT protocol14
NOTE: This modification omits a dilution step (1:10) into PBS, as specified in manufacturer protocol (all versions), and can be implemented under laboratory or field settings.
<ol>
	<li>Use the swab/dropper to collect the equivalent of half a peanut or pea (0.1-0.5 g) of brain material and place it in the buffer sample tube. 
	NOTE: For the modified protocol, all reagents/consumables are included in the kit (no PBS or additional tube is needed) (Figure 4). Document the batch number of the kit and check validity of the expiration date.</li>
	<li>Carefully crush the brain material directly in the tube with the swab or the dropper for about 30 s until a homogeneous suspension is obtained. 
	NOTE: The buffer reaction inactivates the infectivity of the virus in the conditions of the manufacturer&#39;s protocol24.</li>
	<li>Using the dropper, deposit four drops (approximately 100 &#181;L) of the suspension in the sample inlet on the test device.</li>
	<li>Wait for complete sample migration (1-5 min) before reading the test device. The migration should start rapidly after deposit of the sample (1-5 min).</li>
	<li>In case of delay (due to high viscosity suspension) or to accelerate the start of the migration, gently scratch the bottom of the deposit site of the device with the dropper (1-5 times) and eventually add 1-2 more drops. Migration should start immediately thereafter.</li>
	<li>Read the test result in the detection window after 5-10 min, and no more than 20 min, after the end of the migration.</li>
	<li>Interpret the result based on presence or absence of the control line (C-line) and test line (T-line) (purple lines) in the detection window, according to Figure 5. Consider the sample positive when two lines are visible (Figure 5A), negative if only the C-line is present (Figure 5B) and invalid if only the T-line is present or if no lines are visible (Figure 5C). 
	NOTE: Invalid results should be repeated at least once. Other techniques should be performed if results remain invalid. Negative results obtained with RIDT need to be subsequently confirmed using a gold standard reference method, like DFAT, DRIT and molecular methods (polymerase chain reaction or PCR). Even though the sensitivity of this test is high (see representative results), it is not 100%.</li>
	<li>Store used devices at room temperature, or refrigerate/freeze when possible, for subsequent molecular analysis (see section 4). Freeze the remaining sample suspension at -20 &#176;C/-80 &#176;C in the buffer tube to repeat the test if necessary or for subsequent molecular analysis.</li>
</ol>
3. RNA extraction and detection by RT-qPCR from the RIDT device
NOTE: This step can only be implemented under laboratory conditions with adapted environment and suitable equipment for molecular diagnosis. It can be done soon after the RIDT test or retrospectively on archived RIDT devices, stored at room temperature (15-30 &#176;C), refrigerated or frozen.
<ol>
	<li>RNA extraction 
	NOTE: To monitor the extraction step, it is recommended to use an internal control that can be an endogenous mRNA (such as &#223;-actin) or an exogenous control (such as eGFP synthetic RNA) directly spiked into the sample during the first steps of the extraction26,27.
	<ol>
		<li>Carefully open the device and remove the filter paper.</li>
		<li>Cut the deposit area of the sample and place it into a tube containing 1 mL of Tri-Reagent LS. Incubate at RT for 1 hour with gentle regular manual agitation.</li>
		<li>Perform the extraction in accordance with manufacturer recommendations, as previously described27. At this step, the exogenous internal control can be added.</li>
		<li>During the process, add 2 &#181;L of glycogen for facilitating precipitation of RNA, according to the manufacturer recommendations.</li>
		<li>Adjust the final volume for RNA resuspension in nuclease-free water, with a volume of 50 &#181;L generally used. 
		NOTE: At the end of the centrifugation step for aqueous and organic phase separation (after addition of 200 &#181;L of chloroform into the Tri-Reagent LS), the piece of membrane from the device will be at the bottom of the tube and not interfere with collection of the upper aqueous phase. Alternatively, other easy and rapid protocols can be used, for instance, using phenol-based reagents and silica membranes28.</li>
	</ol>
	</li>
	<li>Detection by RT-qPCR26 
