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

Rabies is a hundred percent fatal and rapid, accurate diagnosis is required to get life-saving treatment to those who need it most. Reliable laboratory testing is really crucial for rabies control. The CDC has developed a step-by-step protocol for rabies testing by quantitative reverse transcriptase PCR or RT-PCR, using a validated real-time Alisa virus test called LN34, enabling laboratories to improve diagnostic accuracy and support rabies prevention efforts.

For decades, rabies testing relied on the direct fluorescence antibody tests called DFA or FAT. However, many international laboratories also use RT-PCR for rabies testing due to its excellent sensitivity and accuracy. The DFA test for rabies require specialized microscopes, cold chain storage, specialized training of staff and reagents have experienced shortages over recent years, PCR and RT-PCR are now widely used for pathogen testing.

Many laboratories already have PCR and RT-PCR equipment and expertise and the needed reagents are available from commercial vendors. Cost is a major barrier to rabies testing in areas that have the most human rabies cases. CDC developed a multiplex form of the LN34 assay, which improves quality controls and lowers cost per sample tested.

RT-PCR is increasingly being implemented for rabies diagnostics because it's widely used for testing other pathogens and the results are easy to interpret. But using a validated test is extremely important, and making sure the test is performing as expected is equally critical. Our step-by-step protocol includes quality controls, safety considerations, and PCR best practices to help labs get started.

To begin, wear personal protective equipment including safety glasses, a closed front gown and two pairs of gloves. Clean and disinfect the work surface with quaternary ammonium compound, or another suitable disinfectant for inactivating the rabies virus. Lay out a plastic lined absorbent pad on the work surface, using a clean single use scalpel collect tissue representing a full cross-section of the brainstem and cerebellum.

For homogenization and RNA extraction finally, mince the required tissues using a single use scalpel. Smear the mince tissue with a swab and transfer the swab to a tube prefilled with homogenization buffer and beads. After discarding the outer glove using QAC disinfectant diluted 1:256, clean and disinfect the workstation, equipment, and outside of the sample tubes.

Homogenize samples with a mini bead beater for 60 seconds. And visually inspect the tubes. Allow the samples to sit for five minutes at room temperature.

Finally clean and disinfect the workstation, equipment, and outside of sample tubes with QAC disinfectant before proceeding to RNA extraction. To begin, perform surface decontamination of the biological safety cabinet using QAC diluted to 1:256. Perform additional cleaning to remove dust or other environmental contaminants.

Lay out a plastic lined absorbent work pad on the working platform. Then, arrange the reagents and the samples in the biosafety cabinet. Lay out all collection tubes in a clean rack for micro centrifuge tubes.

Pre-fill one 1.5 milliliter micro centrifuge tube with 300 microliters of 100%ethanol for each brain sample. Centrifuge the homogenized sheet brain samples at 10, 000 to 16, 000 G for two minutes in a tabletop micro centrifuge. Transfer the clear pink supernatant into a new sterile tube containing 100%ethanol, pipette up and down to mix and transfer 600 microliters of the ethanol supernatant mixture to a spin column in a collection tube.

Centrifuge for one minute until the liquid passes through the column, discard the flow through and transfer each column to a new collection tube. Add 400 microliters of RNA pre-wash buffer to each column. Centrifuge the mixture at 10, 000 to 16, 000 G for 30 seconds and discard the flow through.

Return each column to the same collection tube and repeat the pre-wash step a second time. Next, add 700 microliters of RNA wash buffer to each column. Centrifuge at 10, 000 to 16, 000 G for two minutes.

And carefully transfer the into an RNase-free tube. Discard the flow through and the collection tube, then remove and discard outer gloves. Now add 50 microliters of DNase and RNase free water directly to the column matrix and incubate for 30 seconds.

Then centrifuge at 10, 000 to 16, 000 G for one minute. Carefully transfer RNA to a new screw top flat bottom accession labeled micro centrifuge tube for RT-PCR assay. To begin, thaw one step RT-PCR buffer, no template control, nuclease free water, primers, and probes on ice, in the master mix preparation space.

Place one step RT-PCR enzyme on ice. Calculate the volume of each reagent for the LN34 multiplexed and beta A Master Mixes. Designate wells for each sample in triplicate for the LN34 multiplexed assay using a 96 well plate map.

After vortexing and spinning, dispense 23 microliters of LN34 multiplex Master Mix into each LN34 assigned well. To set up negative template control reactions, add two microliters of PCR grade water into each negative template control well. Place the 96 well plate on ice.

Clean the workstation with 70%ethanol. Now at the template edition workspace, thaw RNA samples and positive control single use aliquot of artificial RNA on ice. After vortexing, briefly centrifuge tubes containing RNA samples, pipette two microliters of sample or positive control RNA into the corresponding well.

After adding all samples, place the optical adhesive cover over the wells to seal them completely. Centrifuge the plate at 500 G for one minute at room temperature, place the sealed plate into a real time or quantitative PCR instrument calibrated for fam and VIC Hex reporter dies. Set the instrument to the appropriate cycling parameters.

After the run is completed set the threshold values to 0.2 for LN34 and 0.05 for beta actin. Check the control sample curves for any quality issues. The LN34 assay showed successful amplification curves with positive results crossing the threshold at distinct cycle threshold values when viewed on a logarithmic scale.

And sigmoid amplification curves on a linear scale for both LN34 and beta actin. Negative results exhibited flat lines without amplification. Abnormal amplification curves were identified with some displaying linear rather than sigmoidal increases in fluorescence, indicating atypical results.

The corresponding multi-component plots showed irregular wavy fluorescent signals rather than expected smooth curves. Highlighting the need to analyze amplification plots instead of relying solely on cycle threshold values.