Dr. Pinto is funded by a seed grant from the Saint Louis University School of Medicine and startup funds from Saint Louis University School of Medicine. Dr. Brien is funded by a K22AI104794 early investigator award from the NIH NIAID as well a seed grant from the Saint Louis University School. For all funded individuals the funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

All procedures of the present study are in accordance with the guidelines set by the St. Louis University Animal Care and Use Committee. SLU is fully accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International (AAALAC).
1. Organ Isolation
NOTE: The virus is not stable at room temperature (RT) so the number of animals harvested at one time must be planned carefully to preserve viral titers.
<ol>
	<li>Infect mice using the chosen dose and route, based upon the phenotype required. For this protocol, infect 8-10 week old male and female type I interferon receptor deficient (Ifnar1-/-) C57BL/6 mice.
	<ol>
		<li>Anesthetize Ifnar1-/- mice with a cocktail of Ketamine (90 mg/kg) and Xylazine (10 mg/kg). Test pedal reflex by a firm toe pinch to confirm anesthesia. Administer 1 x 105 focus forming units (FFU) of ZIKV in 50 &#956;L subcutaneously (SC) via the footpad.</li>
	</ol>
	</li>
	<li>Prepare all the materials needed for harvest the day before: 2 scissors, forceps and 20 mL of 70% ethanol (EtOH) in a 50 mL conical for disinfection of tools.
	<ol>
		<li>Prepare syringes and needles: a 20 mL syringe for perfusion, 23 G needles (approximately 1 per cage) and 1 mL syringe with 25 G needle (1 per 3-5 mouse). Prior to weighing tubes, add steel beads (in hood) to 1.5 mL O-ring screw cap tubes (one for each organ). The tubes need to be appropriate for homogenization.</li>
	</ol>
	</li>
	<li>Prepare the day of the harvest the materials needed for the perfusion and organ freezing.
	<ol>
		<li>Fill an ice bucket with dry ice and 70% EtOH to make an ice bath. Prepare sterile phosphate buffered saline (PBS) assuming that 25-30 mL per mouse of PBS for each perfusion is needed 5-10 mL for the spinal cord). Obtain anesthesia, a cocktail of Ketamine and Xylazine, and assume 300 &#956;L per mouse. Place all the materials and any additional tubes required for flow cytometry, etc., in a secondary container to transport to the animal facility.</li>
	</ol>
	</li>
	<li>Follow all necessary procedures for entry including donning and doffing personal protection equipment during entry and exit into the animal facility. 
	CAUTION: All the procedures are to be carried out in a certified biosafety cabinet within a Biosafety level 2 laboratory (BSL-2) facility.</li>
	<li>Administer anesthesia, 200-500 &#956;L, intraperitoneally depending on size and weight of the animal. If working alone, administer anesthesia to one animal at a time to avoid killing the animals before organ harvest.
	<ol>
		<li>Confirm anesthesia dose by pinching toe with forceps to evaluate the pedal reflex. Do not continue unless completely nonresponsive. Use pins to secure the mouse to a wax board. At this point, douse the mouse in 70% ethanol to avoid hair contamination in the organ harvest procedure</li>
	</ol>
	</li>
	<li>Terminally bleed by heart puncture.
	<ol>
		<li>Using the scissors and forceps, open the animal through the chest cavity to expose the heart. Collect blood via heart puncture (~800 &#181;L), this can be used for multiple assays including clinical chemistry, hematology, flow cytometry to detect antigen specific responses, and real time quantitative PCR (qRT-PCR) for the analysis of viral copy number.</li>
		<li>For viral RNA analysis by qRT-PCR, collect blood in an EDTA tube if extracting from whole blood or in a microcentrifuge tube if extracting from serum. (For serum, spin tube at maximum speed in centrifuge, room temp, for 20 min and remove serum to a separate tube). Add a linear polyacrylamide as a carrier to the serum sample after finishing. RNA can then be analyzed or stored at -80 &#176;C to further process at a later date.</li>
	</ol>
	</li>
	<li>Fill the 20 mL syringe with PBS, at room temperature (or 37&#160;&#176;C), and perfuse mice by inserting the butterfly needle into the left ventricle. Puncture the right atrium to allow blood and PBS to exit. Slowly administer the PBS while checking the color of the liver to confirm that the animal is completely perfused. Liver should change from deep red to pink salmon color.
	<ol>
		<li>If preparing the organs for histology, leave butterfly needle in place and then perfuse with 20 mL of ice cold 4% paraformaldehyde. If so, it is convenient to have the syringe connected to a 3-way stopcock with both syringes attached to it, turning the valve ON and OFF as one alternates between PBS and PFA.</li>
	</ol>
	</li>
	<li>Harvest organs into labeled, weighed tubes. For peripheral organs, follow an established order for harvest: liver, spleen, kidney and lungs.
	<ol>
		<li>Take only one lobe from the liver; it does not matter which one, but for all experiments always try to take the same lobe with the same size piece. Similarly, for kidneys and lungs, take the same kidney and lung. If flow cytometry is also to be completed, any organ can be cut in half. Store the half to be used for cytometry in Roswell Park Memorial Institute medium (RPMI) at room temperature until the harvest is complete.</li>
		<li>Immediately after harvest, put each organ in a labeled tube and place in the dry ice bath. Virus titer reduces over time at room temperature, so the amount of time it takes to harvest organs is extremely important. There must be consistency between mice, so after safety, the next priority is speed. 
		NOTE: If harvesting organs for viral titers, it is very helpful to have a second individual harvest the brain at this point, to preserve viral titers.</li>
	</ol>
	</li>
	<li>Remove the remaining organs from the mouse body cavity to gain access to the spine and skull.
	<ol>
		<li>Remove the mouse from the board and remove the pelt, followed by the removal of the arms and legs. 
		&#8203;Remove the head of the mouse with a decapitation, blunt or surgical scissors to harvest the brain by cutting the skull with a serrated LaGrange scissors through the foramen magnum. Then, peel off the skull with forceps and scoop out the brain with a spatula.</li>
		<li>Using strong, blunted scissors, remove the ribs and other bones surrounding the spine. Then, cut across the pelvic bone, exposing the vertebral foramen at the lumbar level. The small tip of the spinal cord should be visible at this point.</li>
		<li>Use a 10 mL syringe filled with PBS and a 18 G needle to expel the spinal cord by &#8220;flushing&#8221; the cord from lumbar to cervical spine over a Petri dish.</li>
		<li>Carefully, place the beveled tip of the needle inside the vertebral foramen, avoiding excessive pressure to prevent the needle to trespassing the vertebral body. Hold strongly to exert pressure on the vertebral body and the needle and press the syringe plunger to expel the cord. Immediately transfer the spinal cord in the labeled tube and place in the dry ice bath.</li>
	</ol>
	</li>
	<li>Repeat the procedure with all the animals until the harvest is completed. Place the harvest tools in the 70% ethanol between animals. Focus on consistency and speed. The length of time between each organ harvest should remain consistent so as to not bias viral titer results.</li>
	<li>When finished, disinfect the biosafety cabinet and all material prior to removing it from the animal facility.</li>
	<li>Remove each tube from the ice bath and weigh tubes to determine organs weight. To determine the organs weight, subtract the weight of the empty tube from the weight of the organ containing tube. 
	NOTE: At this point organs can be frozen at -80 &#176;C to further process at a later date or organs can be homogenized immediately before freezing individual aliquots. Freeze thawing samples multiple times decreases viral titer. Therefore, it is important to do same procedure for all experiments within a single project.</li>
</ol>
2. Organ Homogenization
<ol>
	<li>Prepare 3 labeled, 1.5 mL snap-capped tubes for each organ to be homogenized.</li>
	<li>If samples are not being homogenized immediately after harvest, remove tubes from -80 &#176;C. The samples do not need to be thawed to homogenize.</li>
	<li>Put the samples on ice to keep them cold to minimize viral titer loss. Then, add 1 mL of cold DMEM containing 5% FBS to each organ containing tube.</li>
	<li>Immediately beat tubes in bead beater according to manufacturer&#8217;s instructions. Homogenize all organs with steel beads in a bead-homogenizer instrument. Check to ensure each organ has been completely homogenized.</li>
