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><tbody><tr><td>&lt;em&gt;Paramecium caudatum&lt;/em&gt;, live</td><td>Carolina</td><td>131554</td><td>no not store growing cultures below room temperature</td></tr><tr><td>0.4% Trypan Blue Solution&amp;nbsp;</td><td>Sigma</td><td>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><td>Dimethyl sulfoxide (DMSO)</td><td>Sigma</td><td>276855-100ML</td><td>store in a solvent safety cabinet</td></tr><tr><td>&lt;em&gt;Escherichia coli&lt;/em&gt;, MG1655</td><td>ATCC</td><td>ATCC 700926</td><td>can be replaced by any other non-pathogenic &lt;em&gt;E. coli&lt;/em&gt; strain</td></tr><tr><td>FM 4-64FX stain</td><td>Thermo Fisher</td><td>F34653</td><td>aliquot and store frozen</td></tr><tr><td>Formaldehyde</td><td>Sigma</td><td>F8775-4X25ML</td><td></td></tr><tr><td>LB Broth</td><td>Sigma</td><td>L3397-1KG</td><td></td></tr><tr><td>Phosphate buffered saline tablets</td><td>Thermo Fisher</td><td>18912014</td><td></td></tr><tr><td>Tetracycline</td><td>Sigma</td><td>87128-25G</td><td>toxic, irritant</td></tr><tr><td>Tricaine (Ethyl 3-aminobenzoate methanesulfonate)</td><td>Sigma</td><td>E10521-10G</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>X100-100ML</td><td></td></tr><tr><td>Trypan Blue Solution, 0.4%</td><td>Sigma</td><td>93595-50ML</td><td></td></tr><tr><td>UltraPure Low Melting Point Agarose</td><td>Thermo Fisher</td><td>16520050</td><td></td></tr><tr><td>hemocytometer or cell counter</td><td>any</td><td></td><td></td></tr><tr><td>stereomicroscope</td><td>any</td><td></td><td></td></tr><tr><td>table-top centrifuge</td><td></td><td></td><td></td></tr><tr><td>microwave</td><td></td><td></td><td></td></tr><tr><td>rotator wheel</td><td></td><td></td><td></td></tr><tr><td>heated shaking incubator</td><td></td><td></td><td></td></tr><tr><td>aquatics facilities</td><td></td><td></td><td></td></tr><tr><td>breeding tanks</td><td></td><td></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

28.5K Views.  University of Texas Health Science Center at Houston. Deze methode kan helpen bij het beantwoorden van belangrijke vragen in de microbiologie veld over de dynamiek van kolonisatie en infectie door micro-organismen in het maag-darmkanaal. De belangrijkste voordelen van deze techniek zijn dat het meer representatief is voor door voedsel overgedragen infecties bij de mens en dat het potentiële weefselschade bij vissen vermindert in vergelijking met orale gauas. Visuele demonstratie van deze methode is van cruciaal belang omdat de wasstappen moeili...