	NOTE: Detection of potential viral RNA present in extracted samples can be done using different molecular techniques, such as reverse-transcription PCR, conventional (endpoint) or real time PCR (qPCR). Several methods are available, such as conventional RT-PCR27,29 or RT-qPCR26,30 targeting the viral nucleoprotein or polymerase gene. One example will be presented below based on a dual combined pan-lyssavirus RT-qPCR targeting a conserved region among the viral polymerase. This RT-qPCR technique associates two different RT-qPCR: one based on the TaqMan probe technology (pan-RABV RT-qPCR) and the other using the SyBR Green detection (pan-lyssa RT-qPCR). In addition, the detection of an exogenous internal control (eGFP RNA) directly spiked during the extraction process is done by a specific TaqMan probe-based RT-qPCR (eGFP RT-qPCR). Careful on-site validation of the molecular techniques selected for detection of viral RNA is important, in particular, to verify that primers, and probes for real-time RT-PCR, are adapted for detection of the strains circulating in the region of interest4.
	<ol>
		<li>Dilute RNA sample to 1:10 in nuclease free water. Test each RNA sample in duplicate, using a 96-well reaction plate or other formats. Use positive and negative controls for each assay and test at least in duplicate.</li>
		<li>Prepare the master mix reaction solution for the three different RT-qPCR assays according to Table 1, and with the primers/probes indicated in Table 2.</li>
		<li>Add 5 &#181;L of diluted RNA samples and 15 &#181;L of master mix to each of the three different assays. The pan-RABV RT-qPCR assay and the eGFP RT-qPCR assay can cycle in the same plate.</li>
		<li>Run the different assays following the thermal cycling conditions indicated in Table 3. If only one PCR thermal cycler is available, start with the pan-RABV RT-qPCR and keep the plate for the pan-lyssa RT-qPCR at 4 &#176;C until the end of the pan-RABV RT-qPCR.</li>
		<li>Analyze the results obtained with the three assays according to Table 4.</li>
	</ol>
	</li>
</ol>
4. Genotyping after RNA extraction from the RIDT device
<ol>
	<li>Reverse transcription RT27,29
	<ol>
		<li>Prepare a master mix with 6 &#181;L of RNA, 2 &#181;L of pd(N)6 random primers (200 &#181;g/&#181;L) and 2 &#181;L of nuclease-free water for a final volume of 10 &#181;L.</li>
		<li>Incubate at 65 &#176;C for 10 min in a heat-block and then store on ice.</li>
		<li>Prepare a master mix with 6 &#181;L of 5x First-Strand Buffer, 2 &#181;L of 0.1 M dithiothreitol (DTT), 1 &#181;L (200 U) of Superscript II reverse transcriptase, 2 &#181;L (80 U) of RNasin, 2 &#181;L of dNTP mix (10 &#181;M) and complete with nuclease-free water to obtain a final volume of 20 &#181;L for each sample.</li>
		<li>Add the master mix (20 &#181;L) to the sample (10 &#181;L) (final volume of 30 &#181;L) and incubate at 42 &#176;C for 90 min in a heat-block.</li>
		<li>Proceed to the next step with PCR amplification or store the cDNA at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Conventional PCR27,29,31 
	NOTE: Different techniques of conventional PCR are available for genotyping. Two are presented, both hemi-nested PCR, targeting a part of the nucleoprotein or a part of the viral protein of the lyssavirus. The protocol is the same for each of these assays, except for the primers and cycling conditions. Positive (positive RNA) and negative (negative cDNA and/or nuclease-free water) controls should be included in each series and each round of PCR.
	<ol>
		<li>Prepare for each sample in a 0.2 mL microtube a master mix reaction solution for the first PCR step. This mix contains 5 &#181;L of 10x NH4 Reaction Buffer, 2.5 &#181;L of MgCl2 solution (50 mM), 1 &#181;L of dNTP Mix (10 &#181;M), 1 &#181;L of each primer (10 &#181;M), 0.2 &#181;L (1 U) of Biotaq DNA polymerase and 37.3 &#181;L of nuclease-free water (final volume of 48 &#181;L). The primers are indicated in Table 5.</li>
		<li>Add 2 &#181;L of cDNA in every tube and cycle on a separate conventional PCR thermal cycler for each assay, according to Table 6.</li>
		<li>Prepare a second master mix reaction solution identical to the previous one with using the appropriate primers (Table 5) for the hemi-nested PCR reaction.</li>
		<li>Add 2 &#181;L of the first round PCR product and cycle on a conventional PCR thermal cycler using the cycling parameters indicated in Table 6.</li>
		<li>Visualize the different PCR products (first and second round PCR) after loading them on a 1% agarose gel (100 mL of Tris-acetate EDTA buffer 1x - TAE 1x) with ethidium bromide (final concentration around 0.01%) and run the gel during 30 min at 120 V. A positive PCR result is observed in the form of a bright band of the expected size (Table 5).</li>
	</ol>
	</li>