	<li>Spin down organ debris in microfuge at 12,000 x g for 5 min in a microfuge that has been chilled to 8 &#176;C. Then return the tubes to the ice bucket. In the biosafety cabinet, aliquot samples into tubes for necessary assays. Then return the tubes to the ice bucket.</li>
	<li>For focus forming assay, aliquot 500 &#181;L into a labeled tube. Then place tubes in the rack on an ice bucket. Aliquot 50 &#181;L into a tube for RNA for fluorogenic quantitative RT-PCR (qRT-PCR) to measure viral genome copy number.</li>
	<li>Isolate total RNA from the organs of the infected animals using a commercial RNA isolation kit. Determine flavivirus viral RNA using the primer probe sets specific for ZIKV, which recognizes unique sequences in each flavivirus genome. Determine viral copy number using a copy control plasmid containing a defined positive single-stranded RNA generated in vitro using T7 polymerase containing the ZIKV target sequences.</li>
	<li>Aliquot the remaining sample, which is approximately 300 &#181;L, into the third tube and store at -80 &#176;C if needed. 
	NOTE: If samples were not previously frozen, freeze at -80 &#176;C. If samples had been previously frozen, continue onto the focus forming assay and/or RNA isolation before stopping.</li>
</ol>
3. Zika Virus Focus Forming Assay26
NOTE: It is important to include a no virus control and a positive control. The positive control is a dilution series of a virus stock with a known concentration. Not all controls need to be on the same plate, but as the assay becomes larger than 5 plates, more controls should be added, and spread out among plates. Take care not to scratch the monolayer with either the pipet tips or by vigorous washing. Multiple organs can be titered on the same day or on different days. But an individual organ should not be titered over multiple days because different assay conditions can impact viral titer. It is strongly recommended to run an individual organ on a single day.
<ol>
	<li>Prior to the day of the assay, prepare the cells and reagents needed for the focus forming assay.
	<ol>
		<li>Prepare growth media containing 500 mL of DMEM with 5 mL of HEPES and 25 mL of FBS. Have Vero-World Health Organization (WHO) cells growing in growth media at 37 &#176;C, 5% CO2 prior to the start of the assay. The Vero cells should not be a high passage or ever grown over 100% confluency prior to the start of the assay.</li>
		<li>Prepare 500 mL of a 2% methylcellulose solution by autoclaving a 1 L glass media bottle with 10 g of methylcellulose and a large stir bar and a separate 1 L glass media bottle with 500 mL of H2O. If the autoclaved water has cooled reheat in microwave until bottle is hot to the touch, but not boiling.</li>
		<li>Gently pour warm/hot water into bottle of methylcellulose while in the tissue culture hood. Partially cap bottle and stir on the hotplate until methylcellulose is in solution (1-4 h). Aliquot the 2% methylcellulose solution into sterile 50 mL conical tube. 2% Methylcellulose can be stored at 4 &#176;C until needed.</li>
		<li>Prepare a 5% Paraformaldehyde solution in PBS for fixing the plates and stored at 4 &#176;C until needed. Prepare a 1x focus forming assay wash buffer by adding 0.05% Triton X-100 to PBS and stored at RT. Prepare a 1x FFA staining buffer by adding 1 mg/mL saponin to PBS and stored at 4 &#176;C until needed. These can be prepared one- two weeks in advance.</li>
	</ol>
	</li>
	<li>Calculate the number of flat-bottom 96 well plates needed for the assay such that each organ is plated in triplicate. Include enough additional wells for positive and negative controls.
	<ol>
		<li>Grow up enough Vero cells in growth media to complete the assay designed. Trypsinize the Vero cells and count them resuspending them at 1.5 x 105 cells per mL in growth media. Plate Vero cells in the 96 well flat-bottom plates at 3.0 x 104 Vero-WHO cells/well in growth media by adding 200 &#181;L per well.</li>
		<li>Incubate plates at 37&#176;C at 5% CO2 overnight, make sure the plates are level in the incubator so the cells are equally distributed within the well. 
		NOTE: Each laboratory grows Vero cells slightly differently; the target is a 90-95% confluent monolayer in each well on the day of the assay for Zika virus.</li>
	</ol>
	</li>
	<li>Dilute Zika virus samples the day of the assay. If the organ samples had been previously homogenized, remove samples from the -80 &#176;C freezer and allow them to thaw before placing the samples on ice for the assay.
	<ol>
		<li>On ice, prepare a round bottom 96 well plate by adding 180 &#181;L of cold growth media to rows B through H leaving row A empty. Add 150 &#181;L of each homogenized organ sample to row A of the round bottom plate.</li>
		<li>Prepare serial 10-fold dilutions of each sample, using a multi-channel pipette. Dilute samples in a round bottom 96 well plate by adding 20 &#181;L of sample into 180 &#181;L of growth media, changing pipette tips between each dilution.</li>
	</ol>
	</li>
	<li>Prepare the focus forming plate by removing the media from the flat-bottom 96 well plate covering Vero cells. Do this immediately before adding the virus samples to prevent the monolayer from drying out.
	<ol>
		<li>Add 100 &#181;L of the virus dilution to each well in the Vero plate. Add sample using the same set of tips by going from the lowest to the highest concentration. Rock plates side-to-side 2-4 times being careful not to swirl.</li>
		<li>Incubate at 37 &#176;C, 5% CO2 for 1-2 h. Make sure the plates are level within the incubator. During the incubation warm up 2% methylcellulose to RT.</li>
		<li>Dilute 2% methylcellulose solution in growth media. The dilution should be at a ratio of approximately 2:1 of 2% methylcellulose to growth media. Keep at room temp until it is time to use it. Add 1-2 drops/well (set the pipet to 125 &#181;L) of the methylcellulose: growth media to each well of the 96 well plate.</li>
		<li>Incubate 32-40 h at 37 &#176;C, 5% CO2. As everyone&#8217;s Vero cells grow slightly differently and factors including cell confluency and strain of Zika virus can change the incubation time.</li>
	</ol>
	</li>
	<li>Fix the Vero cells using the prepared 5% paraformaldehyde solution.
	<ol>
		<li>Add 50 &#181;L of 5% paraformaldehyde to each well over the top of the methylcellulose layer in the biosafety cabinet. Incubate for 60 min at RT. The fixing can go overnight at 4 &#176;C but cover the plate with parafilm to reduce evaporation.</li>
		<li>Dump overlay and media off cells into a disposal container inside the biosafety cabinet. Wash gently with PBS, 150 &#181;L/well. Remove PBS from the plates and remove the plate from the BSL-2.</li>
		<li>Repeat the PBS wash 2x adding 150 &#181;L/well. Then remove the PBS. Add 150 &#181;L/well 1x FFA wash buffer and let sit for 5-10 min at RT to permeabilize the fixed cells.</li>
	</ol>
	</li>
	<li>Detect infection using primary Zika antibody. Prepare the primary antibody 4G2 (D1-4G2-4-15) at a concentration of 1 &#956;g/mL in FFA staining buffer. Prepare enough antibody for the whole assay. 
	NOTE: Stay away from lab diapers or other high-lint absorbent material as the fibers will negatively impact the imaging of the foci.
	<ol>
		<li>Remove FFA wash buffer from the plates. Add 50 &#181;L/well of the primary antibody in the FFA staining buffer. Seal the plates with parafilm and incubate overnight at 4 &#176;C on a rocking platform. The assay can be done with an incubation for 2 h at RT.</li>
	</ol>
	</li>
	<li>Visualize the infection by the addition of a secondary HRP conjugated antibody. Prepare the secondary Goat anti-mouse HRP-labeled antibody at a concentration of 1:5,000 in FFA staining buffer. Prepare enough antibody for the whole assay.
	<ol>
		<li>Wash cells 3x with FFA wash buffer, removing the wash buffer by flicking into the sink each time. Stain the cells with the secondary antibody in FFA staining buffer at 50 &#181;L/well. Incubate 1-2 h at RT.</li>
		<li>Wash cells 3x with FFA wash buffer, removing the wash buffer by flicking into the sink each time. Add 50 &#181;L/well of the Trueblue Substrate.</li>
		<li>Watch the plates carefully, waiting 2-15 min until spots are fully defined and minimal background. After the spots are visible, wash gently with water, using a hand to shield the monolayer from the force of the water running. Tap dry on paper towels (NOT DIAPERS) and image as soon as possible.</li>
		<li>Spots may be counted manually or using an automated spot counter. If counted manually, a dissecting scope can be used to aid in visualization.</li>
		<li>For each sample, select a dilution with easily distinguished foci (e.g., 20 to 200 per well) and calculate titer in focus-forming units per mL (FFU/ml), using the average of duplicate wells: FFU/mL = (mean foci/well) &#215; (dilution factor) &#247; (mL inoculum).</li>
	</ol>
	</li>
</ol>