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

Infection of Zebrafish Embryos with Intracellular Bacterial Pathogens

Zebrafish (Danio rerio) embryos are increasingly used as a model for studying the function of the vertebrate innate immune system in host-pathogen interactions 1. The major cell types of the innate immune system, macrophages and neutrophils, develop during the first days of embryogenesis prior to the maturation of lymphocytes that are required for adaptive immune responses. The ease of obtaining large numbers of embryos, their accessibility due to external development, the optical transparency of embryonic and larval stages, a wide range of genetic tools, extensive mutant resources and collections of transgenic reporter lines, all add to the versatility of the zebrafish model. Salmonella enterica serovar Typhimurium (S. typhimurium) and Mycobacterium marinum can reside intracellularly in macrophages and are frequently used to study host-pathogen interactions in zebrafish embryos. The infection processes of these two bacterial pathogens are interesting to compare because S. typhimurium infection is acute and lethal within one day, whereas M. marinum infection is chronic and can be imaged up to the larval stage 2, 3. The site of micro-injection of bacteria into the embryo (Figure 1) determines whether the infection will rapidly become systemic or will initially remain localized. A rapid systemic infection can be established by micro-injecting bacteria directly into the blood circulation via the caudal vein at the posterior blood island or via the Duct of Cuvier, a wide circulation channel on the yolk sac connecting the heart to the trunk vasculature. At 1 dpf, when embryos at this stage have phagocytically active macrophages but neutrophils have not yet matured, injecting into the blood island is preferred. For injections at 2-3 dpf, when embryos also have developed functional (myeloperoxidase-producing) neutrophils, the Duct of Cuvier is preferred as the injection site. To study directed migration of myeloid cells towards local infections, bacteria can be injected into the tail muscle, otic vesicle, or hindbrain ventricle 4-6. In addition, the notochord, a structure that appears to be normally inaccessible to myeloid cells, is highly susceptible to local infection 7. A useful alternative for high-throughput applications is the injection of bacteria into the yolk of embryos within the first hours after fertilization 8. Combining fluorescent bacteria and transgenic zebrafish lines with fluorescent macrophages or neutrophils creates ideal circumstances for multi-color imaging of host-pathogen interactions. This video article will describe detailed protocols for intravenous and local infection of zebrafish embryos with S. typhimurium or M. marinum bacteria and for subsequent fluorescence imaging of the interaction with cells of the innate immune system.

Transparent zebrafish embryos have proved useful model hosts to visualize and functionally study interactions between innate immune cells and intracellular bacterial pathogens, such as Salmonella typhimurium and Mycobacterium marinum. Micro-injection of bacteria and multi-color fluorescence imaging are essential techniques involved in the application of zebrafish embryo infection models.

Zebrafish (Danio rerio) embryos are increasingly used as a model for studying the function of the vertebrate innate immune system in host-pathogen ...

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 ...

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

Chagas disease is a parasitic infection caused by Trypanosoma cruzi, whose motility is not only important for localization, but also for cellular binding and invasion. Current animal models for the study of T. cruzi allow limited observation of parasites in vivo, representing a challenge for understanding parasite behavior during the initial stages of infection in humans. This protozoan has a flagellar stage in both vector and mammalian hosts, but there are no studies describing its motility in vivo.The objective of this project was to establish a live vertebrate zebrafish model to evaluate T. cruzi motility in the vascular system. Transparent zebrafish larvae were injected with fluorescently labeled trypomastigotes and observed using light sheet fluorescence microscopy (LSFM), a noninvasive method to visualize live organisms with high optical resolution. The parasites could be visualized for extended periods of time due to this technique&#39;s relatively low risk of photodamage compared to confocal or epifluorescence microscopy. T. cruzi parasites were observed traveling in the circulatory system of live zebrafish in different-sized blood vessels and the yolk. They could also be seen attached to the yolk sac wall and to the atrioventricular valve despite the strong forces associated with heart contractions. LSFM of T. cruzi-inoculated zebrafish larvae is a valuable method that can be used to visualize circulating parasites and evaluate their tropism, migration patterns, and motility in the dynamic environment of the cardiovascular system of a live animal.

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

In this protocol, fluorescently labeled T. cruzi&#160;were injected into transparent zebrafish larvae, and parasite motility was observed in vivo using light sheet fluorescence microscopy.

Chagas disease is a parasitic infection caused by Trypanosoma cruzi, whose motility is not only important for localization, but also for cellular binding ...

