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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#39;-GACCACTACCAGCAGAACAC-3&#39;.</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&#39;-GAACTCCAGCAGGACCATG-3&#39;.</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&#39;-ATGAGAAGTGG 
			AAYAAYCATCA-3&#39;.</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&#39;-GATCTGTCTGAA 
			TAATAGAYCCARG-3&#39;.</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&#39;-AGCACCCAGT 
			CCGCCCTGAGCA-3&#39;.</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&#39;-AACACYTGATCBA 
			GKACAGARAAYACATC-3&#39;.</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&#39;-AGRGTGTTTTCYAG 
			RACWCAYGAGTTTTTYCA-3&#39;.</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

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Immunology and Infection

13.1K Views.  Institut Pasteur. 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.

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 ...

Rapid Diagnosis of Avian Influenza Virus in Wild Birds: Use of a Portable rRT-PCR and Freeze-dried Reagents in the Field

Wild birds have been implicated in the spread of highly pathogenic avian influenza (HPAI) of the H5N1 subtype, prompting surveillance along migratory flyways. Sampling of wild birds for avian influenza virus (AIV) is often conducted in remote regions, but results are often delayed because of the need to transport samples to a laboratory equipped for molecular testing. Real-time reverse transcriptase polymerase chain reaction (rRT-PCR) is a molecular technique that offers one of the most accurate and sensitive methods for diagnosis of AIV. The previously strict lab protocols needed for rRT-PCR are now being adapted for the field. Development of freeze-dried (lyophilized) reagents that do not require cold chain, with sensitivity at the level of wet reagents has brought on-site remote testing to a practical goal.
Here we present a method for the rapid diagnosis of AIV in wild birds using an rRT-PCR unit (Ruggedized Advanced Pathogen Identification Device or RAPID, Idaho Technologies, Salt Lake City, UT) that employs lyophilized reagents (Influenza A Target 1 Taqman; ASAY-ASY-0109, Idaho Technologies). The reagents contain all of the necessary components for testing at appropriate concentrations in a single tube: primers, probes, enzymes, buffers and internal positive controls, eliminating errors associated with improper storage or handling of wet reagents. The portable unit performs a screen for Influenza A by targeting the matrix gene and yields results in 2-3 hours. Genetic subtyping is also possible with H5 and H7 primer sets that target the hemagglutinin gene.
The system is suitable for use on cloacal and oropharyngeal samples collected from wild birds, as demonstrated here on the migratory shorebird species, the western sandpiper (Calidrus mauri) captured in Northern California. Animal handling followed protocols approved by the Animal Care and Use Committee of the U.S. Geological Survey Western Ecological Research Center and permits of the U.S. Geological Survey Bird Banding Laboratory. The primary advantage of this technique is to expedite diagnosis of wild birds, increasing the chances of containing an outbreak in a remote location. On-site diagnosis would also prove useful for identifying and studying infected individuals in wild populations. The opportunity to collect information on host biology (immunological and physiological response to infection) and spatial ecology (migratory performance of infected birds) will provide insights into the extent to which wild birds can act as vectors for AIV over long distances.

This study describes diagnosis of avian influenza in wild birds using a portable rRT-PCR system. The method takes advantage of freeze-dried reagents to screen wild birds in a non-laboratory setting, typical of an outbreak scenario. Use of molecular tools provides accurate and sensitive alternatives for rapid diagnosis.

Wild birds have been implicated in the spread of highly pathogenic avian influenza (HPAI) of the H5N1 subtype, prompting surveillance along migratory ...

Diagnosing Pulmonary Tuberculosis with the Xpert MTB/RIF Test

Tuberculosis (TB) due to Mycobacterium tuberculosis (MTB) remains a major public health issue: the infection affects up to one third of the world population1, and almost two million people are killed by TB each year.2 Universal access to high-quality, patient-centered treatment for all TB patients is emphasized by WHO's Stop TB Strategy.3 The rapid detection of MTB in respiratory specimens and drug therapy based on reliable drug resistance testing results are a prerequisite for the successful implementation of this strategy. However, in many areas of the world, TB diagnosis still relies on insensitive, poorly standardized sputum microscopy methods. Ineffective TB detection and the emergence and transmission of drug-resistant MTB strains increasingly jeopardize global TB control activities.2
Effective diagnosis of pulmonary TB requires the availability - on a global scale - of standardized, easy-to-use, and robust diagnostic tools that would allow the direct detection of both the MTB complex and resistance to key antibiotics, such as rifampicin (RIF). The latter result can serve as marker for multidrug-resistant MTB (MDR TB) and has been reported in &gt; 95% of the MDR-TB isolates.4, 5 The rapid availability of reliable test results is likely to directly translate into sound patient management decisions that, ultimately, will cure the individual patient and break the chain of TB transmission in the community.2
Cepheid's (Sunnyvale, CA, U.S.A.) Xpert MTB/RIF assay6, 7 meets the demands outlined above in a remarkable manner. It is a nucleic-acids amplification test for 1) the detection of MTB complex DNA in sputum or concentrated sputum sediments; and 2) the detection of RIF resistance-associated mutations of the rpoB gene.8 It is designed for use with Cepheid's GeneXpert Dx System that integrates and automates sample processing, nucleic acid amplification, and detection of the target sequences using real-time PCR and reverse transcriptase PCR. The system consists of an instrument, personal computer, barcode scanner, and preloaded software for running tests and viewing the results.9 It employs single-use disposable Xpert MTB/RIF cartridges that hold PCR reagents and host the PCR process. Because the cartridges are self-contained, cross-contamination between samples is eliminated.6 Current nucleic acid amplification methods used to detect MTB are complex, labor-intensive, and technically demanding. The Xpert MTB/RIF assay has the potential to bring standardized, sensitive and very specific diagnostic testing for both TB and drug resistance to universal-access point-of-care settings3, provided that they will be able to afford it. In order to facilitate access, the Foundation for Innovative New Diagnostics (FIND) has negotiated significant price reductions. Current FIND-negotiated prices, along with the list of countries eligible for the discounts, are available on the web.10