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K., 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=Deficient+IFN+signaling+by+myeloid+cells+leads+to+MAVS-dependent+virus-induced+sepsis.">Deficient IFN signaling by myeloid cells leads to MAVS-dependent virus-induced sepsis.</a> PLoS Pathog. 10 (4), e1004086 (2014).</li><li>Brien, J. D., Lazear, H. M., Diamond, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Propagation,+quantification,+detection,+and+storage+of+West+Nile+virus.">Propagation, quantification, detection, and storage of West Nile virus.</a> Curr Protoc Microbiol. 31, (2013).</li><li>Hassert, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CD4+T+cells+mediate+protection+against+Zika+associated+severe+disease+in+a+mouse+model+of+infection.">CD4+T cells mediate protection against Zika associated severe disease in a mouse model of infection.</a> PLoS Pathogens. 14 (9), e1007237 (2018).</li><li>Fuchs, A., Pinto, A. K., Schwaeble, W. J., Diamond, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+lectin+pathway+of+complement+activation+contributes+to+protection+from+West+Nile+virus+infection.">The lectin pathway of complement activation contributes to protection from West Nile virus infection.</a> Virology. 412 (1), 101-109 (2011).</li><li>Lazear, H. M., Pinto, A. K., Vogt, M. R., Gale, M., Diamond, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Beta+interferon+controls+West+Nile+virus+infection+and+pathogenesis+in+mice.">Beta interferon controls West Nile virus infection and pathogenesis in mice.</a> Journal of Virology. 85 (14), 7186-7194 (2011).</li></ol>

