The authors would like to thank lab members for zebrafish care and maintenance. This work was supported by National Institutes of Health, National Research Service Award A155397 to E. A. Harvie and NIH R01GM074827 to Anna Huttenlocher.

Adult and embryonic zebrafish were maintained in accordance with the University of Wisconsin-Madison Research Animal Resources Center.
1. Preparing Microinjection Needles
<ol>
	<li>Prepare thin wall glass capillary injection needles (1.0 OD/0.75 ID) using a micropipette puller device with the following settings: air pressure 200, heat 502, pull 90, velocity 80, time 70, air time at start of pull 5, air time at end of pull 5.</li>
	<li>Using fine tweezers, break off the tip of the pulled needle so that the tip opening has a diameter of approximately 10 &#181;m.</li>
</ol>
2. Preparing Larval Injection Dishes
<ol>
	<li>Rinse a gel comb with lane width of approximately 4-5 mm in sterile water and allow to dry.</li>
	<li>Prepare a 1.5-2% high melt agarose solution in E3 medium19 and microwave until the solution is clear. Once cooled, pour some of the agarose into a Petri dish (100 x 15 mm) and swirl, adding just enough agarose to completely cover the bottom of the dish.</li>
	<li>Once the agarose layer has solidified, place the rinsed and dried gel comb on top so that the non-combed end is just resting on the top of the Petri dish and the combed end is touching the agarose. Ensure that the comb is as horizontal as possible, creating a 30&#176; angle with respect to the bottom agarose layer.</li>
	<li>Pour an additional small amount of agarose at the interface between the comb and bottom agarose&#160;layer so that the fresh agarose layer covers the wells of the comb. Allow to cool completely before removing the comb. Using a pipet tip, remove any overhanging pieces of agarose from the wells.</li>
	<li>Pour E3 medium on top of the injection mold and store at 4 &#176;C.</li>
	<li>Before each use, replace with fresh medium and warm injection ramps at 28.5 &#176;C for at least an hour before injection.</li>
	<li>Immediately prior to injections, replace the E3 on the injection ramp with E3 medium containing 200 &#181;g/ml ethyl 3-aminobenzoate (tricaine).</li>
</ol>
3. Preparing S. iniae Inoculum
<ol>
	<li>Prepare and autoclave Todd Hewitt broth medium supplemented with 0.2% yeast extract and 2% proteose peptone (THY+P): 30 g/L Todd Hewitt, 2 g/L yeast extract, 20 g/L proteose peptone. For agar plates, add 14 g/L agar.</li>
	<li>Prepare bacterial cultures the night before infections by pipetting a 100 &#181;l aliquot of frozen bacterial stock into 10 ml of THY+P broth in a sealed 15 ml tube. Incubate overnight without agitation at 37 &#176;C. After 14-16 hr of growth, use the overnight culture either to make freezer stocks or prepare for use in injections.</li>
	<li>Make freezer stocks by placing 1 ml of the overnight culture in 500 &#181;l of 80% glycerol in a 1.7 ml centrifuge tube. To avoid freeze-thaw cycles, make 100 &#181;l one-use aliquots from this mix and store at -80 &#176;C.</li>
	<li>For S. iniae used in infections, dilute the overnight culture 1:100 for a total volume of 10 ml culture by adding 0.1 ml of overnight culture to 9.9 ml of THY+P broth. Grow at 37 &#176;C without agitation for approximately 4-5 hr. Monitor the optical density (OD) at 600 nm using a Nanodrop spectrophotometer and harvest the bacteria in mid-logarithmic phase when the OD600 nm reaches 0.250-0.500. An OD600 nm of 0.250 corresponds to approximately 108 colony forming units (CFU)/ml.</li>
	<li>Pellet 1 ml of the bacterial culture in a 1.7 ml centrifuge tube at 1,500 x g for 5 min. Resuspend in 1 ml of fresh PBS and repeat. Measure the OD600 nm of the bacteria in PBS, pellet, and resuspend in PBS to achieve the desired concentration.</li>
	<li>To aid in the visualization of microinjection, add phenol red to the bacterial suspensions prior to injection for a final concentration of 0.1%.</li>
	<li>For experiments involving the injection of heat-killed bacteria, heat the bacteria in PBS at 95 &#176;C for 30 min. Confirm that the heat-killing process reduced the number of viable bacteria to undetectable levels by plating an injection volume (approximately 1 nl) on solid THY+P agar plates and incubating overnight at 37 &#176;C.</li>
</ol>
4. Labeling S. iniae with a CellTracker Red Fluorescent Dye
<ol>
	<li>To label living bacterial cells, prepare a stock solution of a CellTracker fluorescent dye or equivalent. As the dye used comes in 20 x 50 &#181;g aliquots of powder and needs to be resuspended in DMSO, add 7.3 &#181;l DMSO to the tube to obtain a 10 mM stock concentration.</li>
	<li>Test a range of dye concentrations (e.g., 0.5-25 &#181;M) on the bacteria to determine the lowest optimal concentration that stains the cells. Rapidly dividing cells and longer experiments may require a higher concentration of dye.
	<ol>
		<li>Pellet 1.0 ml of bacterial culture in a 1.7 ml tube by centrifugation as described above (section 3). Resuspend the pellet in 1 ml fresh PBS and add the appropriate volume of dye to the bacterial culture.</li>
		<li>Incubate without agitation at 37 &#176;C for 30 min. Spin down the bacteria and resuspend in 1 ml of pre-warmed THY+P broth and incubate without agitation for an additional 30 min at 37 &#176;C.</li>
		<li>Spin down the bacteria and wash two times in PBS before measuring the OD600 nm and diluting the bacteria for microinjection as detailed above (section 3). We typically inject approximately 100 CFU in 1 nl injection volume for our studies of leukocyte recruitment and phagocytosis.</li>
	</ol>
	</li>
</ol>
5. Preparation of Zebrafish Larvae for Infections
<ol>
	<li>Set up breeding pairs the night before and collect embryos as described by Rosen et al.27 Incubate embryos in E3 medium at 28.5 &#176;C until ready to infect.</li>
	<li>For zebrafish that will be imaged, prevent the development of pigment (melanization) by adding N-Phenylthiourea (PTU) to the E3 medium at 24 hr post fertilization (hpf) for a final concentration of 0.2 nM.</li>
