We would like to thank members of the Krachler group for critical reading and comments on the manuscript. This work was funded by a UT Systems STAR award, the BBSRC, and the NIH (R01AI132354).&#160;

Zebrafish care, breeding, and experiments described here are in accordance with the Guide for the Care and Use of Laboratory Animals, and have been approved by the Institutional Animal Welfare Committee of the University of Texas Health Science Center, protocol number AWC-16-0127.
1. Growth and Maintenance of Paramecia
<ol>
	<li>Obtain live paramecia cultures from sources such as the Zebrafish International Resource Center (ZIRC).</li>
	<li>From a live growing culture, add 1 mL of the paramecia culture, 1 mL of an E. coli MG1655 culture (optical density OD600 1.0&#8211;2.0) resuspended in 1x E3 (0.29 g/L NaCl, 13 mg/L KCl, 44 mg/L CaCl2, 81 mg/L MgSO4, 0.48 g/L HEPES, pH 7.0, sterile), and 8 mL of 1x E3 into a 10 mL tissue culture flask. Swirl lightly and then store at 22 &#176;C.</li>
	<li>To maintain a culture, every two weeks, passage 1 mL of a paramecia culture into a 10 mL tissue culture flask with 9 mL of fresh 1x E3 medium containing 108 CFU/mL of E. coli MG1655 resuspended in 1x E3.</li>
</ol>
2. Determination of Bacterial Dose Administered to Zebrafish
<ol>
	<li>Determine bacterial half-life within Paramecium 
	NOTE: The half-life of bacteria within paramecia is determined by plating viable E. coli recovered from paramecium, as described below.
	<ol>
		<li>Following a 2 h incubation of bacteria (OD600 of 1.0) with paramecia at 22 &#176;C, combine the paramecia and bacterial co-culture into a 50 mL conical tube, wash the paramecia with 1x E3, and count the number of paramecia per milliliter (as described below in steps 3.2.7 and 3.2.8).</li>
		<li>After counting the number of paramecia, remove 50 &#181;L of the paramecia and bacterial co-culture every hour (for 6 h post-exposure) and add each sample into a fresh 1.5 mL tube.</li>
		<li>Place 950 &#181;L of 1% nonionic surfactant (see Table of Materials) in phosphate buffered saline (PBS) solution into each of the 1.5 mL tubes and vortex for 1 min to lyse the paramecia. Perform 1:10 dilutions of each sample using sterile PBS as a diluent (i.e., 100 &#181;L in 900 &#181;L).</li>
		<li>Plate 100 &#181;L of each dilution onto selective plates (LB tet medium: 1 g/L tryptone, 0.5 g/L yeast extract, 1 g/L NaCl, 30 &#181;g/mL tetracycline) and incubate at 37 &#176;C for 16 h. The next day, count and record the number of bacterial colonies on the plate (colony forming units, CFUs) by counting only the isolated and distinct individual colonies. Identify a plate with a dilution that gives 30&#8211;300 CFU.</li>
		<li>Determine the dilution factor used. For example, if 1 &#181;L of the bacterial culture is mixed with 99 mL of sterile PBS, this is a 1:100 (0.01) dilution. This number will be the number of CFU in the dilution. Perform the calculation below, for every time point, to determine the CFU in the original sample: 
		<img alt="Equation 1" src="/files/ftp_upload/58949/58949eq1.jpg" style="opacity: 0.9;" /></li>
		<li>Calculate and graph the number of viable E. coli/paramecium corresponding to the hours post exposure using the paramecia concentration calculated in step 3.3.1 and determine the half-life within the paramecia (Figure 1).</li>
		<li>Using the CFU number calculated for each time point, calculate and graph the number of viable E. coli per paramecium on the y-axis versus the hours post incubation on the x-axis, using the paramecia concentration calculated in step 3.3.1. Determine the half-life within the paramecia.</li>
	</ol>
	</li>
	<li>Determine bacterial dosing from bacterial half-life and preying rate. 
	NOTE: To determine the bacterial dosage, the half-life of the bacteria inside the paramecia and the preying rate of zebrafish on paramecia (see step 3.4) have to be taken into account.
	<ol>
		<li>Use the following formula to determine bacterial decay within paramecia: 
		(1)&#160;&#160;<img alt="Equation 2" src="/files/ftp_upload/58949/58949eq2.jpg" style="opacity: 0.9;" /> 
		Where N0 is the initial quantity of bacteria per paramecia after the 2 h incubation, Nt is the remaining quantity after time t, &#964; is the time after which the number of viable bacteria has halved, and k is the decay constant.</li>
		<li>From the degradation experiment, determine the decay constant k using the bacterial half-life (i.e., the time after which the amount of viable bacteria/paramecium has halved). 
		(2)&#160;&#160;<img alt="Equation 3" src="/files/ftp_upload/58949/58949eq3.jpg" style="opacity: 0.9;" /> 
		For the determination of half-life, the term <img alt="Equation 4" src="/files/ftp_upload/58949/58949eq4.jpg" style="opacity: 0.9;" />&#160;and 
		(3)&#160;&#160;<img alt="Equation 5" src="/files/ftp_upload/58949/58949eq5.jpg" style="opacity: 0.9;" />&#160;or 
		(4)&#160;&#160;<img alt="Equation 6" src="/files/ftp_upload/58949/58949eq6.jpg" style="opacity: 0.9;" /> 
		NOTE: Based on Figure 1, the half-life of E. coli in paramecia is approximately 2.3 h. Thus, using formula (4), the decay rate, k, for E. coli is: 
		(5)&#160;&#160;<img alt="Equation 7" src="/files/ftp_upload/58949/58949eq7.jpg" style="opacity: 0.9;" /></li>
		<li>Determine the decay rate (k) from the half-life experiment to find the dose of viable bacteria (Nt) taken up by the zebrafish larvae after preying time (t), where (P) is the preying rate or the number of paramecia eaten by one fish per hour: 
		(6)&#160;&#160;<img alt="Equation 8" src="/files/ftp_upload/58949/58949eq8.jpg" style="opacity: 0.9;" />&#160;or 
		(7)&#160; <img alt="Equation 9" src="/files/ftp_upload/58949/58949eq9.jpg" style="opacity: 0.9;" /> 
		NOTE: Per Figure 1, the initial quantity of bacteria per paramecia (N0) after a 2 h incubation (t) is 790 CFU. Per Movie 1 and Figure 2, the preying rate (P) is 1539.</li>
		<li>Using these values to substitute into equation (7), calculate the bacterial dosage consumed by a zebrafish following a 2 h incubation as: 
		(8)&#160; <img alt="Equation 10" src="/files/ftp_upload/58949/58949eq10.jpg" style="opacity: 0.9;" /></li>
	</ol>
	</li>
</ol>
3. Food-borne Infection of Zebrafish
<ol>
	<li>Incubate bacteria with Paramecia.
	<ol>
		<li>Prepare a co-culture of paramecia and E. coli MG1655 the night prior to infection. Combine 8 mL of E3 media, 1 mL of an ongoing paramecia culture, and 1 mL of an E. coli MG1655 culture (OD600 = 1.0) resuspended in 1x E3 in T25 tissue culture flasks. Incubate the flasks at room temperature (RT) overnight. For each treatment condition, prepare two flasks of paramecia.</li>
		<li>Inoculate bacterial growth media (LB: 1 g/L tryptone, 0.5 g/L yeast extract, 1 g/L NaCl) with infectious strain of bacteria, by picking an individual bacterial colony from a plate using a sterile inoculation loop.&#160;Incubate the liquid culture at 37 &#176;C and leave shaking at 110 rotations per minute (rpm) overnight. 
		NOTE: Personal protective equipment (a laboratory coat and gloves) should be worn and biosafety level 2 facilities should be used when handling infectious agents.</li>
		<li>On the next day, measure the OD600 of the overnight culture. Calculate the volume of culture required to achieve an OD600 of 1 when resuspended in 11 mL of media.</li>
		<li>Harvest the volume of bacteria from step 3.1.2 via centrifugation at 6,000 x g for 5 min, one volume for each flask of paramecia. Discard the supernatant and resuspend the bacterial pellet in 1 mL of E3 media.</li>
		<li>Optionally, pre-stain the bacteria with a fluorescent dye.
		<ol>
			<li>Add 1 &#181;L of FM 4-64FX bacterial stain (5 mg/mL stock solution). Cover tube with foil to protect from photobleaching and incubate rotating end-over-end at RT for 15 min.</li>
			<li>Remove excess dye by washing with 1x E3: Pellet bacteria via centrifugation at 6,000 x g for 1.5 min, then resuspend pellet in 1 mL of E3 media. Repeat wash step two times.</li>
			<li>Harvest stained bacteria via centrifugation at 6,000 x g for 5 min. Discard the supernatant and resuspend the bacterial pellet in 1 mL of E3 media.</li>
		</ol>
		</li>
		<li>Add 1 mL of the bacterial suspensions to each of the two flasks of fresh paramecia. Incubate at RT for 2 h. 
		NOTE: If working with stained bacteria, incubate in the dark at RT for 2 h.</li>
	</ol>
	</li>
	<li>Wash bacteria/paramecia co-culture.
	<ol>
		<li>Combine the contents of both flasks of paramecia/bacteria co-culture into a 50 mL conical tube. Centrifuge samples at 300 x g&#160;at 15 &#176;C for 10 min. Make sure that the centrifuge is pre-cooled prior to this step.</li>
		<li>Remove approximately 10 mL of the E3 supernatant using a serological pipette and add approximately 10 mL of fresh 1x E3 to the conical tube. 