The authors have nothing to disclose

ZIKV infection can cause a neurological disease therefore the current animal models to study pathogenesis, immune responses and protective efficacy of vaccines and antivirals need to focus on viral control within the CNS. One of the challenges in focusing on CNS disease is that it often comes at the expense of studying peripheral infection. The organ isolation methods proposed here focuses on the need to rapidly evaluate ZIKV infection in both the periphery and the CNS in order to assess CNS mediated ZIKV associated disease and establish a model for preclinical testing of antivirals, therapeutic and vaccines. An added benefit of this technique is that it also allows for a high degree of flexibility, including the combined study of immunological responses to ZIKV or histological analysis of infection. This technique, is not restricted to just ZIKV but can also be universally applied to study a range of host-pathogen interactions, including flaviviruses such as Dengue virus21, orthopoxviruses like monkeypox and ectromelia. The considerations and drawbacks to this technique of harvesting focus mainly of the capabilities of the experimenter. As ZIKV is not stable for long periods of time at room temperature, the amount of time it takes to harvest organs after perfusion can significantly impact the quality of the results. For most experiments that we have performed, we compare viral titers from mice treated with two conditions, so we focus our efforts on consistency of time between organ harvests not on speed. In this way the same person performs the same procedure for the whole experiment to maintain consistency. The other major consideration with this procedure is safety, we have readily performed these methods with BSL-2 (ZIKV, Dengue virus) and BSL-3 (WNV, Chikungunya virus) pathogens. It is very important to perform all procedures in a clean, well maintained, certified biosafety cabinet with disinfectant.
An FFA parallels the plaque assay, except that it uses peroxidase immunostaining to identify foci of infected cells, rather than plaques. My laboratory as well as multiple other laboratories have now successfully switched to using FFAs for all our tittering experiments11,14,15,17,26,27,28. The FFA has multiple advantages over the traditional plaque assay: a) The FFA is faster, requiring a shorter incubation compared to a plaque assay, b) it is also higher-throughput, being performed in 96-well plates. The 96 well plate format can also accommodate smaller volumes of starting material. In addition, c) the FFA is compatible with the use of an automated plate washer and automated spot counter, greatly reducing the labor and time required for the assay. The FFA has more steps after infection, but d) with the use of multichannel pipets, or even a pipetting robot, the timing for most of the assay steps after fixation are flexible and the assay can be paused overnight or for longer. Finally, e) it may be especially useful for virus strains that do not form clear plaques, such as Dengue virus. One disadvantage of the FFA is that it requires specific antibodies to detect virus-infected cells, which may be confounding when considering diverse virus strains or mutant viruses. For the FFA as with the plaque assay the density of the cell monolayer at the time of infection is critical for the success of the assay. Cells should be used at higher confluence for a FFA compared to a plaque assay, due to the shortened length of time before fixation. As the FFA is higher throughput, more cost effective and faster than the traditional plaque assay it allows my laboratory to rapidly analyze data for studying emerging infectious diseases. The FFA is more cumulatively more cost effective for several reasons. Although antibodies are more expensive than neutral red or crystal violet, we are able to analyze more samples per plate which eliminates the cost difference. In terms of labor allocation automated spot counting and easy data entry for analysis limits labor costs and the ability to have a long-term image as a record is difficult to quantify. The future of this assay may be to move to a fluorescent-based foci for a readout as opposed to HRP. Quantitating fluorescent intensity along with spot number will extend the utility of the FFA beyond what is currently studied.

Zika virus (ZIKV) is an arbovirus that belongs to the flaviviridae family, which includes important neuroinvasive human pathogens such as Powassan virus (POWV), Japanese encephalitis virus (JEV), and West Nile virus (WNV)1. Following its isolation and identification, there have been periodic reports of human ZIKV infections in Africa and Asia2,3,4,5, and epidemics within Central and South America (reviewed in reference6). However, it was not until recently that ZIKV was thought to cause severe disease7. Now there are thousands of cases of neurological disease and birth defects linked to ZIKV infections. The rapid emergence of ZIKV has prompted many questions relating to: why there is an increase in disease severity, what is the immunological response to ZIKV infection and are there viral and/or immune mediated pathologies linked to the increase in neurological manifestations and birth defects. There is now a rush to understand the central nervous system (CNS) related disease associated with ZIKV as well as the need to rapidly test the efficacy of the antivirals and vaccines against ZIKV. It is against this backdrop that we have developed methods for the rapid analysis of ZIKV titers in both the periphery and CNS using a ZIKV-specific focus forming assays (FFA).
Small animal models are important for understanding disease progression and for the early evaluation of vaccines, therapeutics, and anti-virals. We have established small animal models for the study of arbovirus disease by using various mouse strains to model human infection and protection against viral pathogens8,9,10,11,12,13,14,15,16,17,18,19,20,21,22. Using this prior experience, we began to modify techniques used for the assessment of WNV and Dengue virus, a related flavivirus for the assessment of ZIKV titer in both peripheral organs as well as the CNS21,23,24. The advantages of these methods over other assays are: 1) that they combine the ability to harvest both peripheral and CNS organs for the analysis; 2) the methods are adaptable for flow cytometry, for measurements of innate and adaptive immune responses, along with viral titers on the same animal in the same organ; 3) the harvest technique is adaptable for histological analysis; 4) the ZIKV FFA is a rapid high throughput method for viral titer analysis; and 5) these methods can be applied to the assessment of viral titers in the organs of animals infected with most pathogens25.

