The work was supported in part by FDOH-7ZK15 and NIAID 1R21AI128188-01. Research reported in this publication was supported in part by the National Institutes of Allergy and Infectious Diseases, and in part by Biomedical Research Program of Florida Department of Health. The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH or Florida Department of Health. Dynamic Combinatorial Chemistry LLC is acknowledged for their support and contribution to this project.
Dengue 1 virus (strain BOL-KW010) was kindly provided by the Florida Department of Health Bureau of Laboratories. Zika virus and the Asian lineage of chikungunya virus were graciously provided by the Centers for Disease Control and Prevention. The Indian Ocean lineage of chikungunya virus was kindly provided by Robert Tesh (World Reference Center for Emerging Viruses and Arboviruses, through the University of Texas Medical Branch in Galveston, Texas) to the UF-FMEL. We thank S. Bellamy, B. Eastmond, S. Ortiz, D. Velez, K. Wiggins, R. ZimLer, and K. Zirbel for assistance with the infection studies. We also thank M. S. Kim for providing Q-paper.

NOTE: Mosquitoes were the only animals directly used in this study. The procedures to manage chickens, whose blood was used to feed the infected mosquitoes, were approved as IACUC Protocol #201507682 by the University of Florida Institutional Animal Care and Use Committee.Virus propagation and mosquito infection studies were performed at the BSL-3 facility of the Florida Medical Entomology Laboratory in Vero Beach, FL. RT-LAMP experiments were performed in the BSL-2 laboratory shared by FfAME and Firebird Biomolecular Sciences LLC in Alachua, FL.
1. Design of Primers and Strand Displacing Probes
<ol>
	<li>Extract viral sequences for Dengue 1 from the Broad Institute database23; sequences for Zika and chikungunya from the NIAID Virus Pathogen Database and Analysis Resource24.
	<ol>
		<li>Create multiple sequence alignments (MSAs) for sequences of interest using software MUSCLE v3.8.3125. Use generated MSAs to search for LAMP primers sets within a conserved region of the target, and avoid non-relevant or unintended targets, such as distinctions between subtypes of a target.</li>
	</ol>
	</li>
	<li>Follow the design rules for LAMP primers as described online (http://loopamp.eiken.co.jp/e/lamp/), and allow the use of mixed bases in primers to cover the virus divergence26. To avoid cross-reactivity of primer sets, compare generated LAMP primers to the NCBI RNA virus database using NCBI BLAST27. Compare each set to eliminate primers that would dimerize in a multiplexed assay by using NCBI BLAST as well.</li>
	<li>Screen the double-stranded portion of the strand-displacing probe against any viral genome sequence and mosquito genomic sequence. Modify 5&#39;-ends of target binding probe with fluorescein amidite (FAM), HEX dye, and 5-carboxytetramethylrhodamine (TAMRA) dye for Zika, chikungunya, and Dengue 1, respectively.
	<ol>
		<li>Include a positive control primer set for urine samples. Design LAMP primers to target human mitochondrial DNA. Label strand-displacing probe with TET on 5&#39;-end. Label quencher probes that are partially complementary to each fluorescently labeled probe with quencher (e.g., Iowa Black-FQ) on 3&#39;-end (Table 1).</li>
	</ol>
	</li>
</ol>
2. Virus Isolates and Infected Mosquito Samples
<ol>
	<li>Use isolates of Zika virus (Puerto Rico strain), Chikungunya virus (La R&#233;union or British Virgin Islands strain), and Dengue serotype-1 virus (Key West strain). Determine viral titers using plaque assay28 or quantitative RT-PCR29. Extract viral RNAs using commercially available RNA extraction kits. 
	NOTE: Table 2 represents the viral titers of each virus isolate used in this study.</li>
	<li>Follow the article published by Yaren et al. for a detailed protocol about the infection of Ae. aegypti females with Zika and chikungunya viruses20. See Table 2 for the viral titer of the mosquito sample infected with Zika virus.</li>
</ol>
3. RT-LAMP Coupled with Thermolabile Uracil DNA Glycosylase
<ol>
	<li>10X primer mix preparation.
	<ol>
		<li>Prepare 100 &#181;M stock solution of each primer and probe (see step 1) by adding nuclease free water. Vortex well and store at -20 &#176;C until use.</li>
		<li>For 10X Primer mix for each Zika, Dengue 1, or mitochondrial DNA target, mix 16 &#181;L of FIP, 16 &#181;L BIP, 2 &#181;L of F3, 2 &#181;L of B3, 5 &#181;L of LF, 2 &#181;L of LB, 3 &#181;L of LB-fluorescent labeled probe, 4 &#181;L of quencher probe, and 50 &#181;L of nuclease-free water to give a total of 100 &#181;L volume.</li>
		<li>For 10X Primer mix for Chikungunya, mix 16 &#181;L of FIP, 16 &#181;L BIP, 2 &#181;L of F3, 2 &#181;L of B3, 5 &#181;L of LB, 2 &#181;L of LF, 3 &#181;L of LF-fluorescent labeled probe, 4 &#181;L of quencher probe, and 50 &#181;L of nuclease-free water to give a total of 100 &#181;L volume. 
		NOTE: The probe concentration described above uses 300 nM of final fluorescent probe and 400 nM of quencher probe. Alternatively, use 80 nM of fluorescently labeled probe and 200 nM of its quencher probe for real-time analysis.</li>
	</ol>
	</li>
	<li>Add 5 &#181;L of 10X primer mix (For 3-plex, add 5 &#181;L of each 10X primer mix), 5 &#181;L of 10X isothermal amplification buffer (1X buffer composition: 20 mM Tris-HCl buffer pH 8.8, 50 mM KCl, 10 mM (NH4)2SO4, 8 mM MgSO4, 0.1% Tween-20, 1 mM DTT), 7 &#181;L of dNTP mixture (10 mM of dATP, dCTP and dGTP; 5 mM of dTTP and dUTP), 16 units of DNA Polymerase, 15 units of Reverse Transcriptase, 80 units of recombinant ribonuclease inhibitor, 2 units of thermolabile UDG, 1-2 &#181;L of varying amounts of viral RNAs, and water to bring the total volume up to 50 &#181;L in a 0.25 mL PCR tube (see Table of Materials).</li>
	<li>Include negative control assays at this stage by replacing viral RNA volume with water. Incubate samples between 65-68 &#176;C for 45 min to 1 h, then analyze by running 5 &#181;L of each sample on 2.5% agarose gel in 1X TBE buffer containing ethidium bromide (0.4 &#181;g/mL). Use a 25 bp or 50 bp DNA ladder as marker on the gel. 
	NOTE: Addition of a non-target specific template is optional.</li>
	<li>To detect the presence of pathogenic RNA, test each primer set and its no-template control by varying the temperature and/or magnesium concentration, since each RT-LAMP primer set was designed to work in varying temperatures (65 &#176;C to 68 &#176;C) or magnesium concentrations (6 to 10 mM final). Prepare the experiments at room temperature.</li>
</ol>
4. RT-LAMP Using Viral RNA-Spiked Urine
<ol>