		NOTE: During all wash steps, it is essential to be very quick when removing the supernatant, as the paramecia will begin to swim out of the pellet. Spin and remove supernatant from one tube at a time to ensure quick enough handling at this step, and avoid loss of paramecia in the supernatant.</li>
		<li>Spin samples via centrifugation at 300 x g at 15 &#176;C for 5 min. Remove approximately 10 mL of the E3 supernatant using a serological pipette, and add approximately 10 mL of fresh 1x E3 to the conical tube. Repeat this step twice.</li>
		<li>Centrifuge samples at 300 x g&#160;at 15 &#176;C for 5 min. Remove approximately 10 mL of the E3 supernatant, taking care not to disrupt the pellet.</li>
		<li>Resuspend pellet into the remaining 10 mL of E3 media and transfer 500 &#181;L of the suspension into a new 1.5 mL microcentrifuge tube. Pellet the 500 &#181;L of paramecia by centrifuging at 300 x g for 5 min to count the number of paramecia.</li>
		<li>Remove 400 &#181;L of the E3 supernatant from the 500 &#181;L sample. Add 20 &#181;L of 36.5% formaldehyde solution to the remaining 100 &#181;L of paramecia and gently resuspend, and incubate for 5 min at 22 &#176;C. 
		NOTE: This step kills the paramecia to allow for counting.</li>
		<li>Measure actual total volume using the pipette and record. Dilute the paramecia suspension 1:1 v/v with 0.4% trypan blue solution.</li>
		<li>Use a cell counter or hemocytometer to count the number of dead paramecia/mL. 
		NOTE: Because of the prior fixation step, most paramecia will be dead at this point, but this number reflect the number of live paramecia for the co-incubation experiment. The authors have not found significant paramecia death due to bacterial co-incubation, so this can be disregarded as a factor here.</li>
	</ol>
	</li>
	<li>Co-incubate Paramecia and zebrafish larvae.
	<ol>
		<li>Calculate the concentration of paramecia: 
		<img alt="Equation 11" src="/files/ftp_upload/58949/58949eq11.jpg" style="opacity: 0.9;" /> 
		<img alt="Equation 12" src="/files/ftp_upload/58949/58949eq12.jpg" /> 
		NOTE: This calculation gives the concentration of paramecia in the 50 mL conical tube from step 3.2.5.</li>
		<li>Calculate the volume of washed paramecia required for a concentration of 2 x 105 paramecia/mL in a final volume of 3 mL of E3. 
		NOTE: The concentration of paramecia can be adjusted based on the desired bacterial dosage, which is subject to optimization.</li>
		<li>Anesthetize zebrafish by adding tricaine in 100 mM Tris pH 8.0 to a final concentration of 100 mg/L. Transfer 10 zebrafish into each well of a 6-well plate into a total volume of 3 mL of fresh E3 containing the appropriate concentration of paramecia (calculated in step 3.3.2). Ensure to transfer larvae in a minimal amount of liquid, to ensure they recover from anesthesia in the recipient well.</li>
		<li>Incubate at 30 &#176;C for 2 h in a diurnal incubator under day-light conditions, to ensure optimal lightning conditions for preying.</li>
		<li>Wash zebrafish at least 5 times by transferring fish into a new well containing 3 mL of fresh E3 containing 100 mg/L tricaine each time. 
		NOTE: Do not attempt to omit the tricaine during the washing step. Transferring mobile larvae without anesthesia increases the risk of damage and distress to the animal.</li>
		<li>Optionally, prepare zebrafish for imaging by embedding zebrafish in 3 mL of 1% low-melt agarose in a black-walled 6-well plate: Low-melt agarose is made up in 1xE3 and heated in a microwave. Once molten, add tricaine to a final concentration of 160 mg/mL. Position fish under a stereomicroscope, using a clipped gel loading tip, making sure that the head is on the left and the tail is on the right (Figure 3). Wait for 5 minutes for the agarose to set, then overlay the embedded fish with 1x E3 containing 160 mg/mL tricaine for imaging.</li>
	</ol>
	</li>
	<li>Determine the preying rate. 
	NOTE: No separate experiment needs to be set up to determine the preying rate. Rather, this can be done during step 3.3.4, as described below.
	<ol>
		<li>During step 3.3.4, view preying zebrafish on a stereomicroscope and capture video footage of the prey capture.</li>
		<li>Score the video footage. Prey capture is characterized by striking of zebrafish toward the prey. Count each strike as one prey capture event, although this is only an approximation (see Discussion).</li>
		<li>Calculate the average number of prey capture events per hour from multiple video clips, each representing a different zebrafish larva (Figure 2).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The basic protocol described here has been optimized for pathogenic E. coli, and has been successfully adapted for other bacterial species, including Salmonella enterica and Vibrio cholerae. For some species that do not colonize the zebrafish gut following bath immersion, including some Salmonella enterica strains and some anaerobes, food-borne infection as described here can be used to successfully establish colonization. Compared to microgavage, which is also used to establish high bacterial burdens in the larval intestinal tract, food-borne infection is technically less challenging and requires less specialized equipment. However, critical parameters should be optimized for the bacterial species and strains to be used. Such factors include bacterial and paramecium density for the bacteria-paramecium co-culture step: If bacterial numbers within paramecia are low, this could be improved by increasing the bacterial density in the co-culture step. Some bacterial species may cause damage to the paramecium host, and this should be assessed by microscopy.&#160;
Another important factor in this protocol is prey capture by zebrafish. The preying rate as described here is based on the assumption that every prey capture strike results in the ingestion of one paramecium. High densities of paramecia per fish are used in the protocol to ensure high preying rates. However, prey capture is dependent on the density of paramecia in the system, and in very dilute paramecium cultures, preying rates may be as low as 13&#8211;15 paramecia per hour21,22. A limitation is that prey capture rates are also strongly dependent on lighting conditions and in the dark, capture rates are 80% lower than in light conditions21 and this should be taken into account when setting up experiments. If exposure times to prey have to be expanded to optimize colonization, consideration has to be given to secondary exposure to bacteria through feces. Under the conditions described above &#8211; 2 h of prey exposure &#8211; this exposure is negligible, since gut passage time of bacteria is more than 1 h and the concentration of bacteria in the vehicle is much higher than in feces. However, if prey exposure time is significantly increased, this may become a significant factor.&#160;
Appropriate controls should be included in this protocol, including colonization of zebrafish following feeding with paramecia containing non-pathogenic E. coli MG1655. If multiple bacterial strains are compared for their ability to colonize the zebrafish host, it is important to test whether their half-life within paramecia is comparable. Bacterial mutations, including those compromising cell wall integrity or acid sensing, may compromise bacterial stability within paramecia. In such cases, zebrafish feeding has to be adjusted to account for the differences in dosage.&#160;
The protocol described here can be used to investigate bacterial colonization and its consequences, including by imaging bacterial colonization of zebrafish as described above, as well as by determining CFU per zebrafish from tissue homogenate3, or investigating infection-associated morbidity and mortality. Ideally, for bacterial visualization, bacterial strains expressing fluorescent proteins such as mCherry or red fluorescent protein (RFP) should be used. This will allow the visualization of growing bacterial populations. If the bacterial strain is not genetically tractable or the use of fluorescent protein expression is precluded for other reasons, bacteria may be stained with a fluorescent dye, such as FM 4-64FX, prior to co-culture with paramecia. When using the protocol described here, co-culture with paramecia does not decrease the brightness of the dye and stained bacteria are clearly visible in the zebrafish intestine. However, the dye will be diluted over time should significant bacterial proliferation occur within the zebrafish host. In either case, red-fluorescent bacteria are preferable over green-fluorescent bacteria, since tissue autofluorescence can be higher in the green than in the red channel.&#160;
It has been found that this protocol can be adapted for aerobic and microaerophilic bacterial species. It may be possible to adapt it for the feeding of spores and fungal species, although this remains to be tested experimentally.