<table>
	<tbody>
		<tr>
			<td>1-bromo-3-chloropropane (BCP)</td>
			<td>MRC gene</td>
			<td>BP151</td>
			<td></td>
		</tr>
		<tr>
			<td>10cc syringe</td>
			<td>Thermo Fisher Scientific</td>
			<td>BD 309642</td>
			<td></td>
		</tr>
		<tr>
			<td>18G needle</td>
			<td>Thermo Fisher Scientific</td>
			<td>22-557-145</td>
			<td></td>
		</tr>
		<tr>
			<td>1cc TB syringe</td>
			<td>Thermo Fisher Scientific</td>
			<td>14-823-16H</td>
			<td></td>
		</tr>
		<tr>
			<td>20cc syringe</td>
			<td>Thermo Fisher Scientific</td>
			<td>05-561-66</td>
			<td></td>
		</tr>
		<tr>
			<td>24 tube beadmill</td>
			<td>Thermo Fisher Scientific</td>
			<td>15 340 163</td>
			<td></td>
		</tr>
		<tr>
			<td>3.2 mm stainless steel beads</td>
			<td>Thermo Fisher Scientific</td>
			<td>NC9084634</td>
			<td></td>
		</tr>
		<tr>
			<td>37C Tissue Culture incubator</td>
			<td>Nuair</td>
			<td>5800</td>
			<td></td>
		</tr>
		<tr>
			<td>4G2 antibody</td>
			<td>in house</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>96 well flat bottom plates</td>
			<td>Midsci</td>
			<td>TP92696</td>
			<td></td>
		</tr>
		<tr>
			<td>96well round bottom plates</td>
			<td>Midsci</td>
			<td>TP92697</td>
			<td></td>
		</tr>
		<tr>
			<td>Basix 1.5ml eppendorf tubes</td>
			<td>Thermo Fisher Scientific</td>
			<td>02-682-002</td>
			<td></td>
		</tr>
		<tr>
			<td>Concentrated Germicidal Bleach</td>
			<td>Staples</td>
			<td>30966CT</td>
			<td></td>
		</tr>
		<tr>
			<td>CTL S6 Analyzer</td>
			<td>CTL</td>
			<td>CTL S6 Universal Analyzer</td>
			<td></td>
		</tr>
		<tr>
			<td>curved cutting scissors</td>
			<td>Fine Science Tools</td>
			<td>14061-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#8217;s Modified Eagle&#8217;s Medium - high glucose With 4500 mg/L glucose</td>
			<td>MilliporeSigma</td>
			<td>D5671</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol (molecular biology-grade)</td>
			<td>MilliporeSigma</td>
			<td>e7023</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>MilliporeSigma</td>
			<td>F0926-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Fine Science Tools</td>
			<td>11036-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Glacial acetic acid</td>
			<td>MilliporeSigma</td>
			<td>537020</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti-mouse HRP-labeled antibody</td>
			<td>MilliporeSigma</td>
			<td>8924</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES 1 M</td>
			<td>MilliporeSigma</td>
			<td>H3537-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropanol (molecular biology-grade)</td>
			<td>MilliporeSigma</td>
			<td>I9516</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine/Xylazine cocktail</td>
			<td>Comparative Medicine</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>L-glutamine</td>
			<td>MilliporeSigma</td>
			<td>g7513</td>
			<td></td>
		</tr>
		<tr>
			<td>Magmax RNA purification kit</td>
			<td>Thermo Fisher Scientific</td>
			<td>AM1830</td>
			<td></td>
		</tr>
		<tr>
			<td>Methylcellulose</td>
			<td>MilliporeSigma</td>
			<td>M0512</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge</td>
			<td>Ependorf</td>
			<td>5424R</td>
			<td></td>
		</tr>
		<tr>
			<td>MiniCollect 0.5ml EDTA tubes</td>
			<td>Bio-one</td>
			<td>450480</td>
			<td></td>
		</tr>
		<tr>
			<td>o-ring tubes</td>
			<td>Thermo Fisher Scientific</td>
			<td>21-403-195</td>
			<td></td>
		</tr>
		<tr>
			<td>one step q RT-PCR mix</td>
			<td>Thermo Fisher Scientific</td>
			<td>4392938</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Thermo Fisher Scientific</td>
			<td>EMS- 15713-S</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline</td>
			<td>MilliporeSigma</td>
			<td>d8537-500ml</td>
			<td></td>
		</tr>
		<tr>
			<td>Proline multichannel pipettes</td>
			<td>Sartorius</td>
			<td>72230/72240</td>
			<td></td>
		</tr>
		<tr>
			<td>Proline single channel pipettes</td>
			<td>Sartorius</td>
			<td>728230</td>
			<td></td>
		</tr>
		<tr>
			<td>RNAse free water</td>
			<td>Thermo Fisher Scientific</td>
			<td>10-977-023</td>
			<td></td>
		</tr>
		<tr>
			<td>RNAzol BD</td>
			<td>MRC gene</td>
			<td>RB192</td>
			<td></td>
		</tr>
		<tr>
			<td>Rocking Platform</td>
			<td>Thermo Fisher Scientific</td>
			<td>11-676-333</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI 1640</td>
			<td>Fisher</td>
			<td>MT10040CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Saponin</td>
			<td>MilliporeSigma</td>
			<td>s7900</td>
			<td></td>
		</tr>
		<tr>
			<td>spoon/spatula</td>
			<td>Fine Science Tools</td>
			<td>10090-17</td>
			<td></td>
		</tr>
		<tr>
			<td>straight cutting scissors</td>
			<td>Fine Science Tools</td>
			<td>14060-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>MilliporeSigma</td>
			<td>t8787</td>
			<td></td>
		</tr>
		<tr>
			<td>True Blue Substrate</td>
			<td>VWR</td>
			<td>95059-168</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>MilliporeSigma</td>
			<td>T3924-100ML</td>
			<td></td>
		</tr>
	</tbody>
</table>

isolation and quantification of zika virus from multiple organs in a mouse

The methods being presented demonstrate laboratory procedures for the isolation of organs from Zika virus infected animals and the quantification of viral load. The purpose of the procedure is to quantify viral titers in peripheral and CNS areas of the mouse at different time points post infection or under different experimental conditions to identify virologic and immunological factors that regulate Zika virus infection. The organ isolation procedures demonstrated allow for both focus forming assay quantification and quantitative PCR assessment of viral titers. The rapid organ isolation techniques are designed for the preservation of virus titer. Viral titer quantification by focus forming assay allows for the rapid throughput assessment of Zika virus. The benefit of the focus forming assay is the assessment of infectious virus, the limitation of this assay is the potential for organ toxicity reducing the limit of detection. Viral titer assessment is combined with quantitative PCR, and using a recombinant RNA copy control viral genome copy number within the organ is assessed with low limit of detection. Overall these techniques provide an accurate rapid high throughput method for the analysis of Zika viral titers in the periphery and CNS of Zika virus infected animals and can be applied to the assessment of viral titers in the organs of animals infected with most pathogens, including Dengue virus.