	<li>Sanger sequencing
	<ol>
		<li>Perform a Sanger sequencing of the amplicons obtained with the pan-lyssavirus hemi-nested PCR and complete the genotyping analysis.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The RIDT is a simple, rapid and low-cost method for postmortem rabies diagnosis and a promising field alternative to laboratory testing. The application of such a test, especially for decentralized areas of low- and middle-income countries, would improve understanding of rabies virus prevalence and transmission on a local and potentially national scale. When combined with the rapid brain sample collection method (without full necropsy), a great advantage is that the test can be entirely performed in the field setting, away from laboratory facilities. Brain samples collected via the foramen magnum can be used for testing, thus it is not required to completely open the animal skull. The test is simple to perform and interpret and is particularly suitable for field surveillance activities14. Other advantages of the RIDT over the DFAT or DRIT are no need for positive and negative controls and kit storage at room temperature. In addition, the modified protocol, where the dilution step (1:10) into PBS is omitted, does not require extra reagents to perform the test and further simplifies the procedure under field conditions.
A key point is the quality of the brain samples. Samples should be collected and tested as soon as possible after death of the suspected animal, or kept at cool temperature before testing, to avoid degradation. Decomposed samples should not be tested because it can affect the result (risk of false negative result). Although no data are yet available regarding the loss of sensitivity of RIDT over time for brain samples, we hypothesize that it is similar compared to the DFAT test32. However, time between the death of the animal and performing the test can be reduced, as the test can be done quickly and directly in the field. Thus, there is in general a lower risk of decomposed samples.
Another critical step within the protocol is the sample suspension migration. The migration should start directly after deposit of the sample (1-5 min). High viscosity of the suspension could therefore negatively influence the migration. Gently scratching the bottom of the device deposit site with the dropper and adding 1-2 more drops often solves this problem, and the migration begins immediately after.
Most of the RIDT tests performed in African laboratories (Chad, Ivory Coast and Mali) were performed at ambient temperature which can exceed 30 &#176;C, whereas the range of temperature for storage and use recommended by the manufacturer is 15 &#176;C - 30 &#176;C. Although we did not identify any impact of high temperature on RIDT test performance, it is necessary to evaluate it more carefully. Similarly, the impact of high temperature during storage and transportation of the device after use for viral RNA detection and genotyping needs additional evaluation. The sensitivity of the viral RNA detection by RT-qPCR from the RIDT strip can be affected by the quality of the brain sample initially used in the test, but also by the condition of storage of the RIDT tests after use. For example, the sensitivity of the RNA detection was higher when used RIDT tests were stored under controlled laboratory conditions (94.7%) compared to under field conditions (e.g., Chad) (81.2%)14. These conditions might also affect the integrity (especially the length) of RNA fixed on the strip, possibly explaining the moderate sensitivity for genotyping based on longer PCR amplicons (e.g., &#62;500 nucleotides)14. The sensitivity of RT-qPCR performed on the test strip was lower than that obtained using FTA Whatman cards (80.6%)14. Similar to other molecular techniques, the viral load can also impact the success of genotyping based on RDIT strips, with potential negative results for samples with low viral load14.
The test is not currently recommended by WHO and OIE for routine diagnosis and disease surveillance, and a result cannot be used on its own to guide PEP decision making. Further test validation is still needed. However, accurate quick rabies diagnosis is a crucial element of well-functioning continuous rabies surveillance systems and is instrumental to increase political commitment, which is eminently important for successful sustainable rabies control33. RIDT tests offer new rabies diagnostic opportunities in this context and are a useful tool to expand animal rabies surveillance in the field in low- or middle-income enzootic areas.