	<li>Test RT-LAMP primers in varying final concentrations of urine (0%, 10%, 20%, and 50%) in 50 &#181;L of total reaction volume. For 10% final urine concentration, add 1 &#181;L of viral RNA to 5 &#181;L of urine. For 20% final urine concentration, add 1 &#181;L of viral RNA to 10 &#181;L of urine. For 50% final urine concentration, add 1 &#181;L of viral RNA to 25 &#181;L of urine.</li>
	<li>For real-time monitoring of RT-LAMP in 10% of urine, use a real-time PCR instrument (see Table of Materials) with different fluorophore filters.
	<ol>
		<li>To read fluorescence signals from FAM-labeled amplicons for Zika, use a filter ranging from 483 to 533 nm; for HEX-labeled amplicons for chikungunya, use a filter ranging from 523 to 568 nm; for TAMRA-labeled amplicons for Dengue 1, use a filter ranging from 558 to 610 nm; and for TET-labeled amplicons for mitochondrial DNA, use a filter ranging from 523 to 568 nm. 
		NOTE: FAM has excitation and emission maxima of 495 nm and 520 nm, respectively. HEX has excitation and emission maxima of 538 nm and 555 nm, respectively. TAMRA has excitation and emission maxima of 559 nm and 583 nm, respectively. TET has excitation and emission maxima of 522 nm and 539 nm, respectively.</li>
	</ol>
	</li>
	<li>Use 96-well plates and seal the plate with a clear plastic sheet, then incubate samples at 65-68 &#176;C for 60-90 min and record the fluorescence every 30 s using the light cycler. Initially use 80 nM of fluorescently labeled probes, and then test the 300 nM of fluorescently labeled probes.
	<ol>
		<li>Determine limit of detection for each target by adding serially diluted viral RNAs in final urine concentration of 10%. 
		NOTE: Use real-time RT-LAMP to determine optimum temperature, magnesium concentration, fluorescently labeled probe concentration, and reaction time.</li>
	</ol>
	</li>
	<li>To visualize the fluorescence generated by strand displaceable probes by eye, use a blue LED light source from a light cycler with excitation at 470 nm (any gel electrophoresis box with built-in blue light source will work, see table of materials), and record the image with a cell phone camera at room temperature in the dark. 
	NOTE: Use 300 nM of fluorescently labeled probes, when visualization of all three colors by eye using blue LED is needed.</li>
</ol>
5. RT-LAMP on Detection of Zika-Infected Mosquitoes Using Q-Paper Technology
<ol>
	<li>Prepare quaternary ammonium-modified filter paper (Q-paper), according to previously published methods with slight modifications30.
	<ol>
		<li>Immerse 1 g of cellulose filter paper circles (see Table of Materials) with diameters of 3.5 cm in 50 mL of 1.8% of aqueous NaOH solution for 15 min for activation.</li>
		<li>Collect activated paper circles by vacuum filtration and then immerse them in 40 mL of aqueous solution of (2,3-epoxypropyl) trimethylammonium chloride (0.28 g) overnight at room temperature. 
		NOTE: Keep the mass ratio of (2,3-epoxypropyl) trimethylammonium chloride to filter paper at 0.28 to 1.</li>
		<li>Collect the resulting cationic paper by vacuum filtration, and neutralize with 50 mL of 1% of acetic acid.</li>
		<li>Wash the paper three times with ethanol and air-dry under a hood with constant airflow.</li>
	</ol>
	</li>
	<li>Cut Q-paper sheets into small rectangles (3 mm x 4 mm) using a paper punch or scissors. Crush Ae. aegypti female mosquitoes potentially infected with Zika on each paper with a micro pestle.</li>
	<li>Add to each paper 20 &#181;L of 1 M aqueous ammonia solution (pH &#8776; 12) and wait for 5 min. Then wash each paper once with 20 &#181;L of 50% EtOH and once with 20 &#181;L of nuclease-free water. Air-dry the paper in BSL-2 biosafety cabinet for about 1 h. 
	NOTE: At this point, samples can be stored at -20 &#176;C overnight until use.</li>
	<li>Using tweezers, place each paper in 100 &#181;L of RT-LAMP reaction mixture and incubate at 65-68 &#176;C for 45 min. Make sure to fully submerge the paper in the solution; it should not be floating. Read and record the fluorescence arising from Zika virus using blue LED light source and orange or yellow filter.</li>
</ol>
6. Lyophilization of RT-LAMP Reagents for 100 &#181;L of Reaction Volume
<ol>
	<li>Prepare 10X LAMP primer mixture according to section 3.1, and prepare dNTP mixture containing dUTP according to section 3.2. Store primer mixture and dNTPs at -20 &#176;C until use.</li>
	<li>Prepare 10 mL of enzyme dialysis buffer to remove glycerol, by mixing 100 &#181;L of 1 M Tris-HCl pH 7.5, 500 &#181;L of 1 M KCl, 2 &#181;L of 0.5 M EDTA, 100 &#181;L of 10% Triton X-100, 100 &#181;L of 0.1 M DTT and 9.198 mL of nuclease-free water. Store the dialysis buffer at 4 &#176;C. 
	NOTE: Make sure to filter (0.2-micron filters) all buffer reagent solutions prior to use and store them at 4 &#176;C.
	<ol>
		<li>Use an ultrafiltration membrane with 10 kDa cut-off limit (see Table of Materials). Place 350 &#181;L of dialysis buffer, 4 &#181;L of 32 units of DNA Polymerase, 2 &#181;L of 30 units of Reverse Transcriptase, and 2 &#181;L of 80 units of RNase inhibitor into ultrafiltration membrane and centrifuge at 14,000 x g for 5 min at 4 &#176;C.</li>
		<li>Add another 350 &#181;L of dialysis buffer to the ultrafiltration membrane and centrifuge (14,000 x g) for another 5 min. Empty the collection tube and repeat this step, then centrifuge (14,000 x g) for 3 min.</li>
		<li>For elution, invert the membrane and place in a new collection tube. Spin at 1,000 x g for 2 min. Measure the elution volume; if less than 8 &#181;L, bring the volume up to 8 &#181;L by adding dialysis buffer. Store the elution at 4 &#176;C, and immediately proceed to next step. 
		NOTE: This protocol is for single tube lyophilization, but prepare at least 10 reaction tubes at a time using the same ultrafiltration membrane and use the same amount of dialysis buffer.</li>
	</ol>
	</li>
	<li>Mix 10 &#181;L of 10X LAMP primer if singleplex (for 3-plex, add 10 &#181;L of each 10X primer mix), 14 &#181;L of dNTP mixture, 8 &#181;L of dialyzed enzyme mix, 2 &#181;L of 2 units of thermolabile UDG, and 66 &#181;L of nuclease free water (for 3-plex, use 46 &#181;L) in a 0.5 mL microcentrifuge tube and leave the lid open. Instead, add another lid punctured with a needle to allow vapor to escape.</li>
	<li>Immediately freeze the tubes at -80 &#176;C for 2 h, and lyophilize the tube for 4 h. Cover the lyophilization chamber with aluminum foil for protection from light. When reagents are dried, remove the punctured lids, close the original lid, and store the tubes with lyophilized reagents at 4 &#176;C in the dark. 
	NOTE: Reagents can be lyophilized overnight, if necessary.</li>
</ol>
7. Testing Lyophilized RT-LAMP Reagents on Urine and Mosquito Samples Containing Zika