Zebrafish share morphologically and functionally conserved features with mammals, including granulocytic lineages (e.g., neutrophils), monocyte/macrophage-like cells, Toll-like receptors, pro-inflammatory cytokines, and antimicrobial peptides1. The intestinal tract in zebrafish is fully developed at 6 days post fertilization (dpf) and shows morphological and functional conservation with the mammalian gastrointestinal tract, such as conserved transcriptional regulation in intestinal epithelial cells2. This makes zebrafish an excellent model for intestinal microbial colonization and pathogenesis. A wide range of enteric microbes has been studied in the zebrafish model, including enterohemorrhagic Escherichia coli3, Vibrio cholerae4,5, Salmonella enterica6, the zebrafish microbiota7,8, and the role of probiotics in intestinal immunity9. A distinct advantage of the zebrafish model is that it is colonized by many microbes without disrupting the endogenous microbiota, which allows the investigation of microbial behavior in the context of mixed microbial populations3,6. Currently, most zebrafish models of gastrointestinal colonization and disease rely on the administration of microbes by bath immersion, where zebrafish are incubated in a bacterial suspension for a specific amount of time10. However, this makes it difficult to determine the exact dose of bacteria administered, and leads to limited colonization with some microbes, particularly with non-pathogenic bacteria. Alternatively, a bacterial suspension is administered to fish via oral gavage11, but this is technically challenging and limited to older larvae and adult fish.
This protocol describes the use of the unicellular protozoan Paramecium caudatum as a vehicle for food-borne delivery of microbes to the gastrointestinal tract of zebrafish larvae. Paramecia are easy and cheap to maintain and are capable of feeding on a wide variety of microbes, including algae, fungi, and bacteria, which they internalize through a ciliated oral groove12,13,14. Once internalized, bacteria are held in vacuoles, which eventually acidify and contents are degraded over a time frame of several hours15. Larval zebrafish capture paramecia as natural prey soon after hatching, around 3&#8211;4 dpf depending on temperature16, and take them up with high efficiency. The process of prey capture takes on average 1.2 s from detection to capture17, and captured paramecia are quickly digested in the zebrafish foregut, such that internalized viable bacteria are released into the intestinal tract3. As a result, paramecia can be used as a quick and easy method to deliver a high and consistent dose of bacteria into the gastrointestinal tract of zebrafish. The delivered bacteria can either be transformed to express a fluorescent protein, such as mCherry as described here, or, in the case of genetically intractable bacteria, they can be pre-stained with a fluorescent dye to allow visualization within the gastrointestinal tract.
This protocol describes the food-borne delivery of enteropathogenic E. coli (enterohemorrhagic E. coli [EHEC] and adherent invasive E. coli [AIEC]), and Salmonella enterica ssp. Typhimurium. Both pathogenic E. coli and S. typhimurium are transmitted via the fecal-oral route18,19, and can be acquired via contaminated food, such as meat, vegetables, and dairy. Using P. caudatum as a vehicle, E. coli and S. typhimurium successfully colonize the zebrafish larvae within 30&#8211;60 min of co-incubation with the paramecium vehicle. The achieved bacterial burden is robust enough to visualize colonization and determine burden by plating tissue homogenates.