Canine rabies is the main cause of human rabies, globally responsible for approximately 59,000 human deaths per year, nearly all occurring in low- and middle-income countries (LMICs) in Asia and Africa1. The main etiological agent is a neurotropic canine-associated classical rabies virus (RABV, family Rhabdoviridae, genus Lyssavirus, species Rabies lyssavirus). However, other rabies-related lyssaviruses, mostly circulating in bat species, also cause disease2,3. In affected regions, disease surveillance and control are often hampered by low level political commitment likely due to lack of reliable data4,5,6. One reason for disease underreporting is the absence of laboratory diagnosis, due in part to limited access to equipped laboratories and trained staff as well as the difficulties of shipment of the specimens. Laboratory diagnosis is necessary to confirm rabies cases and additionally allows for genetic characterization of the involved strains, providing insight on virus transmission at the regional level4,5,7.
The current gold standards for postmortem rabies diagnosis, approved by both the World Health Organization (WHO) and the World Organization for Animal Health (OIE), are the direct fluorescent antibody test (DFAT), the direct rapid immunohistochemistry test (DRIT) and molecular methods (e.g., reverse transcription polymerase chain reaction (RT-PCR))4,8. However, proper application in LMICs remains limited due to inadequate laboratory facilities with inconsistent power supply, uncooled sample transportation, and lack of a quality management system. Because animal rabies diagnosis is typically only conducted at central veterinary laboratories in LMICs, existing surveillance data mainly reflects the rabies situation in urban areas.
Recently developed low technology diagnostic alternatives offer opportunities to establish rabies diagnosis in remote areas and decentralized rabies laboratories4,8,9. The rapid immunochromatographic diagnostic test (RIDT) is a lateral flow test based on immunochromatography using gold conjugated detector antibodies and is a very promising rabies diagnostic tool10,11,12,13. The principle is simple: after dilution, brain material is mixed in the provided buffer, and a few drops are applied on the test strip where gold conjugated monoclonal antibodies bind specifically to rabies antigens, mainly the nucleoproteins (Figure 1). The antigen-antibody complexes then undergo lateral flow migration, binding at the test line (T-line) to fixed antibodies against rabies antigens, resulting in a clearly visible purple line. The remaining gold conjugated antibodies not bound to rabies antigens continue migrating and fix to the membrane through additional targeting antibodies, resulting in a clearly visible purple control line (C-line).
The one-step, low cost method is rapid, extremely easy and does not require expensive equipment or special storage conditions. With modification of the manufacturer protocol to eliminate the dilution step, nearly all equipment and reagents required to perform the test are included in the kit14. The result is read after 5-10 minutes without a microscope. This is a major advantage over the DFAT test, which requires a fluorescence microscope and immunofluorescence conjugate, along with refrigerated transportation and sample storage. Even the DRIT test, which can be performed using a light microscope, requires a continuous cold chain to store the anti-rabies antibodies, which are also not yet commercially available. In comparison to the DRIT, the RIDT requires no toxic chemicals, a particular advantage in countries where waste disposal is poorly regulated. The rapid test is less time-consuming with much easier interpretation compared to the gold standard tests DFAT and DRIT. This allows for on-site testing by personnel with limited technical expertise.
Based on these test properties, prompt diagnosis of suspected animals in remote areas becomes feasible, facilitating implementation of post exposure prophylaxis (PEP) for exposed people as soon as possible. In addition, distance transport of rabies samples is not necessary, resulting in better sample quality at the time of testing. However, the results obtained with the RIDT tests should currently be confirmed using a reference diagnostic test such as DFAT or DRIT.
RIDT techniques for detection of RABV and other lyssaviruses have been evaluated. One of the first studies was conducted by Korean researchers in 200710. Compared to the DFAT method, in 51 animal samples and 4 RABV isolates, the RIDT showed a sensitivity and specificity of 91.7% and 100%, respectively. These results were later confirmed with 110 animal brain samples from Korea, with sensitivity and specificity, compared to DFAT, of 95% and 98.9%, respectively15. More recently, other studies assessed the performance of this RIDT using virus isolates and/or infected brain samples from various animals with different geographical origins. A panel of 21 samples, including African RABV and other African lyssaviruses (Duvenhage virus (DUVV), Lagos bat virus (LBV) and Mokola virus (MOKV)), were successfully detected, with sensitivity of 100% compared to the DFAT16. Similar high sensitivity (96.5%) and specificity (100%) values were obtained from a panel of 115 brain samples from Ethiopia17. Another study evaluated European RABV isolates, two other European lyssaviruses (European bat lyssavirus type 1 (EBLV-1) and type 2 (EBLV-2)), and the Australian bat lyssavirus (ABLV)18. Based on analysis of 172 animal brain samples, the RIDT kit had 88.3% sensitivity and 100% specificity compared to DFAT, and the three rabies-related lyssaviruses were successfully detected. In this study, some of the false negative results came from brain samples stored in glycerol buffer, suggesting that improper glycerol removal influenced capillary flow or antibody binding. A recent analysis of 43 clinical samples from Australian bats confirmed previous test results, with complete concordance to DFAT19. Two studies were conducted in India using the RIDT on a limited number of clinical samples (11 and 34 samples). Compared to DFAT, sensitivity was between 85.7% and 91.7% and specificity was 100%20,21. Another evaluation of this kit using 80 animal brain samples from Africa, Europe and the Middle East obtained complete concordance with DFAT for specificity (100%) but a higher sensitivity (96.9%) compared to the previous studies22. In a recent inter-laboratory comparison of this RIDT performed in 22 different laboratories using a panel of 10 samples, overall concordance was 99.5%23.