<ol>
	<li>Use the following items from the kit (see the Table of Materials) for Zika: A 3D-printed observing box with orange filter (long pass filter, cut-on ~540 nm), 2 AAA batteries for observing box, lyophilized reaction tubes labeled ZV for Zika detection, and 1.1X rehydration Buffer in 1.5 mL screw top tubes.</li>
	<li>Prepare 50 mL of 1.1X rehydration buffer by mixing 5.5 mL of 10X isothermal amplification buffer that comes along with DNA Polymerase (see the Table of Materials), 3.3 mL of 100 mM MgSO4, 110 &#181;L of 0.5 M DTT, and 41.09 mL of nuclease-free water. Mix well, aliquot 1 mL of this solution to 1.5 mL screw top tubes, and store at 4 &#176;C until use.</li>
	<li>Add 90 &#181;L of 1.1X rehydration buffer to each reaction tube (0.5 mL). Ensure that the liquid fills the bottom of the tube containing the lyophilized reagents (dissolve them with mixing by shaking, then briefly spin down (1,000 x g)). Add 10 &#181;L of urine sample spiked with viral RNA to the reaction tubes (see section 4.1 for details).
	<ol>
		<li>If testing mosquitoes, add a rectangle of Q-paper holding mosquito carcasses (as prepared in the steps described in sections 5.2 and 5.3), along with 10 &#181;L purified water instead of urine samples. 
		NOTE: Alternatively, add 10 &#181;L of saliva or serum samples instead of urine. If the samples are extracted DNA or RNA, add 2-5 &#181;L of the sample into rehydrated mixture and complete with nuclease-free water to 10 &#181;L of final sample volume.</li>
	</ol>
	</li>
	<li>Close the lid of the reaction tubes. Place them in a heat block/bath (or incubator) pre-set at 65-68 &#176;C. Incubate for 30-45 min, but no more than 1 h. After incubation, remove the tube from the heat. To prevent contamination of future assays, ensure that these tubes remain closed.</li>
	<li>Place reaction tubes in the observing box; the temperature will fall to the ambient temperature (preferably 25 &#176;C). Turn on the switch on the back of the observing box. Observe sample through the orange filter and capture the image by a cell phone camera which is held at a distance where the image fills 80% of the field of view. 
	NOTE: Better visualization is achieved in low light environments.</li>
	<li>After visualization is completed, turn off the switch. Store the sealed tubes in the dark, preferably with refrigeration, if later visualization is needed.</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Basic+local+alignment+search+tool.">Basic local alignment search tool.</a> J Mol Biol. 215, (1990).</li><li>Baer, A., Kehn-Hall, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Viral+Concentration+Determination+Through+Plaque+Assays:+Using+Traditional+and+Novel+Overlay+Systems.">Viral Concentration Determination Through Plaque Assays: Using Traditional and Novel Overlay Systems.</a> Journal of visualized experiments: JoVE. (93), e52065 (2014).</li><li>Reiskind, M. H., Pesko, K., Westbrook, C. J., Mores, C. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Susceptibility+of+Florida+Mosquitoes+to+Infection+with+Chikungunya+Virus.">Susceptibility of Florida Mosquitoes to Infection with Chikungunya Virus.</a> The American journal of tropical medicine and hygiene. 78 (3), 422-425 (2008).</li><li>Glushakova, L. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+chikungunya+viral+RNA+in+mosquito+bodies+on+cationic+(Q)+paper+based+on+innovations+in+synthetic+biology.">Detection of chikungunya viral RNA in mosquito bodies on cationic (Q) paper based on innovations in synthetic biology.</a> Journal of Virological Methods. 246, 104-111 (2017).</li><li>Hsieh, K., Mage, P. L., Csordas, A. T., Eisenstein, M., Tom Soh, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+elimination+of+carryover+contamination+and+detection+of+DNA+with+uracil-DNA-glycosylase-supplemented+loop-mediated+isothermal+amplification+(UDG-LAMP).">Simultaneous elimination of carryover contamination and detection of DNA with uracil-DNA-glycosylase-supplemented loop-mediated isothermal amplification (UDG-LAMP).</a> Chemical Communications. 50 (28), 3747-3749 (2014).</li><li>Tang, Y., Chen, H., Diao, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advanced+uracil+DNA+glycosylase-supplemented+real-time+reverse+transcription+loop-mediated+isothermal+amplification+(UDG-rRT-LAMP)+method+for+universal+and+specific+detection+of+Tembusu+virus.">Advanced uracil DNA glycosylase-supplemented real-time reverse transcription loop-mediated isothermal amplification (UDG-rRT-LAMP) method for universal and specific detection of Tembusu virus.</a> Sci Rep. 6, 27605 (2016).</li><li>Thomas, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+of+pterin,+6-formylpterin,+6-carboxypterin+and+folic+acid+in+aqueous+solution:+pH+effects.">Fluorescence of pterin, 6-formylpterin, 6-carboxypterin and folic acid in aqueous solution: pH effects.</a> Photochem Photobiol Sci. 1, (2002).</li><li>Wu, D., Lehane, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pteridine+fluorescence+for+age+determination+of+anopheles+mosquitoes.">Pteridine fluorescence for age determination of anopheles mosquitoes.</a> Med Vet Entomol. 13, (1999).</li><li>Janis, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inactivation+and+environmental+stability+of+Zika+virus.">Inactivation and environmental stability of Zika virus.</a> Emerg Infect Dis J. 22, (2016).</li><li>Poloni, T. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+dengue+virus+in+saliva+and+urine+by+real+time+RT-PCR.">Detection of dengue virus in saliva and urine by real time RT-PCR.</a> Virology Journal. 7 (1), 22 (2010).</li><li>Jones, P., Okeoma, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+Chikungunya+virus+(CHIKV)+in+urine+of+infected+mice:+a+potential+non-invasive+diagnostic+tool+for+CHIKV.">Detection of Chikungunya virus (CHIKV) in urine of infected mice: a potential non-invasive diagnostic tool for CHIKV.</a> J Infect Dis Ther. 3 (4), 1000226 (2015).</li><li>Glushakova, L. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-throughput+multiplexed+xMAP+Luminex+array+panel+for+detection+of+twenty+two+medically+important+mosquito-borne+arboviruses+based+on+innovations+in+synthetic+biology.">High-throughput multiplexed xMAP Luminex array panel for detection of twenty two medically important mosquito-borne arboviruses based on innovations in synthetic biology.</a> J. Virol. Methods. 214, 60-74 (2015).</li><li>Yaren, O., Benner, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Loop+Mediated+Amplifications+with+Nucleoside+Analogs.">Loop Mediated Amplifications with Nucleoside Analogs.</a> US Provisional patent. , (2016).</li></ol>