The Xpert MTB/RIF test integrates sample decontamination, hands-free operation, on-board sample processing, and ultra-sensitive hemi-nested PCR for the simultaneous detection of Mycobacterium tuberculosis and rifampicin resistance, either in expectorated sputum or concentrated sputum sediments, in approximately two hours. Testing is standardized and requires only moderate laboratory infrastructure and training.

Tuberculosis (TB) due to Mycobacterium tuberculosis (MTB) remains a major public health issue: the infection affects up to one third of the world ...

Monitoring Dendritic Cell Migration using 19F / 1H Magnetic Resonance Imaging

Continuous advancements in noninvasive imaging modalities such as magnetic resonance imaging (MRI) have greatly improved our ability to study physiological or pathological processes in living organisms. MRI is also proving to be a valuable tool for capturing transplanted cells in vivo. Initial cell labeling strategies for MRI made use of contrast agents that influence the MR relaxation times (T1, T2, T2*) and lead to an enhancement (T1) or depletion (T2*) of signal where labeled cells are present. T2* enhancement agents such as ultrasmall iron oxide agents (USPIO) have been employed to study cell migration and some have also been approved by the FDA for clinical application. A drawback of T2* agents is the difficulty to distinguish the signal extinction created by the labeled cells from other artifacts such as blood clots, micro bleeds or air bubbles. In this article, we describe an emerging technique for tracking cells in vivo that is based on labeling the cells with fluorine (19F)-rich particles. These particles are prepared by emulsifying perfluorocarbon (PFC) compounds and then used to label cells, which subsequently can be imaged by 19F MRI. Important advantages of PFCs for cell tracking in vivo include (i) the absence of carbon-bound 19F in vivo, which then yields background-free images and complete cell selectivityand(ii) the possibility to quantify the cell signal by 19F MR spectroscopy.

Monitoring Dendritic Cell Migration using 19F / 1H Magnetic Resonance Imaging

Tracking of cells using MRI has gained remarkable attention in the past years. This protocol describes the labeling of dendritic cells with fluorine (19F)-rich particles, the in vivo application of these cells, and monitoring the extent of their migration to the draining lymph node with 19F/1H MRI and 19F MRS.

Continuous  advancements in noninvasive imaging modalities such as magnetic resonance  imaging (MRI) have greatly improved our ability to study ...

Rapid and Robust Analysis of Cellular and Molecular Polarization Induced by Chemokine Signaling

Cells respond to chemokine stimulation by losing their round shape in a process called polarization, and by altering the subcellular localization of many proteins. Classic imaging techniques have been used to study these phenomena. However, they required the manual acquisition of many cells followed by time consuming quantification of the morphology and the co-localization of the staining of tens of cells. Here, a rapid and powerful method is described to study these phenomena on samples consisting of several thousands of cells using an imaging flow cytometry technology that combines the advantages of a microscope with those of a cytometer. Using T lymphocytes stimulated with CCL19 and staining for MHC Class I molecules and filamentous actin, a gating strategy is presented to measure simultaneously the degree of shape alterations and the extent of co-localization of markers that are affected by CCL19 signaling. Moreover, this gating strategy allowed us to observe the segregation of filamentous actin (at the front) and phosphorylated Ezrin-Radixin-Moesin (phospho-ERM) proteins (at the rear) in polarized T cells after CXCL12 stimulation. This technique was also useful to observe the blocking effect on polarization of two different elements: inhibition of actin polymerization by a pharmacological inhibitor and expression of mutants of the Par6/atypical PKC signaling pathway. Thus, evidence is shown that this technique is useful to analyze both morphological alterations and protein redistributions.

Chemokine signaling elicits marked alterations of cellular morphology and some important redistributions of intracellular proteins. Here, a rapid and detailed protocol is provided to study these events.