Developmental Biology

Modeling Mucosal Candidiasis in Larval Zebrafish by Swimbladder Injection

Early defense against mucosal pathogens consists of both an epithelial barrier and innate immune cells. The immunocompetency of both, and their intercommunication, are paramount for the protection against infections. The interactions of epithelial and innate immune cells with a pathogen are best investigated in vivo, where complex behavior unfolds over time and space. However, existing models do not allow for easy spatio-temporal imaging of the battle with pathogens at the mucosal level.
The model developed here creates a mucosal infection by direct injection of the fungal pathogen, Candida albicans, into the swimbladder of juvenile zebrafish. The resulting infection enables high-resolution imaging of epithelial and innate immune cell behavior throughout the development of mucosal disease. The versatility of this method allows for interrogation of the host to probe the detailed sequence of immune events leading to phagocyte recruitment and to examine the roles of particular cell types and molecular pathways in protection. In addition, the behavior of the pathogen as a function of immune attack can be imaged simultaneously by using fluorescent protein-expressing C. albicans. Increased spatial resolution of the host-pathogen interaction is also possible using the described rapid swimbladder dissection technique.
The mucosal infection model described here is straightforward and highly reproducible, making it a valuable tool for the study of mucosal candidiasis. This system may also be broadly translatable to other mucosal pathogens such as mycobacterial, bacterial or viral microbes that normally infect through epithelial surfaces.

In vivo spatio-temporal interactions of pathogen and immune defenses at the mucosal level are not easily imaged in existing vertebrate hosts. The method presented here describes a versatile platform to study mucosal candidiasis in live vertebrates using the swimbladder of the juvenile zebrafish as an infection site.

Early defense against mucosal pathogens consists of both an epithelial barrier and innate immune cells. The immunocompetency of both, and their ...

Development of a Larval Zebrafish Infection Model for Clostridioides difficile

Clostridioides difficile infection (CDI) is considered to be one of the most common healthcare-associated gastrointestinal infections in the United States. The innate immune response against C. difficile has been described, but the exact roles of neutrophils and macrophages in CDI are less understood. In the current study, Danio rerio (zebrafish) larvae are used to establish a C. difficile infection model for imaging the behavior and cooperation of these innate immune cells in vivo. To monitor C. difficile, a labeling protocol using a fluorescent dye has been established. A localized infection is achieved by microinjecting labeled C. difficile, which actively grows in the zebrafish intestinal tract and mimics the intestinal epithelial damage in CDI. However, this direct infection protocol is invasive and causes microscopic wounds, which can affect experimental results. Hence, a more noninvasive microgavage protocol is described here. The method involves delivery of C. difficile cells directly into the intestine of zebrafish larvae by intubation through the open mouth. This infection method closely mimics the natural infection route of C. difficile.

Development of a Larval Zebrafish Infection Model for Clostridioides difficile

Presented here is a safe and effective method to infect zebrafish larvae with fluorescently labeled anaerobic C. difficile by microinjection and noninvasive microgavage.

Clostridioides difficile infection (CDI) is considered to be one of the most common healthcare-associated gastrointestinal infections in the United ...

Optimizing GF Zebrafish Models: Generation, Maintenance, and Microbial Studies

Generation, Maintenance, and Identification of Germ-Free Zebrafish Models from Larvae to Juvenile Stages

Zebrafish serve as valuable models for research on growth, immunity, and gut microbiota due to their genomic similarities with mammals, transparent embryos developed in a relatively clean chorion environment, and extremely rapid development of larvae compared to rodent models. Germ-free (GF) zebrafish (Danio rerio) are crucial for evaluating pollutant toxicity and establishing human-like disease models related to microbial functions. In comparison to conventionally raised (CR) models (fish in common husbandry), GF zebrafish allow for more accurate manipulation of the host microbiota, aiding in determining the causal relationship between microorganisms and hosts. Consequently, they play a critical role in advancing our understanding of these relationships. However, GF zebrafish models are typically generated and researched during the early life stages (from embryos to larvae) due to limitations in immune function and nutrient absorption. This study optimizes the generation, maintenance, and identification of early GF zebrafish models without feeding and with long-term feeding using GF food (such as Artemia sp., brine shrimp). Throughout the process, daily sampling and culture were performed and identified through multiple detections, including plates and 16S rRNA sequencing. The aseptic rate, survival, and developmental indexes of GF zebrafish were recorded to ensure the quality and quantity of the generated models. Importantly, this study provides details on bacterial isolation and infection techniques for GF fish, enabling the efficient creation of GF fish models from larvae to juvenile stages with GF food support. By applying these procedures in biomedical research, scientists can better understand the relationships between intestinal bacterial functions and host health.