Several of the authors and their institutions own intellectual property associated with this assay.

Mosquito-borne viruses including Zika, chikungunya, and dengue threaten the public health and recent Zika outbreaks highlight the need for low-cost point-of-care detection alternatives for patient diagnostics, as well as for mosquito surveillance. Isothermal amplification methods were developed as affordable alternatives to PCR-based systems. Particularly, RT-LAMP-based platforms have been applied to detect a wide range of pathogens. However, the use of isothermal platforms has been mainly limited to single target detection. The method reported here uses a modified RT-LAMP to allow simultaneous detection of three different targets in a single reaction mixture, which runs at a range of moderate temperatures (65-68 &#176;C), proven to be especially convenient because temperatures higher than 60 &#176;C render Zika and other viruses non-infectious30,35.
Further, data presented here shows that RT-LAMP is capable of tolerating inhibiting substances that might be present in various biological fluids15. This allows viral RNAs to be amplified without an extra step of RNA purification by directly adding the sample into the reaction mixture. Depending on the type of sample, the maximum volume of sample allowed needs to be determined as to not inhibit the reaction and not cause background fluorescence (false positive). The study presented here focuses on the use of 10% final volume for urine samples. However, saliva and serum samples can also be applied to the assay in the same manner20. Limits of detection were found to be well above the reported viral loads in patient urine13, as well as virus-infected mosquitoes20,36,37.
Mosquito surveillance usually has a lower priority for public health resources, so an inexpensive multiplexed kit to survey a household or a public area would have special value. Therefore, this kit was modified by the addition of Q-paper to allow the detection of viruses in mosquito carcasses. Q-paper contains high levels of covalently attached quaternary ammonium groups, which allows the capture of viral nucleic acids on its surface through coulombic interactions. Even though it was a concern that Q-paper might inhibit RT-LAMP, it was found that by adding Q-paper carrying mosquito carcasses directly into RT-LAMP mixture, successful virus detection could be achieved within 30 min. For successful signal generation, Q-paper needs to be small enough (e.g., sized as 3 x 4 mm for 100 &#181;L reaction volume) to be submerged in the RT-LAMP reaction mixture. If it is too big for the reaction tube, or starts floating instead of remaining immersed in the liquid, the reaction may not take place and results may appear as false negatives.
For successful detection and differentiation of each virus, each primer set needs to follow the guidelines for RT-LAMP primer design, the design rules for displacing probes, and include a choice of fluorophores for multiplexing. Each primer set needs to be screened against different targets to avoid cross-reactivity. Additionally, magnesium concentration and incubation temperature need to be optimized for each set. Methods presented here use fluorescence readouts visible by eye, which was accomplished by introducing strand displaceable probes to RT-LAMP architecture. Three distinct colors can be visualized in a multiplexed assay when different fluorophores were assigned to each target virus. This provides advantages over existing isothermal Zika detection methods, since most of the published methods rely on single target detection and the use of intercalating or fluorescence dyes, which are not sequence-specific and are prone to generating non-specific amplicons.
A critical step to consider is the determination of the final concentration of fluorescently labeled displacing probes. It was found that 80 to 100 nM of displacing probes could generate fluorescence signals within 15-20 min, whereas time to signal was delayed when 300 nM of probe concentration was used. Reaction time was further delayed when the assays were run in a multiplexed fashion. The reason for switching from 80 nM to 300 nM was to enable visualization of all three fluorophores using a single LED source. Blue LED with excitation at 470 nm is suitable for FAM-labeled probes, but is less suitable for HEX- or TAMRA-labeled probes. Therefore, a sacrifice was made on incubation time to be able to visualize all three targets.
One of the problems with RT-LAMP is forward contamination. RT-LAMP generates large amounts of amplicon in a short incubation time, and amplicons could be easily released as aerosols. To mitigate this problem, dUTP was included along with other dNTPs followed by UDG digestion during assay set-up. It is important to use a heat-labile version of UDG, since it is necessary to inactivate UDG to prevent the digestion of newly generated RT-LAMP amplicons.
To allow the distribution of the kit without refrigeration, all the components of the assay needed to be lyophilized and the kit supplemented with a rehydration buffer. Therefore, any glycerol present in commercially available enzymes was removed by ultrafiltration because glycerol cannot be removed by lyophilization. At a high concentration in a mixture, glycerol may interfere with enzyme activity. The only enzyme not included in the glycerol removal process was the thermolabile UDG. UDG is a small enzyme with molecular weight of 24.6 kDa, making it unsuitable for ultrafiltration experiments. Therefore, it was added to the lyophilization mixture as purchased. The residual glycerol did not negatively affect the assay&#39;s outcome.
One of the limitations of this fluorescent RT-LAMP technique is the use of the single LED source. When assays were run in singleplex or multiplex format, higher concentrations of displacing probes were needed for signal visualization of all targets. However, this change causes delays in time-to-signal generation. One possible way to overcome this could be the use of two LED sources to better accommodate the other fluorophores (HEX and TAMRA). This way, lower probe concentration could be used to generate a signal with shorter incubation times.
Another limitation that needs consideration is judging the fluorescence color by eye in a dimly lit environment. It is easier if a single target is present in a singleplex or multiplex format, since primers were designed not to interact with each other. The assay would be harder to interpret if multiple targets were present in a single assay tube, such as a pool of mosquitoes or a patient with more than one viral infection. This would require a change of read-out architecture, such as a digital display rather than judging by eye.
A possible future approach would be to increase the level of multiplexing by creating a larger panel of mosquito-borne arboviruses. This requires careful primer design to avoid primer-primer interactions, and may require the use of modified nucleic acid analogs such as self-avoiding molecular recognition systems (SAMRS) and artificially expanded genetic information system (AEGIS) to better accommodate high level of multiplexing38. Accordingly, the solution for signal read-out in a higher multiplex system would be to use a single fluorophore for each displacing probe, and to assign a unique displaced sequence for each target and capture the displaced probes on a solid surface by DNA hybridization. This would create an array-formatted platform with a single fluorophore and LED to excite, but judging the presence of each target would be based on its position in the array39.