	<li>For infections involving embryos aged 2 dpf, dechorionate embryos manually with a pair of fine tweezers. Alternatively, dechorionate embryos by removing E3 and replacing with pronase (2 mg/ml) for about 5 min or until gentle pipetting breaks embryos out of the chorion. Most larvae should have hatched naturally by 3 dpf.</li>
	<li>Anesthetize zebrafish several minutes prior to infection by placing dechorionated larvae into E3 medium containing 200 &#181;g/ml tricaine.</li>
</ol>
6. Otic Vesicle Injection of S. iniae into Three Day Old Larvae
<ol>
	<li>Turn on the microinjector and set the time range to &#8220;millisecond&#8221;. Open the valve on the carbon dioxide tank to let gas into the line. The pressure of the microinjector should read approximately 20 PSI. Adjust the pressure in the microinjector unit by clockwise turning of the black knurled knob for increased pressure or counterclockwise turning for decreased pressure.</li>
	<li>Vortex the prepared S. iniae culture in phenol red and PBS and use a microloader tip to load 2-3 &#181;l of the culture into a pulled capillary injection needle.</li>
	<li>Mount the loaded needle on a micromanipulator connected to a magnetic stand and position it under a stereomicroscope so that the needle is at approximately a 45-65&#176; angle with respect to the base of the microscope.</li>
	<li>Press on the foot pedal of the microinjector to dispense a drop of the inoculum onto the tip of the needle. Measure the diameter of the drop using the scale bar in the ocular lens of the microscope.
	<ol>
		<li>Alternatively, estimate the diameter of the drop by injecting a volume into a drop of mineral oil on a glass microscope slide with a scale bar. The diameter of the drop should be approximately 0.10 mm, which is about a 1 nl volume.</li>
		<li>Adjust the drop size by adjusting the duration setting on the microinjector or by clipping off more of the needle tip with fine tweezers. Note that the injection time should be between 20 - 35 msec to avoid causing too much tissue damage.</li>
	</ol>
	</li>
	<li>Using a plastic transfer pipet, transfer 12 anesthetized larvae into each well of the injection mold.</li>
	<li>Use a glass rod, hair loop, or plastic tip to gently position the larvae so that the heads are pointed towards the back of the microscope and the yolk sacs are against the left side of the well. Point the left ear of the larva up towards the ceiling.</li>
	<li>Looking through the ocular lens of the stereomicroscope, use the knobs on the micromanipulator to line up the loaded needle with the otic vesicle so that both are in the same field of view. Pierce the outer epithelial layer of the otic vesicle with the needle tip so that the needle tip is just inside the vesicle.</li>
	<li>Press on the foot pedal to inject 1 nl of the desired dose of S. iniae. Be sure to use a low enough pressure so as not to rupture the cavity. If the injection is successful, the otic vesicle, but not the surrounding tissue, should fill with the phenol red inoculum (Figure 1 Ai). Immediately remove any mis-injected fish from the injection plate.</li>
	<li>Carefully retract the needle out of the larva and move the injection plate by hand so that the next larva is in view. Do this under 5X magnification. 
	Note: Retraction of the needle may result in the deposition of some bacteria outside of the otic vesicle, which may skew survival and leukocyte recruitment results. To avoid this, it is recommended to inject fluorescent dye-labeled bacteria and visually scan injected larvae under a fluorescent microscope to remove any larvae where this has occurred. A correctly injected larva is shown in (Figure 1 Bi).</li>
	<li>To ensure the injection volume/inoculum remains the same over the course of the experiment, inject a drop of the bacterial suspension into a 1.7 ml centrifuge tube containing 100 &#181;l of sterile PBS after every 48th embryo. Plate the 100 &#181;l on THY+P agar plates at 37 &#176;C overnight to determine the CFU in the injection volume.</li>
	<li>When the entire group of 12 larvae on the injection ramp has been injected, carefully use a plastic transfer pipette to remove the larvae from the wells and place them into a new Petri dish (35 x 10 mm2). Remove the tricaine solution and replace with approximately 2 ml of fresh E3 medium to allow the larvae to recover.</li>
	<li>Pipet injected larvae into single wells of a 96-well plate and incubate at 28.5 &#176;C. Monitor larvae over time for survival in these plates or remove later for imaging or CFU counts.</li>
</ol>
7. Sudan Black Staining of Neutrophils
Note: The following steps can be done at room temperature in a small Petri dish (35 x 10 mm2) on an orbital shaker unless otherwise stated. For each step, the amount of reagent used is approximately 2 ml per dish, using just enough liquid to completely cover the larvae.
<ol>
	<li>For fixation of infected larvae, use a glass Pasteur pipette to remove E3 medium and replace with an ice-cold 4% paraformaldehyde in PBS solution. Immediately place the fish at 4 &#176;C to euthanize. Fix overnight at 4 &#176;C.</li>
	<li>Wash 3 times in PBS for 5 min each.</li>
	<li>Stain with Sudan Black (0.18% stock diluted 1:5 in 70% ethanol, 0.1% phenol) for 30 min to 5 hr. Use a stereomicroscope to check if neutrophil granules have taken up the stain.</li>
	<li>Perform a series of 5 min washes starting with a single wash in 70% ethanol followed by a single wash of a 1:1 ratio of 70% ethanol and PBS, followed by a final wash in PBS.</li>
	<li>Rehydrate into PBS plus 0.1% tween (PBT).</li>
	<li>To clear pigment from the larvae, wash in 1% potassium hydroxide/1% hydrogen peroxide for about 6-10 min. Monitor the sample closely and after about 5 min, check the larvae under a stereomicroscope to see how much of the pigment has cleared. If the solution is left on too long, the yolk sacs will swell and burst.</li>