<table border="0" cellpadding="0" cellspacing="0" style="width:933px;" width="932">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">Paramecium caudatum, live</td>
			<td style="width:133px;">Carolina</td>
			<td style="width:123px;">131554</td>
			<td style="width:411px;">no not store growing cultures below room temperature</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">0.4% Trypan Blue Solution&#160;</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:123px;">T8154-20ML</td>
			<td>liquid, sterile-filtered, suitable for cell culture; prepared in 0.81% sodium chloride and 0.06% potassium phosphate, dibasic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">Dimethyl sulfoxide (DMSO)</td>
			<td style="width:133px;">Sigma</td>
			<td>276855-100ML</td>
			<td>store in a solvent safety cabinet</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">Escherichia coli, MG1655</td>
			<td style="width:133px;">ATCC</td>
			<td style="width:123px;">ATCC 700926</td>
			<td>can be replaced by any other non-pathogenic E. coli strain</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">FM 4-64FX stain</td>
			<td style="width:133px;">Thermo Fisher</td>
			<td style="width:123px;">F34653</td>
			<td>aliquot and store frozen</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">Formaldehyde</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:123px;">F8775-4X25ML</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">LB Broth</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:123px;">L3397-1KG</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">Phosphate buffered saline tablets</td>
			<td style="width:133px;">Thermo Fisher</td>
			<td style="width:123px;">18912014</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">Tetracycline</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:123px;">87128-25G</td>
			<td>toxic, irritant</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:267px;">Tricaine (Ethyl 3-aminobenzoate methanesulfonate)</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:123px;">E10521-10G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">Triton X-100</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:123px;">X100-100ML</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">Trypan Blue Solution, 0.4%</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:123px;">93595-50ML</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">UltraPure Low Melting Point Agarose</td>
			<td style="width:133px;">Thermo Fisher</td>
			<td style="width:123px;">16520050</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">hemocytometer or cell counter</td>
			<td style="width:133px;">any</td>
			<td style="width:123px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">stereomicroscope</td>
			<td style="width:133px;">any</td>
			<td style="width:123px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">table-top centrifuge</td>
			<td style="width:133px;"></td>
			<td style="width:123px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">microwave</td>
			<td style="width:133px;"></td>
			<td style="width:123px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">rotator wheel</td>
			<td style="width:133px;"></td>
			<td style="width:123px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">heated shaking incubator</td>
			<td style="width:133px;"></td>
			<td style="width:123px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">aquatics facilities</td>
			<td style="width:133px;"></td>
			<td style="width:123px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">breeding tanks</td>
			<td style="width:133px;"></td>
			<td style="width:123px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

using the protozoan paramecium caudatum as a vehicle for food-borne infections in zebrafish larvae

Due to their transparency, genetic tractability, and ease of maintenance, zebrafish (Danio rerio) have become a widely-used vertebrate model for infectious diseases. Larval zebrafish naturally prey on the unicellular protozoan Paramecium caudatum. This protocol describes the use of P. caudatum as a vehicle for food-borne infection in larval zebrafish. P. caudatum internalize a wide range of bacteria and bacterial cells remain viable for several hours. Zebrafish then prey on P. caudatum, the bacterial load is released in the foregut upon digestion of the paramecium&#160;vehicle, and the bacteria colonize the intestinal tract. The protocol includes a detailed description of paramecia maintenance, loading with bacteria, determination of bacterial degradation and dose, as well as infection of zebrafish by feeding with paramecia. The advantage of using this method of food-borne infection is that it closely mimics the mode of infection observed in human disease, leads to more robust colonization compared to immersion protocols, and allows the study of a wide range of pathogens. Food-borne infection in the zebrafish model can be used to investigate bacterial gene expression within the host, host-pathogen interactions, and hallmarks of pathogenicity including bacterial burden, localization, dissemination and morbidity.