Mosquito-borne virus infections, including dengue, chikungunya, and Zika viruses are on the rise and demand immediate management strategies. Dengue and chikungunya viruses are already endemic in many of the tropical regions where Zika is now spreading in the Western Hemisphere1. Zika virus, like dengue, is a member of the Flaviviridae family and is native to Africa with one Asian and two African genetic lineages2. Even though the identification of the Zika virus dates back to 1947, Zika infection in humans remained sporadic for a half century before emerging in the Pacific and the Americas. The first reported outbreak of Zika fever occurred on the island of Yap in the Federated States of Micronesia in 2007, followed by French Polynesia in 2013 and 2014. The first major outbreak in the Americas occurred in 2015 in Brazil.
Zika, chikungunya, and dengue viruses are primarily transmitted by Aedes aegypti and Aedes albopictus. However, Zika has additional downstream human-to-human transmission possibilities, likely being spread through sexual contact, mother-to-fetus interaction, and via breast-feeding3,4,5. Zika fever was first believed to cause only mild illness. However, it was later associated with Guillain-Barr&#233; syndrome in adults, microcephaly in neonates, and chronic musculoskeletal diseases that may last months to years. Diagnosis of Zika illness can be challenging, since the symptoms of a Zika infection are similar to those of other mosquito-spread viruses6. Common co-infections of these viruses make differential diagnosis even more challenging7,8. Therefore, rapid and reliable detection of the nucleic acids from Zika and other viruses is needed to understand epidemiology in real time, to initiate control and preventive measures, and to manage patient care9.
Current diagnostic tests for these viruses include serological tests, virus isolation, virus sequencing, and reverse-transcription PCR (RT-PCR). Standard serological approaches often suffer from inadequate sensitivity and results can be complicated by cross-reactivity in patients who have previously been infected by other flaviviruses.
Therefore, nucleic acid testing remains the most reliable way to detect and differentiate these viruses. Detection of Zika and other mosquito-borne viruses is usually performed using RT-PCR or real-time RT-PCR in variety of biological fluids, such as serum, urine, saliva, semen, breast milk, and cerebral fluid10,11. Urine and saliva samples are generally preferred over blood, since they exhibit less PCR-inhibition, higher viral loads, virus presence for longer periods of time, and increased ease of collection and handling12,13. RT-PCR-based diagnostic tests, however, comprise extensive sample preparation steps and expensive thermal cycling equipment, making it less optimal for the point-of-care.
Reverse transcription loop-mediated isothermal amplification (RT-LAMP) has emerged as a powerful RT-PCR alternative due to its high sensitivity and specificity14, its tolerance for inhibitory substances in biological samples15, and operation on single temperature, which significantly lowers assay complexity and associated costs, making it suitable for low resource environments. RT-LAMP, as it is classically implemented, comprises six primers that bind to eight distinct regions within the target RNA. It runs at constant temperatures between 60 &#176;C and 70 &#176;C, and uses a reverse transcriptase and a DNA polymerase with strong strand displacing activity.
During the initial stages of RT-LAMP, forward and backward internal primers (FIP and BIP, Figure 1A) along with outer forward and backward primers (F3 and B3) form a dumbbell structure, the seed structure of exponential LAMP amplification. Amplification is further accelerated by the loop forward and backward primers (LF and LB), which are designed to bind the single stranded regions of the dumbbell, and results in the formation of concatemers with multiple repeating loops16. Classical LAMP assays based on turbidity or readout by DNA intercalating dyes is not entirely suited for point-of-care detection of Zika, where some level of multiplexing is desired17,18,19. Multiplexing is not easily obtained in these systems, as they are prone to generate false-positives due to off-target amplifications.
To manage these issues, the literature adds an additional component in the form of a &#34;strand-displacing probe&#34; to the classical RT-LAMP architecture20,21,22. Each probe has a sequence-specific double-strand region and a single-stranded priming region. The probe with the single-stranded region is tagged with a 5&#39;-fluorophore, and the complementary probe is modified with a 3&#39;-end quencher. In the absence of a target, no fluorescence is observed due to hybridization of the complementary probe strands, which brings the fluorophore and quencher into close proximity. In the presence of a target, the single-stranded portion of the fluorescent probe binds to its complement on the target, and is then extended by a strand displacing polymerase. Further polymerase extension by reverse primers causes the separation of the quencher strand from its complementary fluorescently labeled strand, allowing emission of fluorescence (Figure 1B). With this design, the signal is generated after the dumbbell formation, reducing the chances of false-positive signals.
The double-stranded portion of the strand-displacing probe can be any sequence, and when multiplexing is applied, the same sequence may be used with different fluorophore-quencher pairs. With this architecture, virus-contaminated urine, serum, or mosquito samples squished on paper were directly introduced to the assay without sample preparation. Three-color fluorescence read-outs visible to human eye were generated within 30 - 45 min, and signals were visualized by a 3D-printed observation box that uses a blue LED and an orange filter. Freeze-drying the RT-LAMP reagents enabled deployment of this kit to lower resource settings without a need for refrigeration.

<table><tbody><tr><td>SafeBlue Illuminator/ Electrophoresis System, MBE-150-PLUS</td><td>Major Science</td><td>MBE-150</td><td>Gel electrophoresis</td></tr><tr><td>G:BOX F3</td><td>Syngene</td><td>G:BOX F3</td><td>Gel imaging</td></tr><tr><td>LightCycler 480 Instrument II, 96-well</td><td>Roche Applied Science</td><td>05 015 278 001</td><td>Real-time PCR</td></tr><tr><td>Amicon Ultra-0.5 Centrifugal Filter Unit with Ultracel-10 membrane</td><td>Millipore Sigma</td><td>UFC501096</td><td>ultrafiltraton membrane for dialysis</td></tr><tr><td>Eppendorf 5417C Centrifuge</td><td>Marshall Scientific</td><td>EP-5417C</td><td>centrifuge</td></tr><tr><td>Myblock Mini Drybath</td><td>Benchmark Scientific</td><td>BSH200</td><td>drybath</td></tr><tr><td>FreeZone Plus 6 Liter Cascade Console Freeze Dry System</td><td>Labconco</td><td>7934020</td><td>lyophilizer</td></tr><tr><td>all priers and probes</td><td>IDT</td><td>custom</td><td>RT-LAMP primers and probes</td></tr><tr><td>dNTP set</td><td>Bioline</td><td>BIO-39049</td><td></td></tr><tr><td>Deoxyuridine Triphosphate (dUTP)</td><td>Promega</td><td>U1191</td><td></td></tr><tr><td>Bst&amp;nbsp;2.0 WarmStart&amp;nbsp;DNA Polymerase</td><td>New England Biolabs</td><td>M0538L</td><td>enzyme</td></tr><tr><td>WarmStart RTx Reverse Transcriptase</td><td>New England Biolabs</td><td>M0380L</td><td>enzyme</td></tr><tr><td>RNase Inhibitor, Murine</td><td>New England Biolabs</td><td>M0314L</td><td>enzyme</td></tr><tr><td>Antarctic Thermolabile UDG</td><td>New England Biolabs</td><td>M0372L</td><td>enzyme</td></tr><tr><td>50bp DNA Step Ladder</td><td>Promega</td><td>G4521</td><td>marker</td></tr><tr><td>LightCycler 480 Multiwell Plate 96, white</td><td>Roche Applied Science</td><td>4729692001</td><td>real-time RT-LAMP analysis</td></tr></tbody></table>

multiplexed isothermal amplification based diagnostic platform to detect zika, chikungunya, and dengue 1