Cells respond to chemokine stimulation by losing their round shape in a process called polarization, and by altering the subcellular localization of many ...

Rapid Molecular Detection and Differentiation of Influenza Viruses A and B

Influenza is a contagious respiratory illness caused by influenza viruses A and B in humans and causes a significant amount of morbidity and mortality every year. The Influenza A and B assay was the first CLIA-waived molecular rapid flu test available. The Influenza A and B test works by employing isothermal amplification with influenza-specific primers followed by target detection with molecular beacon probes. Here, the performance of the Influenza A and B assay on frozen, archived nasopharyngeal swab (NPS) specimens stored in viral transport medium (VTM) were compared to a respiratory panel assay.
The performance of the Influenza A and B assay was evaluated by comparing the results to the respiratory panel reference method. The sensitivity for total influenza virus A was 67.5% (95% CI (CI), 56.6-78.5) and the specificity was 86.9% (CI, 71.0-100). For influenza virus B testing, the sensitivity and specificity were 90.2% (CI, 68.5-100) and 98.8% (CI, 68.5-100), respectively.
This system has the advantage of a significantly shorter test time than any other currently available molecular assay and the simple, pipette-free procedure runs on a fully integrated, closed, small-footprint system. Overall, the Influenza A and B assay evaluated in this study has the potential to serve as a point-of-care rapid influenza diagnostic test.

We describe a rapid, molecular-based Influenza A and B assay. The Influenza assay detects each target within 15 min by employing isothermal amplification with influenza-specific primers followed by target detection with molecular beacon probes. The Influenza A and B assay is user-friendly and required minimal hands-on time to perform.

Influenza is a contagious respiratory illness caused by influenza viruses A and B in humans and causes a significant amount of morbidity and mortality ...

Establishment and Analysis of Tumor Slice Explants As a Prerequisite for Diagnostic Testing

Organotypic primary tissue explant cultures, which include precision-cut slices, represent the three-dimensional (3-D) tissue architecture as well as the multicellular interactions of native tissue. Tissue&#160;slices immediately cut from freshly resected tumors preserve spatial aspects of intratumor heterogeneity (ITH), thus making them useful surrogates of in vivo biology. Careful optimization of tissue slice preparation and cultivation conditions is fundamental for the predictive diagnostic potential of tumor slice explants. In this regard, murine models are valuable, as these provide a consistent flow of tumor material to perform replicate and reproducible experiments. This protocol describes the culturing of murine lung tumor-derived tissue slices using a rotating incubation unit, a system that enables intermittent exposure of tissues to oxygen and nutrients. Our previous work showed that the use of rotating incubation units improves the viability of tissue compared to other culture methods, particularly floating slices and stagnant filter supports. Here, we further show that slice thickness influences the viability of cultured slices, suggesting that optimization of slice thickness should be done for different tissue types. Pronounced ITH in relevant oncogenic functions, such as signaling activities, stromal cell infiltration or expression of differentiation markers, necessitates evaluation of adjacent tissue slices for the expression of markers altered by drug treatment or cultivation itself. In summary, this protocol describes the generation of murine lung tumor slices and their culture on a rotating incubation unit and demonstrates how slices should be systematically analyzed for the expression of heterogeneous tissue markers, as a prerequisite prior to drug response studies.

We provide a method for the generation, cultivation and systematic analysis of organotypic slices derived from murine lung tumors. We also describe how to optimize for slice thickness, and how to select drug concentrations to treat tumor slices.

Organotypic primary tissue explant cultures, which include precision-cut slices, represent the three-dimensional (3-D) tissue architecture as well as the ...

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, ...

Application of Consistent Massage-Like Perturbations on Mouse Calves and Monitoring the Resulting Intramuscular Pressure Changes

Massage is generally recognized to be beneficial for relieving pain and inflammation. Although previous studies have reported anti-inflammatory effects of massage on skeletal muscles, the molecular mechanisms behind are poorly understood. We have recently developed a simple device to apply local cyclical compression (LCC), which can generate intramuscular pressure waves with varying amplitudes. Using this device, we have demonstrated that LCC modulates inflammatory responses of macrophages in situ and alleviates immobilization-induced muscle atrophy. Here, we describe protocols for the optimization and application of LCC as a massage-like intervention against immobilization-induced inflammation and atrophy of skeletal muscles of mouse hindlimbs. The protocol that we have developed can be useful for investigating the mechanism underlying beneficial effects of physical exercise and massage. Our experimental system provides a prototype of the analytical approach to elucidate the mechanical regulation of muscle homeostasis, although further development needs to be made for more comprehensive studies.

Here we describe the protocols for applying defined mechanical loads to mouse calves and for monitoring the concomitant intramuscular pressure changes. The experimental systems that we have developed can be useful for investigating the mechanism behind the beneficial effects of physical exercise and massage.

Massage is generally recognized to be beneficial for relieving pain and inflammation. Although previous studies have reported anti-inflammatory effects of ...

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 ...