To evaluate ZIKV titers using the protocol described above Ifnar1-/- mice were infected with ZIKV (PRVABC59) via subcutaneous (SC) injection to the footpad. Here, the administration of 1 x 105 FFU of ZIKV to 8-12 week old Ifnar1-/- mice SC is not lethal but the virus can replicate in both the periphery and CNS. This dose and route are used to study host pathogen immune responses and pathogenicity. Administration of 1 x 105 FFU of ZIKV to a 8-12 week old Ifnar1 -/- mouse intravenous (IV) injection is between 80 to 100% lethal, with the animal succumbing to infection between 8 to 14 days post virus injection. We routinely use this administration route to determine efficacy of antivirals and therapeutics, as well as preclinical vaccine candidate testing.
Four 10-12 week-old Ifnar1-/- mice were infected with 1 x 105 FFU of ZIKV SC and spleens, livers, kidneys, spinal cords and brains were harvested four days post infection by the methods detailed above (Figure 1). The amount of ZIKV in the tissues was assayed by focus forming assay (FFA) using Vero cells in a 96 well format as described above. Using the FFA, tissue viral load is expressed as focus forming units (FFU) per g of tissue. Similar to what was observed in a previous study of ZIKV infection of Ifnar1-/- 26, we saw viremia following a sampling of viral titers in different organs four days post ZIKV infection. These results indicate that the methods used for organ harvest and tittering by focus forming assay can be used to detect titer in both peripheral organs and the CNS within the same animal. Interestingly, we did not expect to see high viral titers in both the periphery and the CNS four days post infection in the Ifnar1-/- mice because all of the Ifnar1-/- survive ZIKV infection with this dose and route. We are continuing to explore this observation to understand how ZIKV can continue to replicate in the CNS of Ifnar1-/- without causing lethality.
When performing the focus forming assay (FFA), there are multiple technical mistakes an investigator can make which will result in suboptimal FFA results. The most common mistakes are: 1) organ toxicity; 2) vigorous pipetting; 3) fiber contamination; and 4) incorrect cell plating density. We discuss each of these issues below and illustrate the outcome in Figure 2. One of the more common issues that occurs with both the FFA and plaque assay is organ toxicity (Figure 2A red arrow). We believe organ toxicity is driven by the high concentration of intracellular components released during organ homogenization. Organ toxicity varies based upon the organ and is seen in organs harvested from uninfected animals, with the liver being the most toxic and the spleen the least. Toxicity is reduced as the organ is serially diluted on the FFA plate. However, toxicity alters the sensitivity of the assay resulting in a change in the limit of detection. As shown in Figure 2A if the viral titer in the organ is lower than the toxicity the FFA will not be able to accurately record the viral titers. Figure 2B illustrates toxicity in wells a1-4, but the viral titer is sufficiently high to overcome organ toxicity as seen in wells b3 and b4. To overcome this limitation in the FFA, we also perform quantitative real-time PCR on organ titer samples. In Figure 2C, we illustrate several common technical errors. Vigorous pipetting or washing can remove the monolayer (Figure 2C, *), if this occurs in wells with foci that data will be lost leading to inaccurate reporting of titer results. Fibers or hairs, that are present in lab bench absorbent paper can contaminate individual wells (Figure 2C, $) this can cause significant errors if using an automated counting program. While most automated counting programs have fiber exclusion options, we have not found it to be highly effective at excluding fibers from the analysis. The solution to this is to manually count the wells, which can be very time consuming and is not practical for the analysis of large assays. Cell density is another issue which can dramatically impact the success of a focus forming assay (Figure 2D). If cells are not at the right density at the start of the assay the number and size of the spots will be impacted. As shown in Figure 2D, columns 1-3, cells at approximately 60% confluency at the start of the assay compared to cells plated at 90% confluency columns 4-6 will dramatically impact the focus forming assay. To overcome this obstacle small pilot assays should be run to optimize cell density and fixation times as individual laboratory conditions will impact the success for the assay.
For studies when different groups of infected animals are compared, the statistical analysis that is performed is dependent on the distribution of data. Either, parametric or nonparametric tests are used to assess statistical significance. For parametric tests, ANOVA is utilized to detect overall effect, and individual treatment groups will be compared using Dunn&#8217;s test. In case the distribution of data does not satisfy requirements for parametric analysis, nonparametric tests are employed. The Kruskal-Wallis test is used to detect the overall treatment effect, and the Mann-Whitney U test is used to perform pair-wise comparisons. For the results present here we did not compare the animals with a second data set harvested at this time point so we did not perform statistical analysis on the data set shown.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59632/59632fig1.jpg" /> 
Figure 1: Viral replication in the periphery and CNS. Viral burden in the peripheral and CNS tissues after IFNAR1-/- mice are given 1x105 FFU of ZIKV SC. On day 4 (n = 4 per group) post-infection organs were harvested, snap frozen, weighed, and homogenized. Levels of virus were quantified by focus forming assay. The limit of detection is 100-500 FFU/gram based upon organ. Data is shown as Log10 focus-forming unit per gram of tissue. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59632/59632fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59632/59632fig2.jpg" /> 
Figure 2: Common Difficulties with the Focus-forming assay. For all the focus forming assays shown viral antigen was detected with an anti-flavivirus MAb, followed by immunoperoxidase staining (purple). (A) Vero cells were grown to a 90% confluency and infected with a 10-fold serial dilution of supernatant from liver harvested from a 8 week old C57BL/6 mouse, IV infected with 5 x 107 FFU of ZIKV 4 days previously. The red arrow indicates the highest or &#8220;neat&#8221; concentration of liver supernatant demonstrating the toxicity at this concentration. (B) Vero cells were grown to a 90% confluency and infected with a 10 fold dilution of supernatant from ZIKV infected kidney cells. In this case samples in column 1 and 2 are from a C57BL/6 mouse and col 3 and 4 are from a Ifnar1-/-. Both mice were 8 weeks old infected with 1 x 105 FFU of ZIKV IV and sacrificed 4 days post infection. Similar to (A) there is some toxicity seen at the highest concentration (row a) but the viral titers observed in column 3 and 4 overcome the limit of detection issues allowing for accurate titers to be detected. (C) Vero cells were plated. The selected wells show to common technical errors. The * demonstrates an area where the monolayer was removed due to vigorous pipetting. The $ is placed over a well where a fiber can be seen. (D) Vero cell concentration affects the sensitivity of the assay. In this plate Vero cells were plated at 1.0 x 104 cells/well in column 1-3 and 3.0 x 104 Vero-WHO cells/well in column 4-6. Then the plate was infected with 10-fold dilutions of ZIKV PRVABC59 stock. The highest viral concentration samples are in row A and diluted down, with each row representing a 10-fold dilution. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59632/59632fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Isolation and Quantification of Zika Virus from Multiple Organs in a Mouse at JoVE.com

Isolation and Quantification of Zika Virus from Multiple Organs in a Mouse

The goal of the protocol is to demonstrate the techniques used to investigate viral disease by isolating and quantifying Zika virus, from multiple organs in a mouse following infection.

The methods being presented demonstrate laboratory procedures for the isolation of organs from Zika virus infected animals and the quantification of viral ...