Zika, dengue, and chikungunya viruses are transmitted by mosquitoes, causing diseases with similar patient symptoms. However, they have different downstream patient-to-patient transmission potentials, and require very different patient treatments. Thus, recent Zika outbreaks make it urgent to develop tools that rapidly discriminate these viruses in patients and trapped mosquitoes, to select the correct patient treatment, and to understand and manage their epidemiology in real time.
Unfortunately, current diagnostic tests, including those receiving 2016 emergency use authorizations and fast-track status, detect viral RNA by reverse transcription polymerase chain reaction (RT-PCR), which requires instrumentation, trained users, and considerable sample preparation. Thus, they must be sent to &#34;approved&#34; reference laboratories, requiring time. Indeed, in August 2016, the Center for Disease Control (CDC) was asking pregnant women who had been bitten by a mosquito and developed a Zika-indicating rash to wait an unacceptable 2 to 4 weeks before learning whether they were infected. We very much need tests that can be done on site, with few resources, and by trained but not necessarily licensed personnel.
This video demonstrates an assay that meets these specifications, working with urine or serum (for patients) or crushed mosquito carcasses (for environmental surveillance), all without much sample preparation. Mosquito carcasses are captured on paper carrying quaternary ammonium groups (Q-paper) followed by ammonia treatment to manage biohazards. These are then directly, without RNA isolation, put into assay tubes containing freeze-dried reagents that need no chain of refrigeration. A modified form of reverse transcription loop-mediated isothermal amplification with target-specific fluorescently tagged displaceable probes produces readout, in 30 min, as a three-color fluorescence signal. This is visualized with a handheld, battery-powered device with an orange filter. Forward contamination is prevented with sealed tubes, and the use of thermolabile uracil DNA glycosylase (UDG) in the presence of dUTP in the amplification mixture.

Watch this Scientific Journal Video about Multiplexed Isothermal Amplification Based Diagnostic Platform to Detect Zika, Chikungunya, and Dengue 1 at JoVE.com

Multiplexed Isothermal Amplification Based Diagnostic Platform to Detect Zika, Chikungunya, and Dengue 1

Current multiplexed diagnostics to detect Zika, chikungunya, and dengue viruses require complex sample preparation and expensive instrumentation, and are difficult to use in low resource environments. We show a diagnostic that uses isothermal amplification with target-specific strand displaceable probes to detect and differentiate these viruses with high sensitivity and specificity.

Zika, dengue, and chikungunya viruses are transmitted by mosquitoes, causing diseases with similar patient symptoms. However, they have different ...

multiplexed-isothermal-amplification-based-diagnostic-platform-to

Nanyang Technological University

Research

JoVE Journal

Biochemistry

14.2K Views.  Foundation for Applied Molecular Evolution (FfAME). Current multiplexed diagnostics to detect Zika, chikungunya, and dengue viruses require complex sample preparation and expensive instrumentation, and are difficult to use in low resource environments. We show a diagnostic that uses isothermal amplification with target-specific strand displaceable probes to detect and differentiate these viruses with high sensitivity and specificity.

Video: Multiplexed Isothermal Amplification Based Diagnostic Platform to Detect Zika, Chikungunya, and Dengue 1

Reverse Transcription-Loop-mediated Isothermal Amplification (RT-LAMP) Assay for Zika Virus and Housekeeping Genes in Urine, Serum, and Mosquito Samples

Infection with Zika virus (ZIKV) can be asymptomatic in adults, however, infection during pregnancy can lead to miscarriage and severe neurological birth defects. The goal of this protocol is to quickly detect ZIKV in both human and mosquito samples. The current gold standard for ZIKV detection is quantitative reverse transcription PCR (qRT-PCR); reverse transcription loop-mediated isothermal amplification (RT-LAMP) may allow for a more efficient and low-cost testing without the need for expensive equipment. In this study, RT-LAMP is used for ZIKV detection in various biological samples within 30 min, without first isolating the RNA from the sample. This technique is demonstrated using ZIKV infected patient urine and serum, and infected mosquito samples. 18S ribosomal ribonucleic acid and actin are used as controls in human and mosquito samples, respectively.

Reverse Transcription-Loop-mediated Isothermal Amplification RT-LAMP Assay for Zika Virus and Housekeeping Genes in Urine, Serum, and Mosquito Samples

This protocol provides an efficient and low-cost method to detect Zika virus or control targets in human urine and serum samples or in mosquitoes by reverse transcription loop-mediated isothermal amplification (RT-LAMP). This method does not require RNA isolation and can be done within 30 min.

Infection with Zika virus (ZIKV) can be asymptomatic in adults, however, infection during pregnancy can lead to miscarriage and severe neurological birth ...

Genetics

Visual Detection of Multiple Nucleic Acids in a Capillary Array

Multi-target, short time, and resource-affordable methodologies for the detection of multiple nucleic acids in a single, easy to operate test are urgently needed in disease diagnosis, microbial monitoring, genetically modified organism (GMO) detection, and forensic analysis. We have previously described the platform called CALM (Capillary Array-based Loop-mediated isothermal amplification for Multiplex visual detection of nucleic acids). Herein, we describe improved fabrication and performance processes for this platform. Here, we apply a small, ready-to-use cassette assembled by capillary array for multiplex visual detection of nucleic acids. The capillary array is pre-treated into a hydrophobic and hydrophilic pattern before fixing loop-mediated isothermal amplification (LAMP) primer sets in capillaries. After assembly of the loading adaptor, LAMP reaction mixture is loaded and isolated into each capillary, due to capillary force by a single pipetting step. The LAMP reactions are performed in parallel in the capillaries. The results are visually read out by illumination with a hand-held UV flashlight. Using this platform, we demonstrate monitoring of 8 frequently appearing elements and genes in GMO samples with high specificity and sensitivity. In summary, the platform described herein is intended to facilitate the detection of multiple nucleic acids. We believe it will be widely applicable in fields where high-throughput nucleic acid analysis is required.

This protocol describes the fabrication of a small, ready-to-use cassette that can be applied for visual detection of multiple nucleic acids in a single, test that is easy to operate. In this approach, a capillary array was used for multiplex and highly efficient detection of GMO targets.

Multi-target, short time, and resource-affordable methodologies for the detection of multiple nucleic acids in a single, easy to operate test are urgently ...

Bioengineering

Zika Virus Specific Diagnostic Epitope Discovery

High-density peptide microarrays allow screening of more than six thousand peptides on a single standard microscopy slide. This method can be applied for drug discovery, therapeutic target identification, and developing of diagnostics. Here, we present a protocol to discover specific Zika virus (ZIKV) diagnostic peptides using a high-density peptide microarray. A human serum sample validated for ZIKV infection was incubated with a high-density peptide microarray containing the entire ZIKV protein translated into 3,423 unique 15 linear amino acid (aa) residues with a 14-aa residue overlap printed in duplicate. Staining with different secondary antibodies within the same array, we detected peptides that bind to Immunoglobulin M (IgM) and Immunoglobulin G (IgG) antibodies present in serum. These peptides were selected for further validation experiments. In this protocol, we describe the strategy followed to design, process, and analyze a high-density peptide microarray.

In this protocol, we describe a technique to discover Zika virus specific diagnostic peptides using a high-density peptide microarray. This protocol can readly be adapted for other emerging infectious diseases.