	<li>Wash 3 times for 5 min each in PBS.</li>
	<li>Store the samples in either PBS or PBT at 4 &#176;C for up to 1 week until ready to image and count.</li>
	<li>To image Sudan Black-stained larvae, transfer the fish into PBT and then into 80% glycerol and place onto a single cavity depression slide. Position fixed larvae under a stereomicroscope using a pipette tip. Image using a color digital camera on a stereomicroscope (Figure 1 Aii-iv).</li>
</ol>
8. Enumeration of Viable Bacteria from Infected Larvae
<ol>
	<li>Euthanize larvae at the desired times post infection in an ice-cold 200-300 mg/L solution of tricaine in E3 medium.</li>
	<li>Pipet euthanized larvae into 1.7 ml centrifuge tubes. Remove the tricaine solution and replace with 100 &#181;l of 0.2% Triton X-100 in PBS (PBSTx).</li>
	<li>Homogenize larvae by passing up and down 10 times through a 27 G needle.</li>
	<li>Prepare serial dilutions of homogenates in sterile PBS. For example, if the infectious dose is 100 CFU, pipet 100 &#181;l of the homogenate into 900 &#181;l sterile PBS. Using a new pipet tip, transfer 100 &#181;l of the diluted homogenate into 900 &#181;l sterile PBS.</li>
	<li>Plate 100 &#181;l of the dilutions on Columbia CNA agar for selective isolation of gram-positive bacteria, and incubate at 37 &#176;C for 48 hr. This medium will provide greater selection than the THY+P agar plates, which will support growth of both gram-negative and gram-positive bacteria.</li>
	<li>On plates with fewer than 500 individual colonies, count the number of colonies and multiply by the dilution factor to determine the number of CFU.</li>
</ol>
9. Fixation of Larvae for Imaging
<ol>
	<li>Fix larvae in an ice-cold 4% paraformaldehyde in PBS solution as described in section&#160;7.</li>
	<li>At room temperature, wash 3 times for 5 min each in PBS before imaging.</li>
</ol>
10.&#160;Preparation of Larvae for Live Imaging
<ol>
	<li>Prepare a&#160;1.5% low-melting-point agarose solution in E3 by heating in the microwave until the solution is clear.</li>
	<li>Place agarose solution in a 55 &#176;C water bath to let it cool but not harden.</li>
	<li>Add tricaine to the agarose to a final concentration of 0.016%.</li>
	<li>Using a plastic transfer pipette, place 4-5 anesthetized larvae in a glass bottom dish on the stage of a stereomicroscope.</li>
	<li>Remove the tricaine and agarose solution from the 55 &#176;C water bath and let it cool at room temperature for 1-2 min. While the agarose is cooling, remove as much liquid from the anesthetized larvae as possible.</li>
	<li>Pour the cooled agarose into the dish until about half of the surface is covered. Swirl the dish to spread the agarose. Note that if the agarose is too hot, it will kill the larvae. Also, if too much agarose is added to the dish, the objective lens of the microscope may hit the bottom of the dish when focusing through the agarose.</li>
	<li>Using a transfer pipette, pick up the larvae that have floated to the sides of the dish and pipet them back into the center.</li>
	<li>Under the stereomicroscope, gently position the larvae as desired with a hair loop, a long pipette tip or a glass rod. For imaging of the otic vesicle, position the larvae so that the left otic vesicle is flat against the bottom of the dish.</li>
	<li>Let the agarose cool for about 10 min before moving the dish. The larvae may shift positions if the agarose is not solid. Gently pipet some tricaine and E3 solution to the top of the agarose layer to keep it moist.</li>
</ol>
11. Confocal Imaging of Infection
<ol>
	<li>Place the glass bottom dish with larvae onto the stage of an inverted microscope with an FV-1000 laser scanning confocal system.</li>
	<li>Set the pinhole to 200-300 &#181;m and using a numeric aperture 0.75/20X objective lens, set z-stacks with 3-6 &#181;m slices.</li>
	<li>Use continuous line scanning to adjust the laser power and detector gain for each channel.</li>
	<li>Using sequential line scanning for each fluorescence channel (e.g., 488 and 543 nm) and differential interference contrast (DIC), conduct a time lapse movie of the left otic vesicle every 3 min for 2-6 hr to observe initial recruitment and phagocytosis by neutrophils and macrophages. For longer time courses, place a lid on the Petri dish to prevent evaporation and drying out of the agarose.</li>
	<li>Acquire still images of fixed (Figure 1B) or live (Figure 2) larvae at 20X or 40X magnification.</li>
</ol>
12. Photoconversion of Dendra2-labeled Leukocytes at the Otic Vesicle
Note: Dendra2 can be photoconverted from green to red fluorescence by focusing a 405 nm laser (50-70% laser power should be sufficient) on the region of interest (ROI) for 1 min. Below is the step-by-step protocol used for the FV-1000 laser scanning confocal system:
<ol>
	<li>Visualize the sample using a z-stack scan with the 488 nm and 543 nm lasers. Use continuous line scanning to adjust the laser power and detector gain. Check to make sure there is no accidentally photoconverted red fluorescence.</li>
	<li>On the &#8220;Image Acquisition Control&#8221; window, under &#8220;Stimulus Setting&#8221; select the &#8220;Use Scanner&#8221; tool and choose &#8220;main&#8221;. Select the 405 nm laser and set it at 70% power. Using the circle option, define the ROI&#160;in the otic vesicle.</li>
	<li>Under &#8220;Stimulus Start Setting&#8221; select &#8220;Activation in series&#8221; with a preactivation of 1 frame and an activation time of 60,000 msec. On the &#8220;Acquisition Setting&#8221; window under the &#8220;Time Scan heading&#8221;, choose 2 intervals of 00:01:00 (one for pre- and one for post- photoconversion). 
	Note: After the time lapse series scan is complete, the Dendra2 should have been photoconverted to its red fluorescent state.</li>
	<li>Scan the sample by a z-stack using the 488 nm and 543 nm lasers to visualize the photoconverted red fluorescence as well as any remaining green fluorescence (Figure 3).</li>
</ol>