Paramecium caudatum readily internalizes a wide range of bacteria into storage vacuoles. The intracellular bacterial density depends on the densities of bacteria and paramecia in the co-culture, as well as the bacterial species used. Over time, the vacuoles acidify and bacterial degradation ensues. The rate of degradation has to be individually determined for all strains used. For pathogenic E. coli, the initial bacterial density is 790 bacteria/paramecium, and bacteria are degraded with a half-life of approximately 2.3 h (Figure 1).&#160;
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58949/58949fig1.jpg" /> 
Figure 1: Determination of bacterial half-life in paramecia. (A) Following 2 h of co-incubation with infectious E. coli, P. caudatum was washed and transferred to medium without bacteria. At the indicated time points, numbers of viable E. coli cells were determined by dilution plating on selective agar. Results are means &#177; standard error of the mean (SEM; n = 3). (B) Typical image of paramecium carrying internalized bacteria, with bright field (Bi), fluorescent bacteria (Bii), and merged channels (Biii). Scale bar = 20 &#956;m.&#160;<a href="https://www.jove.com/files/ftp_upload/58949/58949fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Further, the zebrafish preying rate, that is, the rate at which zebrafish internalize bacteria-loaded paramecia upon co-incubation, was studied. Larval zebrafish start to hunt and capture live prey from 5 dpf20, although it was found that, when raised at 30 &#176;C, larval development is accelerated and animals display preying behavior from 4 dpf. Preying is accompanied by a characteristic striking behavior20 (Figure 2A), and the determination of the preying rate is based on the assumption that each strike leads to internalization of one paramecium, although this can only be regarded an approximation (see&#160;Discussion). Based on the herein described observations of preying zebrafish larvae, the rate of paramecia uptake is approximately 1,539 per h (Figure 2B).&#160;
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58949/58949fig2.jpg" /> 
Figure 2: Determination of zebrafish preying rate. (A) Still images from a preying video, showing a zebrafish larvae (5 dpf) preying on paramecia carrying fluorescent bacteria. Time in [seconds]. Arrow indicates the main axis of movement during striking. (B) Quantification of preying rate (paramecia intake per hour), based on n = 10 videos taken over the full 2 h exposure time.&#160;<a href="https://www.jove.com/files/ftp_upload/58949/58949fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Following internalization of paramecia, the zebrafish efficiently degrades the prey in the foregut, releasing infectious bacteria into the digestive system. As described herein, paramecia degradation proceeds quickly, and free bacteria can be detected in the intestinal tract within 30 minutes of preying. Free bacteria then move from the foregut to the mid- and posterior intestine, where they are detected approximately 1&#8211;2 h after the beginning of preying (Figure 3). Bacterial persistence in the intestine depends on species and dose but ranges from several h to several days in the case of E. coli and S. enterica. S. enterica localizes primarily in the intestinal mucosae, with some epithelial invasion (Figure 3D), leading to infiltration of neutrophils into the epithelium (Figure 3C).&#160;
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58949/58949fig3.jpg" /> 
Figure 3: Colonization of zebrafish with bacteria. Zebrafish at 5 dpf were left uninfected (A) or colonized with mCherry expressing (B) E. coli or (C) S. enterica. Infection experiments may be performed in wild type (A and B) fish or transgenic lines (e.g., the line Tg(MPO::EGFP)i114 expressing green fluorescent neutrophils shown in (C). The rectal opening is marked by an arrow. (D) Higher magnification of intestinal section from whole-mount embedded larvae infected with Salmonella enterica infection. (Di) Blue = Hoechst marking nuclei, (Dii) Purple = phalloidin marking F-actin, (Diii) Red = Salmonella, (Div) merge. Scale bar = 5 &#956;m.&#160;<a href="https://www.jove.com/files/ftp_upload/58949/58949fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Movie 1" src="/files/ftp_upload/58949/58949mov1.jpg" /> 
Movie 1: Video footage of the prey capture.&#160;<a href="https://www.jove.com/files/ftp_upload/58949/PreyCapture.mp4" target="_blank">Please click here to download this video.</a>

Watch this Scientific Journal Video about Using the Protozoan Paramecium caudatum as a Vehicle for Food-borne Infections in Zebrafish Larvae at JoVE.com

Using the Protozoan Paramecium caudatum as a Vehicle for Food-borne Infections in Zebrafish Larvae

Zebrafish (Danio rerio) are becoming a widely-used vertebrate animal model for microbial colonization and pathogenesis. This protocol describes the use of the protozoan Paramecium caudatum as a vehicle for food-borne infection in zebrafish larvae. P. caudatum readily internalizes bacteria and get taken up by larval zebrafish through natural preying behavior.

Using the Protozoan Paramecium caudatum as a Vehicle for Food-borne Infections in Zebrafish Larvae

Due to their transparency, genetic tractability, and ease of maintenance, zebrafish (Danio rerio) have become a widely-used vertebrate model for ...

using-protozoan-paramecium-caudatum-as-vehicle-for-food-borne

Research

JoVE Journal

Immunology and Infection

28.3K Views.  University of Texas Health Science Center at Houston. Zebrafish (Danio rerio) are becoming a widely-used vertebrate animal model for microbial colonization and pathogenesis. This protocol describes the use of the protozoan Paramecium caudatum as a vehicle for food-borne infection in zebrafish larvae. P. caudatum readily internalizes bacteria and get taken up by larval zebrafish through natural preying behavior.

Video: Using the Protozoan Paramecium caudatum as a Vehicle for Food-borne Infections in Zebrafish Larvae

Using Luciferase to Image Bacterial Infections in Mice

Imaging is a valuable technique that can be used to monitor biological processes. In particular, the presence of cancer cells, stem cells, specific immune cell types, viral pathogens, parasites and bacteria can be followed in real-time within living animals 1-2. Application of bioluminescence imaging to the study of pathogens has advantages as compared to conventional strategies for analysis of infections in animal models3-4. Infections can be visualized within individual animals over time, without requiring euthanasia to determine the location and quantity of the pathogen. Optical imaging allows comprehensive examination of all tissues and organs, rather than sampling of sites previously known to be infected. In addition, the accuracy of inoculation into specific tissues can be directly determined prior to carrying forward animals that were unsuccessfully inoculated throughout the entire experiment. Variability between animals can be controlled for, since imaging allows each animal to be followed individually. Imaging has the potential to greatly reduce animal numbers needed because of the ability to obtain data from numerous time points without having to sample tissues to determine pathogen load3-4.
This protocol describes methods to visualize infections in live animals using bioluminescence imaging for recombinant strains of bacteria expressing luciferase. The click beetle (CBRLuc) and firefly luciferases (FFluc) utilize luciferin as a substrate5-6. The light produced by both CBRluc and FFluc has a broad wavelength from 500 nm to 700 nm, making these luciferases excellent reporters for the optical imaging in living animal models7-9. This is primarily because wavelengths of light greater than 600 nm are required to avoid absorption by hemoglobin and, thus, travel through mammalian tissue efficiently. Luciferase is genetically introduced into the bacteria to produce light signal10. Mice are pulmonary inoculated with bioluminescent bacteria intratracheally to allow monitoring of infections in real time. After luciferin injection, images are acquired using the IVIS Imaging System. During imaging, mice are anesthetized with isoflurane using an XGI-8 Gas Anethesia System. Images can be analyzed to localize and quantify the signal source, which represents the bacterial infection site(s) and number, respectively. After imaging, CFU determination is carried out on homogenized tissue to confirm the presence of bacteria. Several doses of bacteria are used to correlate bacterial numbers with luminescence. Imaging can be applied to study of pathogenesis and evaluation of the efficacy of antibacterial compounds and vaccines.