Author Spotlight: Establishing Germ-Free Zebrafish for Microbiota-Host Relationship Studies

This protocol outlines the primary steps for obtaining germ-free (GF) fish embryos and maintaining them from larvae until the juvenile stage, including sampling and detecting their sterile status. The use of GF models with infection is important for understanding the role of microbes in host health.

Zebrafish serve as valuable models for research on growth, immunity, and gut microbiota due to their genomic similarities with mammals, transparent ...

Infection of Zebrafish Larvae with Aspergillus Spores for Analysis of Host-Pathogen Interactions

Invasive aspergillosis (IA) is one of&#160;the most common fungal infections among immunocompromised individuals. Despite the availability of antifungal drugs, IA can cause &#62;50% mortality in infected immunocompromised patients. It is crucial to determine both host and pathogen factors that contribute to infection susceptibility and low survival rates in infected patients in order to develop novel therapeutics. Innate immune responses play a pivotal role in recognition and clearance of Aspergillus spores, though little is known about the exact cellular and molecular mechanisms. Reliable models are required to investigate detailed mechanistic interactions between the host and pathogen. The optical clarity and genetic tractability of zebrafish larvae make them an intriguing model&#160;to study host-pathogen interactions of multiple human bacterial and fungal infections in a live and intact host. This protocol describes a larval zebrafish Aspergillus infection model. First, Aspergillus spores are isolated and injected into the zebrafish hindbrain ventricle via microinjection. Then, chemical inhibitors such as immunosuppressive drugs are added directly to the larval water. Two methods to monitor the infection in injected larvae are described, including the 1) homogenization of larvae for colony forming unit (CFU) enumeration and 2) a repeated, daily live imaging setup. Overall, these techniques can be used to mechanistically analyze the progression of Aspergillus infection in vivo and can be applied to different host backgrounds and Aspergillus strains to interrogate host-pathogen interactions.

Infection of Zebrafish Larvae with Aspergillus Spores for Analysis of Host-Pathogen Interactions

This protocol describes an Aspergillus infection model in zebrafish larvae. Aspergillus spores are microinjected into the hindbrain of larvae, and chemical treatment is used to induce immunosuppression. Infection progression is monitored via a daily imaging setup to monitor fungal growth and immune responses as well as enumeration of live spores by colony forming unit plating.

Invasive aspergillosis (IA) is one of&#160;the most common fungal infections among immunocompromised individuals. Despite the availability of antifungal ...

High-fat Feeding Paradigm for Larval Zebrafish: Feeding, Live Imaging, and Quantification of Food Intake

Zebrafish are emerging as a model of dietary lipid processing and metabolic disease. This protocol describes how to feed larval zebrafish a lipid-rich meal, which consists of an emulsion of chicken egg yolk liposomes created by sonicating egg yolk in embryo media. Detailed instructions are provided to screen larvae for egg yolk consumption so that larvae that fail to feed will not confound experimental results. The chicken egg yolk liposomes can be spiked with fluorescent lipid analogs, including fatty acids and cholesterol, enabling both systemic and subcellular visualization of dietary lipid processing. Several methods are described to mount larvae that are conducive to short- and long-term live imaging with both upright and inverted objectives at high and low magnification. Additionally presented is an assay to quantify larval food intake by extracting the lipids of larvae fed fluorescent lipid analogs, spotting the lipids on a thin layer chromatography plate, and quantifying the fluorescence. Finally, critical aspects of the procedures, important controls, options for modifying the protocols to address specific experimental questions, and potential limitations are discussed. These techniques can be applied not only to focused, hypothesis driven inquiries, but also to a variety of screens and live imaging techniques to study dietary lipid metabolism and the control of food intake.