Only one recent multicentric study showed unsatisfactory overall RIDT performance24. Samples from three different datasets were tested and provided variable sensitivity and specificity values compared to DFAT. For example, sensitivity and specificity obtained with the first panel (n=51) and the second panel (n=31) of samples from experimental infected animals, all tested in laboratory A, gave a sensitivity of 16% and 43%, respectively, whereas the specificity was 100% for both. Conversely, the results of the third panel (n=30) of field clinical samples analyzed by laboratory B provided a complete concordance with the results of DFAT, which was further nearly completely confirmed by laboratory A (85% sensitivity and 100% specificity). Batch-to-batch variation was suggested as a possible explanation for the fluctuating relatively low sensitivity with RIDT24.
At the same time, another study performed a similar validation process of the above described RIDT, with a modification of the manufacturer recommended protocol14. The pre-dilution step (1:10) in PBS was omitted during preparation of the brain material. Based on this simpler modified protocol, the authors obtained sensitivity and specificity of 95.3% and 93.3%, respectively, compared to DFAT by testing, under laboratory conditions, a dataset of 73 animal brain samples, naturally or experimentally infected with various RABV strains. The study presented the first evaluation of this RIDT in a field setting (Chad, Africa). In 48 clinical brain samples, sensitivity and specificity were 94.4% and 100%, respectively. The discrepancies between DFAT and RIDT were due to false positive results with DFAT, determined after confirmation with RT-PCR. When these results were deleted, there was complete concordance, and it demonstrated that the RIDT was more reliable that DFAT under these field conditions14. No batch-to-batch variation was observed using the modified protocol. When the modified protocol was applied to a small number of the DFAT/ RIDT divergent samples (n=8) in the study of Eggerbauer et al.24, all were found concordant (100% sensitivity).
Another major advantage of the RIDT is secondary use for detecting viral RNA fixed on the strip using molecular techniques (such as RT-PCR) and subsequent genotyping14,24. Following an extraction step, L&#233;chenne et al.14 demonstrated viral RNA fixed on the Anigen device membrane using RT-PCR with 86.3% sensitivity in a panel of 51 samples (including 18 samples tested and shipped from Chad at ambient temperature). Subsequent genotyping was possible in 93% of the 14 samples tested. Sanger sequencing of PCR amplicons of at least 500 nucleotides in length were used. In addition to RABV isolates, the test detected four other lyssavirus species, DUVV, EBLV-1, EBLV-2 and Bokeloh bat lyssavirus (BBLV), during a fully concordant international inter laboratory test14. The sensitivity of viral RNA detection was even higher (100%) in the study of Eggerbauer et al., based on laboratory samples examination24. The latter study also demonstrated that the buffer used in the RIDT kit inactivated virus. Thereby, the devices can be shipped easily, at ambient temperature without specific biosafety precautions to reference laboratories, for molecular confirmation and genotyping.
Based on the previous evaluations, RIDT tools offer numerous advantages for use in field settings, especially when the reference diagnostic techniques are not available. However, this test also has some limitations, in particular, low sensitivity of antigen detection14,24. The test is applicable for samples containing high quantities of viral antigens, such as brain samples. However, it is not appropriate for other samples such as saliva or other body fluids. Another drawback is cost of the device (around 5-10 Euros in Europe), which is less expensive compared to the cost of performing DFAT, RT-PCR or DRIT, but which still remains high for LMICs. However, future development and validation of similar RIDTs from other companies could lead to a price decrease. One study reported batch-to-batch variations. Although not reported by others, strict quality controls should nevertheless be performed when testing a new batch, as for any reagent used in a quality management environment. The use of the modified protocol was not altered when using different batches14. All except one study demonstrated that the sensitivity of RDIT was high compared to DFAT (around 90%-95%). Because rabies is always fatal, it is still strongly recommended to confirm any negative results with RDIT using a reference diagnostic test such as DFAT, DRIT or RT-PCR14.
In this manuscript, we present a complete protocol for field postmortem diagnosis of animal rabies based on an example of a commercialized RIDT, from brain sample collection to application of a modified protocol compared to the manufacturer recommendations (which were previously validated14) and subsequent molecular analysis. This protocol was applied and validated many times under field conditions in West- and Central Africa, where the RIDT was used routinely for rabies diagnosis alongside the DFAT test. We additionally demonstrate a second application for the device, in laboratory settings, for extraction and detection using RT-PCR of viral RNA fixed on the device.