Methods for bioluminescence imaging of bacterial infections in living animals are decribed. Pathogens are modified to express luciferase allowing optical whole body imaging of infections in live animals. Animal models can be infected with luciferase expressing pathogens and the resulting course of disease visualized in real-time by bioluminescence imaging.

Imaging is a valuable technique that can be used to monitor biological processes.  In particular, the presence of cancer cells, stem cells, specific ...

Introducing Shear Stress in the Study of Bacterial Adhesion

During bacterial infections a sequence of interactions occur between the pathogen and its host. Bacterial adhesion to the host cell surface is often the initial and determining step of the pathogenesis. Although experimentally adhesion is mostly studied in static conditions adhesion actually takes place in the presence of flowing liquid. First encounters between bacteria and their host often occur at the mucosal level, mouth, lung, gut, eye, etc. where mucus flows along the surface of epithelial cells. Later in infection, pathogens occasionally access the blood circulation causing life-threatening illnesses such as septicemia, sepsis and meningitis. A defining feature of these infections is the ability of these pathogens to interact with endothelial cells in presence of circulating blood. The presence of flowing liquid, mucus or blood for instance, determines adhesion because it generates a mechanical force on the pathogen. To characterize the effect of flowing liquid one usually refers to the notion of shear stress, which is the tangential force exerted per unit area by a fluid moving near a stationary wall, expressed in dynes/cm2. Intensities of shear stress vary widely according to the different vessels type, size, organ, location etc. (0-100 dynes/cm2). Circulation in capillaries can reach very low shear stress values and even temporarily stop during periods ranging between a few seconds to several minutes 1. On the other end of the spectrum shear stress in arterioles can reach 100 dynes/cm2 2. The impact of shear stress on different biological processes has been clearly demonstrated as for instance during the interaction of leukocytes with the endothelium 3. To take into account this mechanical parameter in the process of bacterial adhesion we took advantage of an experimental procedure based on the use of a disposable flow chamber 4. Host cells are grown in the flow chamber and fluorescent bacteria are introduced in the flow controlled by a syringe pump. We initially focused our investigations on the bacterial pathogen Neisseria meningitidis, a Gram-negative bacterium responsible for septicemia and meningitis. The procedure described here allowed us to study the impact of shear stress on the ability of the bacteria to: adhere to cells 1, to proliferate on the cell surface 5and to detach to colonize new sites 6 (Figure 1). Complementary technical information can be found in reference 7. Shear stress values presented here were chosen based on our previous experience1 and to represent values found in the literature. The protocol should be applicable to a wide range of pathogens with specific adjustments depending on the objectives of the study.

During the infection process, a key step is the adhesion of pathogens with host cells. In most instances this adhesion step occurs in the presence of mechanical stress generated by flowing liquid. We describe a technique that introduces shear stress as an important parameter in the study of bacterial adhesion.

During bacterial infections a sequence of interactions occur between the pathogen and its host. Bacterial adhesion to the host cell surface is often the ...

Non-invasive Imaging of Disseminated Candidiasis in Zebrafish Larvae

Disseminated candidiasis caused by the pathogen Candida albicans is a clinically important problem in hospitalized individuals and is associated with a 30 to 40% attributable mortality6. Systemic candidiasis is normally controlled by innate immunity, and individuals with genetic defects in innate immune cell components such as phagocyte NADPH oxidase are more susceptible to candidemia7-9. Very little is known about the dynamics of C. albicans interaction with innate immune cells in vivo. Extensive in vitro studies have established that outside of the host C. albicans germinates inside of macrophages, and is quickly destroyed by neutrophils10-14. In vitro studies, though useful, cannot recapitulate the complex in vivo environment, which includes time-dependent dynamics of cytokine levels, extracellular matrix attachments, and intercellular contacts10, 15-18. To probe the contribution of these factors in host-pathogen interaction, it is critical to find a model organism to visualize these aspects of infection non-invasively in a live intact host. 
The zebrafish larva offers a unique and versatile vertebrate host for the study of infection. For the first 30 days of development zebrafish larvae have only innate immune defenses2, 19-21, simplifying the study of diseases such as disseminated candidiasis that are highly dependent on innate immunity. The small size and transparency of zebrafish larvae enable imaging of infection dynamics at the cellular level for both host and pathogen. Transgenic larvae with fluorescing innate immune cells can be used to identify specific cells types involved in infection22-24. Modified anti-sense oligonucleotides (Morpholinos) can be used to knock down various immune components such as phagocyte NADPH oxidase and study the changes in response to fungal infection5. In addition to the ethical and practical advantages of using a small lower vertebrate, the zebrafish larvae offers the unique possibility to image the pitched battle between pathogen and host both intravitally and in color. 
The zebrafish has been used to model infection for a number of human pathogenic bacteria, and has been instrumental in major advances in our understanding of mycobacterial infection3, 25. However, only recently have much larger pathogens such as fungi been used to infect larva5, 23, 26, and to date there has not been a detailed visual description of the infection methodology. Here we present our techniques for hindbrain ventricle microinjection of prim25 zebrafish, including our modifications to previous protocols. Our findings using the larval zebrafish model for fungal infection diverge from in vitro studies and reinforce the need to examine the host-pathogen interaction in the complex environment of the host rather than the simplified system of the Petri dish5.

The rapid development, small size and transparency of zebrafish are tremendous advantages for the study of innate immune control of infection1-4. Here we demonstrate techniques for infecting zebrafish larvae using the fungal pathogen Candida albicans by microinjection, methodology recently used to implicate phagocyte NADPH oxidase activity in control of fungal dimorphism5.

Disseminated candidiasis caused by the pathogen Candida albicans is a clinically important problem in hospitalized individuals and is associated with a 30 ...