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J., Mulvey, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+Zebrafish+to+Probe+the+Divergent+Virulence+Potentials+and+Toxin+Requirements+of+Extraintestinal+Pathogenic+Escherichia+coli.">Use of Zebrafish to Probe the Divergent Virulence Potentials and Toxin Requirements of Extraintestinal Pathogenic Escherichia coli.</a> PLoS Pathog. 5 (12), e1000697 (2009).</li><li>Harvie, E. A., Green, J. M., Neely, M. N., Huttenlocher, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Innate+Immune+Response+to+Streptococcus+iniae+Infection+in+Zebrafish+Larvae.">Innate Immune Response to Streptococcus iniae Infection in Zebrafish Larvae.</a> Infect. Immun. 81 (1), 110-121 (2013).</li><li>Haddon, C., Lewis, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+Ear+Development+in+the+Embryo+of+the+Zebrafish,+Danio+rerio.">Early Ear Development in the Embryo of the Zebrafish, Danio rerio.</a> J. Comp. Neurol. 365, 113-128 (1996).</li><li>Whitfield, T. T., Riley, B. B., Chiang, M. -. Y., Phillips, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+zebrafish+inner+ear.">Development of the zebrafish inner ear.</a> Dev. Dyn. 223 (4), 427-458 (2002).</li><li>Rosen, J. N., Sweeney, M. F., Mably, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+of+zebrafish+embryos+to+analyze+gene+function.">Microinjection of zebrafish embryos to analyze gene function.</a> J. Vis. Exp. (25), (2009).</li><li>Colucci-Guyon, E., Tinevez, J. Y., Renshaw, S. A., Herbomel, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strategies+of+professional+phagocytes+in+vivo:+unlike+macrophages,+neutrophils+engulf+only+surface-associated+microbes.">Strategies of professional phagocytes in vivo: unlike macrophages, neutrophils engulf only surface-associated microbes.</a> J. Cell Sci. 124 (18), 3053-3059 (2011).</li><li>Gurskaya, N. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineering+of+a+monomeric+green-to-red+photoactivatable+fluorescent+protein+induced+by+blue+light.">Engineering of a monomeric green-to-red photoactivatable fluorescent protein induced by blue light.</a> Nat. Biotechnol. 24 (4), 461-465 (2006).</li><li>Deng, Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Localized+bacterial+infection+induces+systemic+activation+of+neutrophils+through+Cxcr2+signaling+in+zebrafish.">Localized bacterial infection induces systemic activation of neutrophils through Cxcr2 signaling in zebrafish.</a> J. Leukoc. Biol. 93 (5), 761-769 (2013).</li></ol>