<table><tbody><tr><td>Anti-Rabies Nucleocapsid Conjugate (lyophilizied, adsorbed)</td><td>Bio-Rad, France</td><td>3572112</td><td>Fluorescein-5-isothiocyanate (FITC) conjugated polyclonal antibody against the nucleocapsid of rabies virus. Use for the DFAT reference tecnnique.</td></tr><tr><td>Applied Biosystems 7500 Real-Time PCR System</td><td>Applied Biosystems, France</td><td>4351104</td><td>Amplification of the viral RNA by RT-qPCR after extraction from the RIDT device.</td></tr><tr><td>Disposable plastic pipette, drinking straw, clamp, dropper</td><td>-</td><td>-</td><td>Equipment used for the collection of the brain stem (medulla oblongata) via the foramen magnus (occipital route).</td></tr><tr><td>Evans Blue Solution 1%</td><td>Bio-Rad, France</td><td>3574911</td><td>Counter-coloration used for the DFAT to facilite the reading under UV microscope.</td></tr><tr><td>Primer eGFPF1</td><td>Eurofins Genomics, Germany</td><td>-</td><td>Forward primer for the amplification of the internal control eGFP by RT-qPCR after RNA extraction from the RIDT device, sequence: 5&#039;-GACCACTACCAGCAGAACAC-3&#039;.</td></tr><tr><td>Primer eGFPR2</td><td>Eurofins Genomics, Germany</td><td>-</td><td>Reverse primer for the amplification of the internal control eGFP by RT-qPCR after extraction from the RIDT device, sequence: 5&#039;-GAACTCCAGCAGGACCATG-3&#039;.</td></tr><tr><td>Primer Taq17 revlong</td><td>Eurofins Genomics, Germany</td><td>-</td><td>Reverse primer for the amplification of the viral RNA by RT-qPCR after RNA extraction from the RIDT device, sequence: 5&#039;-ATGAGAAGTGG&lt;br/&gt; AAYAAYCATCA-3&#039;.</td></tr><tr><td>Primer Taq3 long</td><td>Eurofins Genomics, Germany</td><td>-</td><td>Forward primer for the amplification of the viral RNA by RT-qPCR after RNA extraction from the RIDT device, sequence: 5&#039;-GATCTGTCTGAA&lt;br/&gt; TAATAGAYCCARG-3&#039;.</td></tr><tr><td>Probe eGFP FAM/TAMRA</td><td>Eurofins Genomics, Germany</td><td>-</td><td>Forward primer for the amplification of the internal control by RT-qPCR after extraction from the RIDT device, sequence: 5&#039;-AGCACCCAGT&lt;br/&gt; CCGCCCTGAGCA-3&#039;.</td></tr><tr><td>Probe RABV4 FAM/TAMRA</td><td>Eurofins Genomics, Germany</td><td>-</td><td>Probe for the amplification of the viral RNA by RT-qPCR after RNA extraction from the RIDT device, sequence: 5&#039;-AACACYTGATCBA&lt;br/&gt; GKACAGARAAYACATC-3&#039;.</td></tr><tr><td>Probe RABV5 FAM/TAMRA</td><td>Eurofins Genomics, Germany</td><td>-</td><td>Probe for the amplification of the viral RNA by RT-qPCR after RNA extraction from the RIDT device, sequence: 5&#039;-AGRGTGTTTTCYAG&lt;br/&gt; RACWCAYGAGTTTTTYCA-3&#039;.</td></tr><tr><td>Rapid Rabies Ag Test Kit</td><td>BioNote Inc., Republic of Korea</td><td>RG18-01DD</td><td>Rapid immunochromatographic diagnostic test (RIDT, also named lateral flow device or LFD) for the post-mortem diagnosis of rabies.</td></tr><tr><td>Recombinant RNasin Ribonuclease Inhibitor</td><td>Promega, USA</td><td>N2515</td><td>Enzyme used with the kit for the amplification of the viral RNA by RT-qPCR after extraction from the RIDT device.</td></tr><tr><td>SuperScript III Platinum One-Step qRT-PCR Kit</td><td>Invitrogen, France</td><td>11732-020</td><td>Kit for the amplification of the viral RNA by RT-qPCR after extraction from the RIDT device.</td></tr><tr><td>TRIzol Reagent</td><td>Invitrogen, France</td><td>15596026</td><td>Phenol/chloroforme based total RNA extraction using the cellulose membrane of the RIDT.</td></tr></tbody></table>