High-density peptide microarrays allow screening of more than six thousand peptides on a single standard microscopy slide. This method can be applied for ...

Immunology and Infection

A Simple Flow Cytometry Based Assay to Determine In Vitro Antibody Dependent Enhancement of Dengue Virus Using Zika Virus Convalescent Serum

Antibody dependent enhancement of infection has been shown to play a major role in Dengue viral pathogenesis. Traditional assays that measure the capacity of antibodies or serum to enhance infection in impermissible cell lines have relied on using viral output in the media followed by plaque assays to quantify infection. More recently, these assays have examined Dengue virus (DENV) infection in the cell lines using fluorescently labeled antibodies. Both these approaches have limitations that restrict the widespread use of these techniques. Here, we describe a simple in vitro assay using Dengue virus reporter viral particles (RVPs) that express green fluorescent protein and K562 cells to examine antibody dependent enhancement (ADE) of DENV infection using serum that was obtained from rhesus macaques 16 weeks after infection with Zika virus (ZIKV). This technique is reliable, involves minimal manipulation of cells, does not involve the use of live replication competent virus, and can be performed in a high throughput format to get a quantitative readout using flow cytometry. Additionally, this assay can be easily adapted to examine antibody dependent enhancement (ADE) of other flavivirus infections such as Yellow Fever virus (YFV), Japanese Equine Encephalitis virus (JEEV), West Nile virus (WNV) etc. where RVPs are available. The ease of setting up the assay, analyzing the data, and interpreting results makes it highly amenable to most laboratory settings.

A Simple Flow Cytometry Based Assay to Determine In Vitro Antibody Dependent Enhancement of Dengue Virus Using Zika Virus Convalescent Serum

We describe a simple and easy protocol for measuring antibody dependent enhancement of infection by Zika virus convalescent serum using Dengue virus Reporter Viral Particles.

Antibody dependent enhancement of infection has been shown to play a major role in Dengue viral pathogenesis. Traditional assays that measure the capacity ...

Quantification of Antibody-dependent Enhancement of the Zika Virus in Primary Human Cells

The recent emergence of the flavivirus Zika and neurological complications, such as Guillain-Barr&#233; syndrome and microcephaly in infants, has brought serious public safety concerns. Among the risk factors, antibody-dependent enhancement (ADE) poses the most significant threat, as the recent re-emergence of the Zika virus (ZIKV) is primarily in areas where the population has been exposed and is in a state of pre-immunity to other closely related flaviviruses, especially dengue virus (DENV). Here, we describe a protocol for quantifying the effect of human serum antibodies against DENV on ZIKV infection in primary human cells or cell lines.

We describe a method to evaluate the effect of pre-existing immunity against dengue virus on the Zika virus infection by using human serum, primary human cells, and infection quantification by quantitative real-time polymerase chain reaction.

The recent emergence of the flavivirus Zika and neurological complications, such as Guillain-Barr&#233; syndrome and microcephaly in infants, has brought ...

Design to Implementation Study for Development and Patient Validation of Paper-Based Toehold Switch Diagnostics

Access to low-burden molecular diagnostics that can be deployed into the community for testing is increasingly important and has meaningful wider implications for the well-being of societies and economic stability. Recent years have seen several new isothermal diagnostic modalities emerge to meet the need for rapid, low-cost molecular diagnostics. We have contributed to this effort through the development and patient validation of toehold switch-based diagnostics, including diagnostics for the mosquito-borne Zika and chikungunya viruses, which provided performance comparable to gold-standard reverse transcription-quantitative polymerase chain reaction (RT-qPCR) based assays. These diagnostics are inexpensive to develop and manufacture, and they have the potential to provide diagnostic capacity to low-resource environments. Here the protocol provides all the steps necessary for the development of a switch-based assay for Zika virus detection. The article takes readers through the stepwise diagnostic development process. First, genomic sequences of Zika virus serve as inputs for the computational design of candidate switches using open-source software. Next, the assembly of the sensors for empirical screening with synthetic RNA sequences and optimization of diagnostic sensitivity is shown. Once complete, validation is performed with patient samples in parallel with RT-qPCR, and a purpose-built optical reader, PLUM. This work provides a technical roadmap to researchers for the development of low-cost toehold switch-based sensors for applications in human health, agriculture, and environmental monitoring.

Access to decentralized, low-cost, and high-capacity diagnostics that can be deployed into the community for decentralized testing is critical for combating global health crises. This manuscript describes how to build paper-based diagnostics for viral RNA sequences that can be detected with a portable optical reader.

Access to low-burden molecular diagnostics that can be deployed into the community for testing is increasingly important and has meaningful wider ...

High-throughput Antiviral Assays to Screen for Inhibitors of Zika Virus Replication

Antiviral drug discovery requires the development of reliable biochemical and cellular assays that can be performed in high-throughput screening (HTS) formats. The flavivirus non-structural (NS) proteins are thought to co-translationally assemble on the endoplasmic reticulum (ER) membranes, forming the replication complex (RC). The NS3 and NS5 are the most studied enzymes of the RC and constitute the main targets for drug development due to their crucial roles in viral genome replication. NS3 protease domain, which requires NS2B as its cofactor, is responsible for the cleavage of the immature viral polyprotein into the mature NS proteins, whereas NS5 RdRp domain is responsible for the RNA replication. Herein, we describe in detail the protocols used in replicon-based screenings and enzymatic assays to test large compound libraries for inhibitors of the Zika virus (ZIKV) replication. Replicons are self-replicating subgenomic systems expressed in mammalian cells, in which the viral structural genes are replaced by a reporter gene. The inhibitory effects of compounds on viral RNA replication can be easily evaluated by measuring the reduction in the reporter protein activity. The replicon-based screenings were performed using a BHK-21 ZIKV replicon cell line expressing Renilla luciferase as a reporter gene. To characterize the specific targets of identified compounds, we established in-vitro fluorescence-based assays for recombinantly expressed NS3 protease and NS5 RdRp. The proteolytic activity of the viral protease was measured by using the fluorogenic peptide substrate Bz-nKRR-AMC, while the NS5 RdRp elongation activity was directly detected by the increase of the fluorescent signal of SYBR Green I during RNA elongation, using the synthetic biotinylated self-priming template 3&#8242;UTR-U30 (5&#39;-biotin-U30-ACUGGAGAUCGAUCUCCAGU-3&#39;).

In this work, we describe the protocols used in replicon-based and viral enzyme-based assays to screen for inhibitors of Zika virus replication in a high-throughput screening format.

Antiviral drug discovery requires the development of reliable biochemical and cellular assays that can be performed in high-throughput screening (HTS) ...