The authors declare that they have no competing financial interests.

The infection method used here is useful for the study of the host immune response to an initially localized infection in 2-3 dpf embryos and larvae. The focus of an inflammatory stimulus, such as infection, in a closed cavity such as the otic vesicle allows for the study of neutrophil and macrophage chemotaxis and phagocytosis. One caveat of injecting bacteria into the otic vesicle is that the ability of neutrophils to efficiently phagocytose bacteria in fluid-filled cavities may be dependent on the particular microbe. ...

Streptococcus iniae is a major aquatic pathogen that is capable of causing systemic disease in both fish and humans1. While S. iniae is responsible for large losses in the aquaculture industry, it is also a potential zoonotic pathogen, capable of causing disease in immunocompromised human hosts with clinical pathologies similar to those caused by other streptococcal human pathogens. Given its similarities with human pathogens, it is important to study S. iniae disease pathogenesis in the context of a natural host. An adult zebrafish model of S. iniae infection revealed robust infiltration of host leukocytes to the localized site of infection as well as a rapid time to host death, a time too short to involve the adaptive immune system7. In order to gain an in-depth look into the innate immune response to S. iniae infection in vivo, it is necessary to use a model that is more amenable to non-invasive live imaging.
The larval zebrafish has a number of advantages that make it an increasingly attractive vertebrate model for studying host-pathogen interactions. Zebrafish are relatively inexpensive and easy to use and maintain compared to mammalian models. Adaptive immunity is not functionally mature until 4-6 weeks post fertilization, but larvae have a highly conserved vertebrate innate immune system with complement, Toll-like receptors, cytokines, and neutrophils and macrophages with antimicrobial capabilities including phagocytosis and respiratory burst2-6,8-11. In addition, the genetic tractability and optical transparency of the embryonic and larval stages of development allow for the generation of stable transgenic lines with fluorescently labeled immune cells making it possible to examine host-pathogen interactions in real time in vivo. The generation of these transgenic lines using a photoconvertible protein such as Dendra2 allows for the tracking of individual host cell origin and fate over the course of infection12.
When developing a zebrafish larval infection model, the chosen site of microinjection will determine whether an infection is initially localized or systemic. Systemic blood infections into the caudal vein or Duct of Cuvier are most commonly used to study microbial pathogens in zebrafish and are useful for studying interactions between host and microbial cells, cytokine responses, and differences in virulence between pathogen strains. For slower growing microorganisms, early injection into the yolk sac of an embryo at the 16-1,000 cell stage can be used to generate a systemic infection13,14, with the optimal developmental stage for microinjection of a slow-growing microorganism found to be between the 16 to 128 cell stage15. However, yolk sac injections of many microbes at later stages of host development tend to be lethal to the host due to the nutrient-rich environment for the microbe and lack of infiltrating leukocytes16-18.
A localized infection usually results in directed migration of leukocytes towards the site of infection that can be easily quantified with non-invasive imaging. This type of infection can allow for dissection of the mechanisms that mediate leukocyte migration as well as investigation of different migratory and phagocytic capabilities of various leukocyte populations. Localized infections are also useful when examining differences in virulence between bacterial strains as well as studying microbe invasion mechanisms since physical host barriers must be crossed for a localized infection to become systemic. Zebrafish are typically raised at temperatures of 25-31 &#176;C19, but they can also be maintained at temperatures as high as 34-35 &#176;C for studies of the invasiveness of certain human pathogens with strict temperature requirements for virulence20,21.
Many different sites have been used to generate an initially localized bacterial infection including the hindbrain ventricle22, dorsal tail muscle18, pericardial cavity23, and otic vesicle (ear)5,16,24. However, it has been found that injection of bacteria into tail muscle can cause tissue damage and inflammation independent of the bacteria, which may skew results when investigating leukocyte response13. Although less damage is associated with injection into the hindbrain and although it is initially devoid of leukocytes in young embryos, the hindbrain ventricle steadily gains more immune cells over time as microglia take up residence. The hindbrain ventricle is also a more difficult location to image. The otic vesicle is a closed hollow cavity with no direct access to the vasculature25,26. It is normally devoid of leukocytes, but leukocytes can be recruited to the otic vesicle in response to inflammatory stimuli such as infection. It is also a preferred site of microinjection of bacteria in zebrafish aged 2-3 days post fertilization (dpf) because of the ease of imaging and the visualization of the injection. Therefore, we chose the otic vesicle as our site of localized bacterial infection.