Tractable Mammalian Cell Infections with Protozoan-primed Bacteria

Many intracellular bacterial pathogens use freshwater protozoans as a natural reservoir for proliferation in the environment. Legionella pneumophila, the causative agent of Legionnaires' pneumonia, gains a pathogenic advantage over in vitro cultured bacteria when first harvested from protozoan cells prior to infection of mammalian macrophages. This suggests that important virulence factors may not be properly expressed in vitro. We have developed a tractable system for priming L. pneumophila through its natural protozoan host Acanthamoeba castellanii prior to mammalian cell infection. The contribution of any virulence factor can be examined by comparing intracellular growth of a mutant strain to wild-type bacteria after protozoan priming. GFP-expressing wild-type and mutant L. pneumophila strains are used to infect protozoan monolayers in a priming step and allowed to reach late stages of intracellular growth. Fluorescent bacteria are then harvested from these infected cells and normalized by spectrophotometry to generate comparable numbers of bacteria for a subsequent infection into mammalian macrophages. For quantification, live bacteria are monitored after infection using fluorescence microscopy, flow cytometry, and by colony plating. This technique highlights and relies on the contribution of host cell-dependent gene expression by mimicking the environment that would be encountered in a natural acquisition route. This approach can be modified to accommodate any bacterium that uses an intermediary host as a means for gaining a pathogenic advantage.

This technique provides a method to harvest, normalize and quantify intracellular growth of bacterial pathogens that are pre-cultivated in natural protozoan host cells prior to infections of mammalian cells. This method can be modified to accommodate a wide variety of host cells for the priming stage as well as target cell types.

Many intracellular bacterial pathogens use freshwater protozoans as a natural reservoir for proliferation in the environment. Legionella pneumophila, the ...

4D Multimodality Imaging of Citrobacter rodentium Infections in Mice

This protocol outlines the steps required to longitudinally monitor a bioluminescent bacterial infection using composite 3D diffuse light imaging tomography with integrated &mu;CT (DLIT-&mu;CT) and the subsequent use of this data to generate a four dimensional (4D) movie of the infection cycle. To develop the 4D infection movies and to validate the DLIT-&mu;CT imaging for bacterial infection studies using an IVIS Spectrum CT, we used infection with bioluminescent C. rodentium, which causes self-limiting colitis in mice. In this protocol, we outline the infection of mice with bioluminescent C. rodentium and non-invasive monitoring of colonization by daily DLIT-&mu;CT imaging and bacterial enumeration from feces for 8 days.
The use of the IVIS Spectrum CT facilitates seamless co-registration of optical and &mu;CT scans using a single imaging platform. The low dose &mu;CT modality enables the imaging of mice at multiple time points during infection, providing detailed anatomical localization of bioluminescent bacterial foci in 3D without causing artifacts from the cumulative radiation. Importantly, the 4D movies of infected mice provide a powerful analytical tool to monitor bacterial colonization dynamics in vivo.

4D Multimodality Imaging of Citrobacter rodentium Infections in Mice

Multi-modality imaging is a valuable approach for studying bacterial colonization in small animal models. This protocol outlines infection of mice with bioluminescent Citrobacter rodentium and the longitudinal monitoring of bacterial colonization using composite 3D diffuse light imaging tomography with &mu;CT imaging to create a 4D movie of C. rodentium infection.

This protocol outlines the steps required to  longitudinally monitor a bioluminescent bacterial infection using composite 3D diffuse light imaging ...

Zebrafish as a Model to Assess the Teratogenic Potential of Nitrite

High nitrate levels in the environment may&#160;result in congenital defects or miscarriages in humans. Presumably, this is due to the conversion of nitrate to nitrite by gut and salivary bacteria. However, in other mammalian studies, high nitrite levels do not cause birth defects, although they can lead to poor reproductive outcomes. Thus, the teratogenic potential of nitrite is not clear. It would be useful to have a vertebrate model system to easily assess teratogenic effects of nitrite or any other chemical of interest. Here, we demonstrate the utility of zebrafish (Danio rerio) to screen compounds for toxicity and embryonic defects. Zebrafish embryos are fertilized externally and have rapid development, making them a good model for teratogenic studies. We show that increasing the time of exposure to nitrite negatively affects survival. Increasing the concentration of nitrite also adversely affects survival, whereas nitrate does not. For embryos that survive nitrite exposure, various defects can occur, including pericardial and yolk sac&#160;edema, swim bladder noninflation, and craniofacial malformation. Our results indicate that the zebrafish is a convenient system for studying the teratogenic potential of nitrite. This approach can easily be adapted to test other chemicals for their effects on early vertebrate development.

Exposure to teratogens may cause birth defects. Zebrafish are useful for determining the teratogenic potential of chemicals. We demonstrate the utility of zebrafish by exposing embryos to various levels of nitrite and also at different times of exposure. We show that nitrite can be toxic and cause severe developmental defects.

High nitrate levels in the environment may&#160;result in congenital defects or miscarriages in humans. Presumably, this is due to the conversion of ...

Human Placental and Decidual Organ Cultures to Study Infections at the Maternal-fetal Interface

The placenta shows a large degree of interspecies anatomic variability. To best understand biology and pathophysiology of the human placenta, it is imperative to design experiments using human cells and tissues. An advantage of organ culture is maintenance of three-dimensional (3D) structural organization and extracellular matrix. The goal of the method described here is successful establishment of ex vivo human gestational tissue organ cultures and their healthy culture maintenance for 72-96 hr. The protocol details the immediate processing of research-consented, placental and decidual specimens fresh from the operating suite. These are abundant specimens that would otherwise be discarded. Detailed instructions on the sterile collection of these samples, including morphologic details on how to select appropriate tissues to establish 3D organ cultures, is provided. Placental villous and decidual tissues are microdissected into 2-3 mm3 pieces and placed separately on matrix-lined transwell filters and cultured for several days. Villous and decidual organ cultures are well suited for the study of human host-pathogen interaction. As compared to other model organisms, these human cultures are particularly advantageous to examine mechanism of infection for pathogens that demonstrate variable patterns of host specificity. As an example, we demonstrate infection of placental and decidual organ cultures with the clinically relevant, facultative intracellular bacterial pathogen Listeria monocytogenes.

A simple method to establish primary human placental (villous) and decidual organ cultures is described. Villous and decidual organ cultures are invaluable tools for studying pathogenesis at the human maternal-fetal interface. Infection with the facultative intracellular bacterium Listeria monocytogenes is demonstrated.

The placenta shows a large degree of interspecies anatomic variability. To best understand biology and pathophysiology of the human placenta, it is ...