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Immunology and Infection

7.4K Views.  Saint Louis University. The goal of the protocol is to demonstrate the techniques used to investigate viral disease by isolating and quantifying Zika virus, from multiple organs in a mouse following infection.

Video: Isolation and Quantification of Zika Virus from Multiple Organs in a Mouse

Isolation of Lymphocytes from Mouse Genital Tract Mucosa

Mucosal surfaces, including in the gastrointestinal, urogenital, and respiratory tracts, provide portals of entry for pathogens, such as viruses and bacteria 1. Mucosae are also inductive sites in the host to generate immunity against pathogens, such as the Peyers patches in the intestinal tract and the nasal-associated lymphoreticular tissue in the respiratory tract. This unique feature brings mucosal immunity as a crucial player of the host defense system. Many studies have been focused on gastrointestinal and respiratory mucosal sites. However, there has been little investigation of reproductive mucosal sites. The genital tract mucosa is the primary infection site for sexually transmitted diseases (STD), including bacterial and viral infections. STDs are one of the most critical health challenges facing the world today. Centers for Disease Control and Prevention estimates that there are 19 million new infectious every year in the United States. STDs cost the U.S. health care system $17 billion every year 2, and cost individuals even more in immediate and life-long health consequences. In order to confront this challenge, a greater understanding of reproductive mucosal immunity is needed and isolating lymphocytes is an essential component of these studies. Here, we present a method to reproducibly isolate lymphocytes from murine female genital tracts for immunological studies that can be modified for adaption to other species. The method described below is based on one mouse.&#x2003;

An efficient way to isolate lymphocytes from mouse genital tract is described. This method takes advantage of enzyme digestion and Percoll gradient separation to allow efficient isolation. This technique is also adaptable to for use in other species

Mucosal surfaces, including in the gastrointestinal, urogenital, and respiratory tracts, provide portals of entry for pathogens, such as viruses and ...

Nasal Wipes for Influenza A Virus Detection and Isolation from Swine

Surveillance for influenza A viruses in swine is critical to human and animal health because influenza A virus rapidly evolves in swine populations and new strains are continually emerging. Swine are able to be infected by diverse lineages of influenza A virus making them important hosts for the emergence and maintenance of novel influenza A virus strains. Sampling pigs in diverse settings such as commercial swine farms, agricultural fairs, and live animal markets is important to provide a comprehensive view of currently circulating IAV strains. The current gold-standard ante-mortem sampling technique (i.e. collection of nasal swabs) is labor intensive because it requires physical restraint of the pigs. Nasal wipes involve rubbing a piece of fabric across the snout of the pig with minimal to no restraint of the animal. The nasal wipe procedure is simple to perform and does not require personnel with professional veterinary or animal handling training. While slightly less sensitive than nasal swabs, virus detection and isolation rates are adequate to make nasal wipes a viable alternative for sampling individual pigs when low stress sampling methods are required. The proceeding protocol outlines the steps needed to collect a viable nasal wipe from an individual pig.

The authors present a protocol to collect swine nasal wipes to detect and isolate influenza A viruses.

Surveillance for influenza A viruses in swine is critical to human and animal health because influenza A virus rapidly evolves in swine populations and ...

Neutrophil Isolation and Analysis to Determine their Role in Lymphoma Cell Sensitivity to Therapeutic Agents

Neutrophils are the most abundant (40% to 75%) type of white blood cells and among the first inflammatory cells to migrate towards the site of inflammation. They are key players in the innate immune system and play major roles in cancer biology. Neutrophils have been proposed as key mediators of malignant transformation, tumor progression, angiogenesis and in the modulation of the antitumor immunity; through their release of soluble factors or their interaction with tumor cells. To characterize the specific functions of neutrophils, a fast and reliable method is coveted for in vitro isolation of neutrophils from human blood. Here, a density gradient separation method is demonstrated to isolate neutrophils as well as mononuclear cells from the blood. The procedure consists of layering the density gradient solution such as Ficoll carefully above the diluted blood obtained from patients diagnosed with chronic lymphocytic leukemia (CLL), followed by centrifugation, isolation of mononuclear layer, separation of neutrophils from RBCsby dextran then lysis of residual erythrocytes. This method has been shown to isolate neutrophils &#8805; 90 % pure. To mimic the tumor microenvironment, 3-dimensional (3D) experiments were performed using basement membrane matrix such as Matrigel. Given the short half-life of neutrophils in vitro, 3D experiments with fresh human neutrophils cannot be performed. For this reason promyelocytic HL60 cells are differentiated along the granulocytic pathway using the differentiation inducers dimethyl sulfoxide (DMSO) and retinoic acid (RA). The aim of our experiments is to study the role of neutrophils on the sensitivity of lymphoma cells to anti-lymphoma agents. However these methods can be generalized to study the interactions of neutrophils or neutrophil-like cells with a large range of cell types in different situations.

Neutrophils are the most abundant type of white blood cells. The isolation of neutrophils from human blood by density gradient separation method and the differentiation of human promyelocytic (HL60) cells along the granulocytic pathway are described here; to test their role on sensitivity of lymphoma cells to anti-lymphoma agents.

Neutrophils are the most abundant (40% to 75%) type of white blood cells and among the first inflammatory cells to migrate towards the site of ...

Zika Virus Infectious Cell Culture System and the In Vitro Prophylactic Effect of Interferons

Zika Virus (ZIKV) is an emerging pathogen that is linked to fetal developmental abnormalities such as microcephaly, eye defects, and impaired growth. ZIKV is an RNA virus of the Flaviviridae family. ZIKV is mainly transmitted by mosquitoes, but can also be spread by maternal to fetal vertical transmission as well as sexual contact. To date, there are no reliable treatment or vaccine options available to protect those infected by the virus. The development of a reproducible, effective Zika virus infectious cell culture system is critical for studying the molecular mechanisms of ZIKV replication as well as drug and vaccine development. In this regard, a protocol describing a mammalian cell-based in vitro Zika virus culture system for viral production and growth analysis is reported here. Details on the formation of plaques by Zika virus on a cell monolayer and plaque assay for measuring viral titer are presented. Viral genome replication kinetics and double-stranded RNA genome replicatory intermediates are determined. This culture platform was utilized to screen against a library of a small set of cytokines resulting in the identification of interferon-&#945; (IFN-&#945;), IFN-&#946; and IFN-&#947; as potent inhibitors of Zika viral growth. In summary, an in vitro infectious Zika viral culture system and various virological assays are demonstrated in this study, which has the potential to greatly benefit the research community in elucidating further the mechanisms of viral pathogenesis and the evolution of viral virulence. Antiviral IFN-alpha can further be evaluated as a prophylactic, post-exposure prophylactic, and treatment option for Zika virus infections in high-risk populations, including infected pregnant women.