<table><tbody><tr><td>1.7 ml Eppendorfs</td><td>MidSci</td><td>AVSS1700</td><td></td></tr><tr><td>14 ml Falcon tube</td><td>BD Falcon</td><td>352059</td><td></td></tr><tr><td>27 G x &amp;frac12; in. needle</td><td>BD Biosciences</td><td>305109</td><td></td></tr><tr><td>96-well Plate</td><td>Corning Incorporated</td><td>3596</td><td></td></tr><tr><td>Agar</td><td>BD Biosciences</td><td>214030</td><td></td></tr><tr><td>CellTracker Red</td><td>Molecular Probes, Invitrogen</td><td>C34552</td><td></td></tr><tr><td>CNA agar</td><td>Dot Scientific, Inc</td><td>7126A</td><td></td></tr><tr><td>Disposable transfer pipets</td><td>Fisher Scientific</td><td>13-711-7m</td><td></td></tr><tr><td>Dissecting scope</td><td>Nikon</td><td>SMZ745</td><td></td></tr><tr><td>DMSO</td><td>Sigma Aldrich</td><td>D2650</td><td></td></tr><tr><td>Ethanol 200 proof</td><td>MDS</td><td>2292</td><td></td></tr><tr><td>Fine tweezers</td><td>Fine Science Tools</td><td>11251-20</td><td></td></tr><tr><td>Gel comb</td><td>VWR</td><td>27372-482</td><td>4.2 mm width, 1.5 mm thick</td></tr><tr><td>Glass bottom dishes</td><td></td><td></td><td>Custom made by drilling a 16 - 18 mm hole in the center of a 35 mm tissue culture dish bottom and placing a 22 mm round #1 coverslip in the hole and sealing with a thin layer of Norland Optical Adhesive 68 cured by UV light.</td></tr><tr><td>Glycerol</td><td>Fisher Scientific</td><td>G33-4</td><td></td></tr><tr><td>High melt agarose</td><td>Denville Scientific, Inc.</td><td>CA3510-6</td><td></td></tr><tr><td>Hydrogen peroxide</td><td>Fisher Scientific</td><td>H325</td><td></td></tr><tr><td>Laser scanning confocal microscope</td><td>Olympus</td><td>with FV-1000 system</td><td></td></tr><tr><td>Low melt agarose</td><td>Fisher</td><td>BP165-25</td><td></td></tr><tr><td>Magnetic stand</td><td>Tritech (Narishige)</td><td>GJ-1</td><td></td></tr><tr><td>Microinjection system</td><td>Parker</td><td>Picospritzer III</td><td></td></tr><tr><td>Microloader pipet tips</td><td>Eppendorf</td><td>930001007</td><td></td></tr><tr><td>Micromanipulator</td><td>Tritech (Narishige)</td><td>M-152</td><td></td></tr><tr><td>Micropipette puller</td><td>Sutter Instrument Company</td><td>Flaming/Brown P-97</td><td></td></tr><tr><td>Nanodrop spectrophotmeter</td><td>Thermo Scientific</td><td>ND-1000</td><td></td></tr><tr><td>N-Phenylthiourea (PTU)</td><td>Sigma Aldrich</td><td>P7629</td><td></td></tr><tr><td>Paraformaldheyde</td><td>Electron Microscopy Sciences</td><td>15710</td><td></td></tr><tr><td>Petri dishes</td><td>Fisher Scientific</td><td>FB0875712</td><td>100 x 15 mm</td></tr><tr><td>Phenol</td><td>Sigma Aldrich</td><td>P-4557</td><td></td></tr><tr><td>Phenol Red</td><td>Ricca Chemoical Company</td><td>572516</td><td></td></tr><tr><td>Phosphate buffered saline</td><td>Fisher Scientific</td><td>BP665-1</td><td></td></tr><tr><td>Potassium hydroxide</td><td>Sigma Aldrich</td><td>P-6310</td><td></td></tr><tr><td>Pronase</td><td>Roche</td><td>165921</td><td></td></tr><tr><td>Protease peptone</td><td>Fluka Biochemika</td><td>29185</td><td></td></tr><tr><td>Small cell culture dish</td><td>Corning Incorporated</td><td>430165</td><td>35 x 10 mm</td></tr><tr><td>Sudan Black</td><td>Sigma Aldrich</td><td>S2380</td><td></td></tr><tr><td>Thin wall glass capillary injection needles</td><td>World Precision Instruments, Inc.</td><td>TW100-3</td><td></td></tr><tr><td>Todd Hewitt</td><td>Sigma Aldrich/Fluka Analytical</td><td>T1438</td><td></td></tr><tr><td>Tricaine (ethyl 3-aminobenzoate)</td><td>Argent Chemical Laboratory/Finquel</td><td>C-FINQ-UE-100G</td><td></td></tr><tr><td>Triton X-100</td><td>Fisher Scientific</td><td>BP151-500</td><td></td></tr><tr><td>Tween 20</td><td>Fisher Scientific</td><td>BP337-500</td><td></td></tr><tr><td>Yeast extract</td><td>Fluka Biochemika</td><td>92144</td><td></td></tr></tbody></table>