Using Zebrafish Models of Human Influenza A Virus Infections to Screen Antiviral Drugs and Characterize Host Immune Cell Responses

Each year, seasonal influenza outbreaks profoundly affect societies worldwide. In spite of global efforts, influenza remains an intractable healthcare burden. The principle strategy to curtail infections is yearly vaccination. In individuals who have contracted influenza, antiviral drugs can mitigate symptoms. There is a clear and unmet need to develop alternative strategies to combat influenza. Several animal models have been created to model host-influenza interactions. Here, protocols for generating zebrafish models for systemic and localized human influenza A virus (IAV) infection are described. Using a systemic IAV infection model, small molecules with potential antiviral activity can be screened. As a proof-of-principle, a protocol that demonstrates the efficacy of the antiviral drug Zanamivir in IAV-infected zebrafish is described. It shows how disease phenotypes can be quantified to score the relative efficacy of potential antivirals in IAV-infected zebrafish. In recent years, there has been increased appreciation for the critical role neutrophils play in the human host response to influenza infection. The zebrafish has proven to be an indispensable model for the study of neutrophil biology, with direct impacts on human medicine. A protocol to generate a localized IAV infection in the Tg(mpx:mCherry) zebrafish line to study neutrophil biology in the context of a localized viral infection is described. Neutrophil recruitment to localized infection sites provides an additional quantifiable phenotype for assessing experimental manipulations that may have therapeutic applications. Both zebrafish protocols described faithfully recapitulate aspects of human IAV infection. The zebrafish model possesses numerous inherent advantages, including high fecundity, optical clarity, amenability to drug screening, and availability of transgenic lines, including those in which immune cells such as neutrophils are labeled with fluorescent proteins. The protocols detailed here exploit these advantages and have the potential to reveal critical insights into host-IAV interactions that may ultimately translate into the clinic.

Systemic and localized zebrafish infection models for human influenza A virus are demonstrated. Using a systemic infection model, zebrafish can be used to screen antiviral drugs. Using a localized infection model, zebrafish can be used to characterize host immune cell responses.

Each year, seasonal influenza outbreaks profoundly affect societies worldwide. In spite of global efforts, influenza remains an intractable healthcare ...

Arbovirus Infections As Screening Tools for the Identification of Viral Immunomodulators and Host Antiviral Factors

RNA interference- and genome editing-based screening platforms have been widely used to identify host cell factors that restrict virus replication. However, these screens are typically conducted in cells that are naturally permissive to the viral pathogen under study. Therefore, the robust replication of viruses in control conditions may limit the dynamic range of these screens. Furthermore, these screens may be unable to easily identify cellular defense pathways that restrict virus replication if the virus is well-adapted to the host and capable of countering antiviral defenses. In this article, we describe a new paradigm for exploring virus-host interactions through the use of screens that center on naturally abortive infections by arboviruses such as vesicular stomatitis virus (VSV). Despite the ability of VSV to replicate in a wide range of dipteran insect and mammalian hosts, VSV undergoes a post-entry, abortive infection in a variety of cell lines derived from lepidopteran insects, such as the gypsy moth (Lymantria dispar). However, these abortive VSV infections can be &#34;rescued&#34; when host cell antiviral defenses are compromised. We describe how VSV strains encoding convenient reporter genes and restrictive L. dispar cell lines can be paired to set-up screens to identify host factors involved in arbovirus restriction. Furthermore, we also show the utility of these screening tools in the identification of virally encoded factors that rescue VSV replication during coinfection or through ectopic expression, including those encoded by mammalian viruses. The natural restriction of VSV replication in L. dispar cells provides a high signal-to-noise ratio when screening for the conditions that promote VSV rescue, thus enabling the use of simplistic luminescence- and fluorescence-based assays to monitor the changes in VSV replication. These methodologies are valuable for understanding the interplay between host antiviral responses and viral immune evasion factors.

Here, we present the protocols to identify 1) virus-encoded immunomodulators that promote arbovirus replication and 2) eukaryotic host factors that restrict arbovirus replication. These fluorescence- and luminescence-based methods allow researchers to rapidly obtain quantitative readouts of arbovirus replication in simplistic assays with low signal-to-noise ratios.

RNA interference- and genome editing-based screening platforms have been widely used to identify host cell factors that restrict virus replication. ...

Confocal Imaging of Double-Stranded RNA and Pattern Recognition Receptors in Negative-Sense RNA Virus Infection

Double-stranded (ds) RNA is produced as a replicative intermediate during RNA virus infection. Recognition of dsRNA by host pattern recognition receptors (PRRs) such as the retinoic acid (RIG-I) like receptors (RLRs) RIG-I and melanoma differentiation-associated protein 5 (MDA-5) leads to the induction of the innate immune response. The formation and intracellular distribution of dsRNA in positive-sense RNA virus infection has been well characterized by microscopy. Many negative-sense RNA viruses, including some arenaviruses, trigger the innate immune response during infection. However, negative-sense RNA viruses were thought to produce low levels of dsRNA, which hinders the imaging study of PRR recognition of viral dsRNA. Additionally, infection experiments with highly pathogenic arenaviruses must be performed in high containment biosafety level facilities (BSL-4). The interaction between viral RNA and PRRs for highly pathogenic RNA virus is largely unknown due to the additional technical challenges that researchers need to face in the BSL-4 facilities. Recently, a monoclonal antibody (Mab) (clone 9D5) originally used for pan-enterovirus detection has been found to specifically detect dsRNA with a higher sensitivity than the traditional J2 or K1 anti-dsRNA antibodies. Herein, by utilizing the 9D5 antibody, we describe a confocal microscopy protocol that has been used successfully to visualize dsRNA, viral protein and PRR simultaneously in individual cells infected by arenavirus. The protocol is also suitable for imaging studies of dsRNA and PRR distribution in pathogenic arenavirus infected cells in BSL4 facilities.

Double-stranded RNA produced during RNA virus replication can be recognized by pattern recognition receptors to induce an innate immune response. For negative-sense RNA viruses, the interaction between the low-level dsRNA and PRRs remains unclear. We have developed a confocal microscopy method to visualize arenavirus dsRNA and PRR in individual cells.

Double-stranded (ds) RNA is produced as a replicative intermediate during RNA virus infection. Recognition of dsRNA by host pattern recognition receptors ...