Zebrafish are emerging as a valuable model of dietary lipid processing and metabolic disease. Described are protocols of lipid-rich larval feeds, live imaging of dietary fluorescent lipid analogs, and quantification of food intake. These techniques can be applied to a variety of screening, imaging, and hypothesis driven inquiry techniques.

Zebrafish are emerging as a model of dietary lipid processing and metabolic disease. This protocol describes how to feed larval zebrafish a lipid-rich ...

Biology

Environmental Screening of Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa in Zebrafish Systems

Health monitoring systems are developed and used in zebrafish research facilities because pathogens of Danio rerio such as Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa have the potential to impair animal welfare and research. The fish are typically analyzed post mortem to detect microbes. The use of sentinels is a suggested way to improve the sensitivity of the surveillance and to reduce the number of animals to sample. The setting of a pre-filtration sentinel tank out of a recirculating system is described. The technique is developed to prevent water pollution and to represent the fish population by a careful selection of age, gender, and strains. In order to use the minimum number of animals, techniques to screen the environment are also detailed. Polymerase Chain Reaction (PCR) on surface sump swabs is used to significantly improve the detection of some prevalent and pathogenic mycobacterial species such as Mycobacterium fortuitum, Mycobacterium haemophilum, and Mycobacterium chelonae. Another environmental method consists of processing the sludge at the bottom of a holding tank or sump to look for P. tomentosa eggs. This is a cheap and fast technique that can be applied in quarantine where a breeding device is submerged into the holding tank of imported animals. Finally, PCR is applied to the sludge sample and A. hydrophila is detected at the sump&#39;s bottom and surface. Generally, these environmental screening techniques applied to these specific pathogens have led to an increased sensitivity compared to the testing of pre-filtration sentinels.

Environmental Screening of Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa in Zebrafish Systems

This protocol describes the use of sump swabs and sludge analysis of zebrafish systems, which leads to increased detection compared to the sole use of sentinels to detect pathogens such as Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa. A system to monitor P. tomentosa eggs in quarantine is also proposed.

Health monitoring systems are developed and used in zebrafish research facilities because pathogens of Danio rerio such as Aeromonas hydrophila, ...

High-throughput Measurement of Gut Transit Time Using Larval Zebrafish

Zebrafish are used as alternative model organisms for drug safety testing. The gastrointestinal (GI) tract of zebrafish has genetic, neuronal, and pharmacological similarities to that of the mammals. GI intolerance during clinical testing of drug candidates is common and may pose a serious threat to human health. Testing for GI toxicity in preclinical mammalian models can be expensive in terms of time, test compound, and labor. The high-throughput method presented here may be used to predict GI safety issues. Compared to mammalian models, this method allows for more expedient assessment of test compound effects on GI transit while using low quantities of compound. In this method, larval zebrafish (7 days post fertilization) are fed food containing a fluorescent label. After feeding, each larval fish is placed into a well of a 96-conical-bottom-well plate and dosed with test compound (dissolved in water) or the vehicle. As gut transit occurs, fecal matter accumulates on the bottom of the wells, and the rate at which this happens is monitored by measuring fluorescence from the bottom of the well repeatedly over time using a plate spectrophotometer. The fluorescence from larvae in a given treatment group are averaged and these values are graphed along with standard error, for each measurement time, yielding a curve representing average transit of food over time. Effects on gut transit time are identified by comparing the area under the curve for each treatment group to that of the vehicle-treated group. This method detected changes in zebrafish GI transit time induced by drugs with known clinical GI effects; it can be employed to interrogate dozens of treatments for GI effects per day. As such, safer compounds can be quickly prioritized for mammalian testing, which expedites discovery and proffers 3Rs advancement.

The goal of this protocol is to measure the transit time of fluorescently labeled food through the gut of larval zebrafish in a high throughput fashion.

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

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