non-invasive imaging of the innate immune response in a zebrafish larval model of streptococcus iniae infection

The aquatic pathogen, Streptococcus iniae, is responsible for over 100 million dollars in annual losses for the aquaculture industry and is capable of causing systemic disease in both fish and humans. A better understanding of S. iniae disease pathogenesis requires an appropriate model system. The genetic tractability and the optical transparency of the early developmental stages of zebrafish allow for the generation and non-invasive imaging of transgenic lines with fluorescently tagged immune cells. The adaptive immune system is not fully functional until several weeks post fertilization, but zebrafish larvae have a conserved vertebrate innate immune system with both neutrophils and macrophages. Thus, the generation of a larval infection model allows the study of the specific contribution of innate immunity in controlling S. iniae infection.
The site of microinjection will determine whether an infection is systemic or initially localized. Here, we present our protocols for otic vesicle injection of zebrafish aged 2-3 days post fertilization as well as our techniques for fluorescent confocal imaging of infection. A localized infection site allows observation of initial microbe invasion, recruitment of host cells and dissemination of infection. Our findings using the zebrafish larval model of S. iniae infection indicate that zebrafish can be used to examine the differing contributions of host neutrophils and macrophages in localized bacterial infections. In addition, we describe how photolabeling of immune cells can be used to track individual host cell fate during the course of infection.

Microinjection of S. iniae into the otic vesicle (Figure 1 and Figure 2) results in an initially localized host response. When injected correctly, the bacteria should only be seen in the otic vesicle and not in the surrounding tissue or blood. This can be visualized during microinjection using phenol red dye (Figure 1A). Alternatively, if labeled bacteria are injected, a quick scan of infected larvae immediately post injection can confirm the bacteria are only i...

Watch this Scientific Journal Video about Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection at JoVE.com

Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection

Here, we present a protocol for the generation and imaging of a localized bacterial infection in the zebrafish otic vesicle.

Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection

The aquatic pathogen, Streptococcus iniae, is responsible for over 100 million dollars in annual losses for the aquaculture industry and is capable of ...

non-invasive-imaging-innate-immune-response-zebrafish-larval-model

Research

JoVE Journal

Immunology and Infection

10.7K Views.  University of Wisconsin-Madison. Here, we present a protocol for the generation and imaging of a localized bacterial infection in the zebrafish otic vesicle. 