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.

Watch this Scientific Journal Video about Field Postmortem Rabies Rapid Immunochromatographic Diagnostic Test for Resource-Limited Settings with Further Molecular Applications at JoVE.com

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

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, ...

field-postmortem-rabies-rapid-immunochromatographic-diagnostic-test

Research

JoVE Journal

Immunology and Infection

13.3K Views.  Institut Pasteur. Dit protocol kan worden gebruikt voor de postmortemdiagnose van dierlijke hondsdolheid vanaf de eerste weergave van de dictie in het veld tot de moleculaire conformatie in een gecentraliseerd niveau van activiteit. Snelle immunochromatografische diagnosetesten is een nuttig alternatief voor landelijke en landelijke eigenschappen in Afrika en Azië, waarbij referentiematerialen voor rabiësdiagnose niet beschikbaar zijn. Het aantonen van de procedure zal Simon Bonas een technicus van mijn labo...

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

Combination of Adhesive-tape-based Sampling and Fluorescence in situ Hybridization for Rapid Detection of Salmonella on Fresh Produce

This protocol describes a simple approach for adhesive-tape-based sampling of tomato and other fresh produce surfaces, followed by on-tape fluorescence in situ hybridization (FISH) for rapid culture-independent detection of Salmonella spp. Cell-charged tapes can also be placed face-down on selective agar for solid-phase enrichment prior to detection. Alternatively, low-volume liquid enrichments (liquid surface miniculture) can be performed on the surface of the tape in non-selective broth, followed by FISH and analysis via flow cytometry. To begin, sterile adhesive tape is brought into contact with fresh produce, gentle pressure is applied, and the tape is removed, physically extracting microbes present on these surfaces. Tapes are mounted sticky-side up onto glass microscope slides and the sampled cells are fixed with 10% formalin (30 min) and dehydrated using a graded ethanol series (50, 80, and 95%; 3 min each concentration). Next, cell-charged tapes are spotted with buffer containing a Salmonella-targeted DNA probe cocktail and hybridized for 15 - 30 min at 55&deg;C, followed by a brief rinse in a washing buffer to remove unbound probe. Adherent, FISH-labeled cells are then counterstained with the DNA dye 4',6-diamidino-2-phenylindole (DAPI) and results are viewed using fluorescence microscopy. For solid-phase enrichment, cell-charged tapes are placed face-down on a suitable selective agar surface and incubated to allow in situ growth of Salmonella microcolonies, followed by FISH and microscopy as described above. For liquid surface miniculture, cell-charged tapes are placed sticky side up and a silicone perfusion chamber is applied so that the tape and microscope slide form the bottom of a water-tight chamber into which a small volume (&le; 500 &mu;L) of Trypticase Soy Broth (TSB) is introduced. The inlet ports are sealed and the chambers are incubated at 35 - 37&deg;C, allowing growth-based amplification of tape-extracted microbes. Following incubation, inlet ports are unsealed, cells are detached and mixed with vigorous back and forth pipetting, harvested via centrifugation and fixed in 10% neutral buffered formalin. Finally, samples are hybridized and examined via flow cytometry to reveal the presence of Salmonella spp. As described here, our "tape-FISH" approach can provide simple and rapid sampling and detection of Salmonella on tomato surfaces. We have also used this approach for sampling other types of fresh produce, including spinach and jalape&ntilde;o peppers.