Zika Virus Infectious Cell Culture System and the In Vitro Prophylactic Effect of Interferons

Zika Virus (ZIKV), an emerging pathogen, is linked to fetal developmental abnormalities and microcephaly. The establishment of an effective infectious cell culture system is crucial for studies of ZIKV replication as well as vaccine and drug development. In this study, various virological assays pertaining to ZIKV are illustrated and discussed.

Zika Virus (ZIKV) is an emerging pathogen that is linked to fetal developmental abnormalities such as microcephaly, eye defects, and impaired growth. ZIKV ...

Rescue and Characterization of Recombinant Virus from a New World Zika Virus Infectious Clone

Infectious cDNA clones allow for genetic manipulation of a virus, thus facilitating work on vaccines, pathogenesis, replication, transmission and viral evolution. Here we describe the construction of an infectious clone for Zika virus (ZIKV), which is currently causing an explosive outbreak in the Americas. To prevent toxicity to bacteria that is commonly observed with flavivirus-derived plasmids, we generated a two-plasmid system which separates the genome at the NS1 gene and is more stable than full-length constructs that could not be successfully recovered without mutations. After digestion and ligation to join the two fragments, full-length viral RNA can be generated by in vitro transcription with T7 RNA polymerase. Following electroporation of transcribed RNA into cells, virus was recovered that exhibited similar in vitro growth kinetics and in vivo virulence and infection phenotypes in mice and mosquitoes, respectively.

This protocol describes the recovery of infectious Zika virus from a two-plasmid infectious cDNA clone.

Infectious cDNA clones allow for genetic manipulation of a virus, thus facilitating work on vaccines, pathogenesis, replication, transmission and viral ...

Isolation of Salmonella typhimurium-containing Phagosomes from Macrophages

Salmonella typhimurium is a facultative intracellular bacterium that causes gastroenteritis in humans. After invasion of the lamina propria, S. typhimurium bacteria are quickly detected and phagocytized by macrophages, and contained in vesicles known as phagosomes in order to be degraded. Isolation of S. typhimurium-containing phagosomes have been widely used to study how S. typhimurium infection alters the process of phagosome maturation to prevent bacterial degradation. Classically, the isolation of bacteria-containing phagosomes has been performed by sucrose gradient centrifugation. However, this process is time-consuming, and requires specialized equipment and a certain degree of dexterity. Described here is a simple and quick method for the isolation of S. typhimurium-containing phagosomes from macrophages by coating the bacteria with biotin-streptavidin-conjugated magnetic beads. Phagosomes obtained by this method can be suspended in any buffer of choice, allowing the utilization of isolated phagosomes for a broad range of assays, such as protein, metabolite, and lipid analysis. In summary, this method for the isolation of S. typhimurium-containing phagosomes is specific, efficient, rapid, requires minimum equipment, and is more versatile than the classical method of isolation by sucrose gradient-ultracentrifugation.

Isolation of Salmonella typhimurium-containing Phagosomes from Macrophages

We describe here a simple and quick method for the isolation of Salmonella typhimurium-containing phagosomes from macrophages by coating the bacteria with biotin and streptavidin.

Salmonella typhimurium is a facultative intracellular bacterium that causes gastroenteritis in humans. After invasion of the lamina propria, S. ...

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 ...

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 ...

Evaluation of Zika Virus-specific T-cell Responses in Immunoprivileged Organs of Infected Ifnar1-/- Mice

The Zika virus (ZIKV) can induce inflammation in immunoprivileged organs (e.g., the brain and testis), leading to the Guillain-Barr&#233; syndrome and damaging the testes. During an infection with the ZIKV, immune cells have been shown to infiltrate into the tissues. However, the cellular mechanisms that define the protection and/or immunopathogenesis of these immune cells during a ZIKV infection are still largely unknown. Herein, we describe methods to evaluate the virus-specific T-cell functionality in these immunoprivileged organs of ZIKV-infected mice. These methods include a) a ZIKV infection and vaccine inoculation in Ifnar1-/- mice; b) histopathology, immunofluorescence, and immunohistochemistry assays to detect the virus infection and inflammation in the brain, testes, and spleen; c) the preparation of a tetramer of ZIKV-derived T-cell epitopes; d) the detection of ZIKV-specific T cells in the monocytes isolated from the brain, testes, and spleen. Using these approaches, it is possible to detect the antigen-specific T cells that have infiltrated into the immunoprivileged organs and to evaluate the functions of these T cells during the infection: potential immune protection via virus clearance and/or immunopathogenesis to exacerbate the inflammation. These findings may also help to clarify the contribution of T cells induced by the immunization against ZIKV.

Evaluation of Zika Virus-specific T-cell Responses in Immunoprivileged Organs of Infected Ifnar1-/- Mice

A protocol to evaluate antigen-specific T-cell responses in the immunoprivileged organs of the Ifnar1-/- murine model for the Zika virus (ZIKV) infection is described. This method is pivotal for investigating the cellular mechanisms of the protection and immunopathogenesis of ZIKV vaccines and is also valuable for their efficacy evaluation.

The Zika virus (ZIKV) can induce inflammation in immunoprivileged organs (e.g., the brain and testis), leading to the Guillain-Barr&#233; syndrome and ...

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 ...

Isolation and Quantification of Zika Virus from Multiple Organs in a Mouse

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Isolation of Lymphocytes from Mouse Genital Tract Mucosa

Nasal Wipes for Influenza A Virus Detection and Isolation from Swine

Neutrophil Isolation and Analysis to Determine their Role in Lymphoma Cell Sensitivity to Therapeutic Agents

Zika Virus Infectious Cell Culture System and the In Vitro Prophylactic Effect of Interferons

Rescue and Characterization of Recombinant Virus from a New World Zika Virus Infectious Clone

Isolation of Salmonella typhimurium-containing Phagosomes from Macrophages

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

Evaluation of Zika Virus-specific T-cell Responses in Immunoprivileged Organs of Infected Ifnar1^-/- Mice

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