A Multi-detection Assay for Malaria Transmitting Mosquitoes

The Anopheles gambiae species complex includes the major malaria transmitting mosquitoes in Africa. Because these species are of such medical importance, several traits are typically characterized using molecular assays to aid in epidemiological studies. These traits include species identification, insecticide resistance, parasite infection status, and host preference. Since populations of the Anopheles gambiae complex are morphologically indistinguishable, a polymerase chain reaction (PCR) is traditionally used to identify species. Once the species is known, several downstream assays are routinely performed to elucidate further characteristics. For instance, mutations known as KDR in a para gene confer resistance against DDT and pyrethroid insecticides. Additionally, enzyme-linked immunosorbent assays (ELISAs) or Plasmodium parasite DNA detection PCR assays are used to detect parasites present in mosquito tissues. Lastly, a combination of PCR and restriction enzyme digests can be used to elucidate host preference (e.g., human vs. animal blood) by screening the mosquito bloodmeal for host-specific DNA. We have developed a multi-detection assay (MDA) that combines all of the aforementioned assays into a single multiplex reaction genotyping 33SNPs for 96 or 384 samples at a time. Because the MDA includes multiple markers for species, Plasmodium detection, and host blood identification, the likelihood of generating false positives or negatives is greatly reduced from previous assays that include only one marker per trait. This robust and simple assay can detect these key mosquito traits cost-effectively and in a fraction of the time of existing assays.

Malaria transmitting mosquitoes have a number of epidemiologically important characteristics that can only be detected using molecular techniques. Utilizing a MALDI-TOF based SNP genotyping platform, we developed an assay for simultaneously detecting multiple key traits (species, insecticide resistance, parasite infection and host choice) of malaria vectors.

The Anopheles gambiae species complex includes the major malaria transmitting mosquitoes in Africa. Because these species are of such medical importance, ...

Biology

Single-Cell Multiplexed Fluorescence Imaging to Visualize Viral Nucleic Acids and Proteins and Monitor HIV, HTLV, HBV, HCV, Zika Virus, and Influenza Infection

Capturing the dynamic replication and assembly processes of viruses has been hindered by the lack of robust in situ hybridization (ISH) technologies that enable sensitive and simultaneous labeling of viral nucleic acid and protein. Conventional DNA fluorescence in situ hybridization (FISH) methods are often not compatible with immunostaining. We have therefore developed an imaging approach, MICDDRP (multiplex immunofluorescent cell-based detection of DNA, RNA and protein), which enables simultaneous single-cell visualization of DNA, RNA, and protein. Compared to conventional DNA FISH, MICDDRP utilizes branched DNA (bDNA) ISH technology, which dramatically improves oligonucleotide probe sensitivity and detection. Small modifications of MICDDRP enable imaging of viral proteins concomitantly with nucleic acids (RNA or DNA) of different strandedness. We have applied these protocols to study the life cycles of multiple viral pathogens, including human immunodeficiency virus (HIV)-1, human T-lymphotropic virus (HTLV)-1, hepatitis B virus (HBV), hepatitis C virus (HCV), Zika virus (ZKV), and influenza A virus (IAV). We demonstrated that we can efficiently label viral nucleic acids and proteins across a diverse range of viruses. These studies can provide us with improved mechanistic understanding of multiple viral systems, and in addition, serve as a template for application of multiplexed fluorescence imaging of DNA, RNA, and protein across a broad spectrum of cellular systems.

Presented here is a protocol for a fluorescence imaging approach, multiplex immunofluorescent cell-based detection of DNA, RNA, and protein (MICDDRP), a method capable of simultaneous fluorescence single-cell visualization of viral protein and nucleic acids of different type and strandedness. This approach can be applied to a diverse range of systems.

Capturing the dynamic replication and assembly processes of viruses has been hindered by the lack of robust in situ hybridization (ISH) technologies that ...

Multiplex RT-qPCR Assay for Respiratory Virus Detection

Development of Multiplex Real-Time RT-qPCR Assays for the Detection of SARS-CoV-2, Influenza A/B, and MERS-CoV

The severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) that causes Coronavirus disease 2019 (COVID-19) is a serious threat to the general public&#39;s health. During influenza seasons, the spread of SARS-CoV-2 and other respiratory viruses may cause a population-wide burden of respiratory disease that is difficult to manage. For that, the respiratory viruses SARS-CoV-2, Influenza A, Influenza B, and Middle East respiratory syndrome (MERS-CoV) will need to be carefully watched over in the upcoming fall and winter seasons, particularly in the case of SARS-CoV-2, Influenza A, and Influenza B, which share similar epidemiological factors like susceptible populations, mode of transmission, and clinical syndromes. Without target-specific assays, it can be challenging to differentiate among cases of these viruses owing to their similarities. Accordingly, a sensitive and targeted multiplex assay that can easily differentiate between these viral targets will be useful for healthcare practitioners. In this study, we developed a real-time reverse transcriptase-PCR-based assay utilizing an in-house developed R3T one-step RT-qPCR kit for simultaneous detection of SARS-CoV-2, Influenza A, Influenza B, and SARS-CoV-2, MERS-CoV. With as few as 10 copies of their synthetic RNAs, we can successfully identify SARS-CoV-2, Influenza A, Influenza B, and MERS-CoV targets simultaneously with 100% specificity. This assay is found to be accurate, reliable, simple, sensitive, and specific. The developed method can be used as an optimized SARS-CoV-2, Influenza A, Influenza B, and SARS-CoV-2, MERS-CoV diagnostic assay in hospitals, medical centers, and diagnostic laboratories as well as for research purposes.

Author Spotlight: Advancements in Multiplex Detection of Respiratory Viruses 

We present two probe-based in-house one-step RT-qPCR kits for common respiratory viruses. The first assay is for SARS-CoV-2 (N), Influenza A (H1N1 and H3N2), and Influenza B. The second is for SARS-Cov-2 (N) and MERS (UpE and ORF1a). These assays can be successfully implemented in any specialized laboratory.

The severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) that causes Coronavirus disease 2019 (COVID-19) is a serious threat to the general ...

Multiplexed Isothermal Amplification Based Diagnostic Platform to Detect Zika, Chikungunya, and Dengue 1

Summary

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Chapters in this video

Reverse Transcription-Loop-mediated Isothermal Amplification (RT-LAMP) Assay for Zika Virus and Housekeeping Genes in Urine, Serum, and Mosquito Samples

Visual Detection of Multiple Nucleic Acids in a Capillary Array

Zika Virus Specific Diagnostic Epitope Discovery

A Simple Flow Cytometry Based Assay to Determine In Vitro Antibody Dependent Enhancement of Dengue Virus Using Zika Virus Convalescent Serum

Quantification of Antibody-dependent Enhancement of the Zika Virus in Primary Human Cells

Design to Implementation Study for Development and Patient Validation of Paper-Based Toehold Switch Diagnostics

High-throughput Antiviral Assays to Screen for Inhibitors of Zika Virus Replication

A Multi-detection Assay for Malaria Transmitting Mosquitoes

Single-Cell Multiplexed Fluorescence Imaging to Visualize Viral Nucleic Acids and Proteins and Monitor HIV, HTLV, HBV, HCV, Zika Virus, and Influenza Infection

Development of Multiplex Real-Time RT-qPCR Assays for the Detection of SARS-CoV-2, Influenza A/B, and MERS-CoV