Combination of Adhesive-tape-based Sampling and Fluorescence in situ Hybridization for Rapid Detection of Salmonella on Fresh Produce

This protocol describes a simple adhesive-tape-based approach for sampling of tomato and other fresh produce surfaces, followed by rapid whole cell detection of Salmonella using fluorescence in situ hybridization (FISH).

This protocol describes a simple approach for adhesive-tape-based sampling of tomato and other fresh produce surfaces, followed by on-tape fluorescence in ...

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

Enhanced Rabies Surveillance Using a Direct Rapid Immunohistochemical Test

Laboratory-based surveillance is integral for rabies prevention, control and management efforts. While the DFA is the gold standard for rabies diagnosis, there is a need to validate additional diagnostic techniques to improve rabies surveillance, particularly in developing countries. Here, we present a standard protocol for the DRIT as an alternative, laboratory or field-based testing option that uses light microscopy as compared to the DFA. Touch impressions of brain tissue collected from suspect animals are fixed in 10% buffered formalin. The DRIT uses rabies virus-specific monoclonal or polyclonal antibodies (conjugated to biotin), a streptavidin-peroxidase enzyme, and a chromogen reporter (such as acetyl 3-amino-9-ethylcarbazole) to detect viral inclusions within infected tissue. In approximately 1 h, a brain tissue sample can be tested and interpreted by the DRIT. Evaluation of suspect animal brains tested from a variety of species in North America, Asia, Africa, and Europe have illustrated high sensitivity and specificity by the DRIT approaching 100% with results compared to DFA. Since 2005, the United States Department of Agriculture&#8217;s Wildlife Services (USDA WS) program has conducted large-scale enhanced rabies surveillance efforts using the DRIT to test &#62;94,000 samples collected from wildlife in strategic rabies management areas. The DRIT provides a powerful, economical tool for rabies diagnosis that can be used by laboratorians and field biologists to improve current rabies surveillance, prevention and control programs globally.

The direct rapid immunohistochemical test (DRIT) offers a World Organization for Animal Health and World Health Organization (OIE/WHO) recognized alternative to the direct fluorescent antibody (DFA) test for rabies diagnosis. This test allows for field-based applications that can be performed in approximately 1 h upon brain impressions using light microscopy.

Laboratory-based surveillance is integral for rabies prevention, control and management efforts. While the DFA is the gold standard for rabies diagnosis, ...

Pan-lyssavirus Real Time RT-PCR for Rabies Diagnosis

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

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

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

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 ...

Quick and Early Detection of Rabies Antibodies to Evaluate the Immune Response in a Patient

Detection of Rabies IgG and IgM Antibodies Using the Rabies Indirect Fluorescent Antibody Test

The rabies indirect fluorescent antibody (IFA) test was developed to detect various rabies-specific antibody isotypes in sera or cerebral spinal fluid. This test provides rapid results and can be used to detect rabies antibodies in several different scenarios. The rabies IFA test is especially useful for the quick and early detection of antibodies to evaluate the immune response in a patient who has developed rabies. Although other methods for antemortem rabies diagnosis take precedence, this test may be utilized to demonstrate recent rabies virus exposure through antibody detection. The IFA test does not provide a virus-neutralizing antibody (VNA) titer, but the pre-exposure prophylaxis (PrEP) response can be evaluated through positive or negative antibody presence. This test can be utilized in various situations and can provide results for a number of different targets. In this study, we used several paired serum samples from individuals who received PrEP and demonstrated their rabies antibody presence over time using the IFA test.

Author Spotlight: Rabies-Specific Antibody Isotypes Detection in Sera or Cerebral Spinal Fluid Using an IFA Test 

The aim of this manuscript is to examine the use of the rabies indirect fluorescent antibody test for the detection of rabies-specific IgG and IgM antibodies.

The rabies indirect fluorescent antibody (IFA) test was developed to detect various rabies-specific antibody isotypes in sera or cerebral spinal fluid. ...

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.

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

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