Video: Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection

Live Imaging of Innate Immune and Preneoplastic Cell Interactions Using an Inducible Gal4/UAS Expression System in Larval Zebrafish Skin

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS expression system1 in zebrafish larvae, and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells. The KalTA4-ERT2/UAS system is both inducible and reversible which allows us to induce cell transformation with precise temporal/spatial resolution in vivo. This provides us with a unique opportunity to live image how individual preneoplastic cells interact with host tissues as soon as they emerge, then follow their progression as well as regression. Recent studies in zebrafish larvae have shown a trophic function of innate immunity in the earliest stages of tumorigenesis2,3. Our inducible system would allow us to live image the onset of cellular transformation and the subsequent host response, which may lead to important insights on the underlying mechanisms for the regulation of oncogenic trophic inflammatory responses. We also discuss how one might adapt our protocol to achieve temporal and spatial control of ectopic gene expression in any tissue of interest.

Studying the earliest events of preneoplastic cell progression and innate immune cell interaction is pivotal to understand and treat cancer. Here we describe a method to conditionally induce epithelial cell transformations and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells in zebrafish larvae.

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS ...

Developmental Biology

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 ...

Quantification of the Respiratory Burst Response as an Indicator of Innate Immune Health in Zebrafish

The phagocyte respiratory burst is part of the innate immune response to pathogen infection and involves the production of reactive oxygen species (ROS). ROS are toxic and function to kill phagocytized microorganisms. In vivo quantification of phagocyte-derived ROS provides information regarding an organism's ability to mount a robust innate immune response. Here we describe a protocol to quantify and compare ROS in whole zebrafish embryos upon chemical induction of the phagocyte respiratory burst. This method makes use of a non-fluorescent compound that becomes fluorescent upon oxidation by ROS. Individual zebrafish embryos are pipetted into the wells of a microplate and incubated in this fluorogenic substrate with or without a chemical inducer of the respiratory burst. Fluorescence in each well is quantified at desired time points using a microplate reader. Fluorescence readings are adjusted to eliminate background fluorescence and then compared using an unpaired t-test. This method allows for comparison of the respiratory burst potential of zebrafish embryos at different developmental stages and in response to experimental manipulations such as protein knockdown, overexpression, or treatment with pharmacological agents. This method can also be used to monitor the respiratory burst response in whole dissected kidneys or cell preparations from kidneys of adult zebrafish and some other fish species. We believe that the relative simplicity and adaptability of this protocol will complement existing protocols and will be of interest to researchers who seek to better understand the innate immune response.

The innate immune response protects organisms against pathogen infection. A critical component of the innate immune response, the phagocyte respiratory burst, generates reactive oxygen species that kill invading microorganisms. We describe a respiratory burst assay that quantifies reactive oxygen species produced when the innate immune response is chemically induced.

The phagocyte respiratory burst is part of the innate immune response to  pathogen infection and involves the production of reactive oxygen species (ROS). ...

Deciphering and Imaging Pathogenesis and Cording of Mycobacterium abscessus in Zebrafish Embryos

Zebrafish (Danio rerio) embryos are increasingly used as an infection model to study the function of the vertebrate innate immune system in host-pathogen interactions.&#160;The ease of obtaining large numbers of embryos, their accessibility due to external development, their optical transparency as well as the availability of a wide panoply of genetic/immunological tools and transgenic reporter line collections, contribute to the versatility of this model.&#160;In this respect, the present manuscript describes the use of zebrafish as an&#160;in vivo&#160;model system to investigate the chronology of Mycobacterium abscessus infection. This human pathogen can exist either as smooth (S) or rough (R) variants, depending on cell wall composition, and their respective virulence can be imaged and compared in zebrafish embryos and larvae. Micro-injection of either S or R fluorescent variants directly in the blood circulation via the caudal vein, leads to chronic or acute/lethal infections, respectively. This biological system allows high resolution visualization and analysis of the role of mycobacterial cording in promoting abscess formation. In addition, the use of fluorescent bacteria along with transgenic zebrafish lines harbouring fluorescent macrophages produces a unique opportunity for multi-color imaging of the host-pathogen interactions. This article describes detailed protocols for the preparation of homogenous M. abscessus inoculum and for intravenous injection of zebrafish embryos for subsequent fluorescence imaging of the interaction with macrophages. These techniques open the avenue to future investigations involving mutants defective in cord formation and are dedicated to understand how this impacts on M. abscessus pathogenicity in a whole vertebrate.

Deciphering and Imaging Pathogenesis and Cording of Mycobacterium abscessus in Zebrafish Embryos

Optically transparent zebrafish embryos are widely used to study and visualize in real time the interactions between pathogenic microorganisms and the innate immune cells. Micro-injection of Mycobacterium abscessus, combined with fluorescence imaging, is used to scrutinize essential pathogenic features such as cord formation in zebrafish embryos.

Zebrafish (Danio rerio) embryos are increasingly used as an infection model to study the function of the vertebrate innate immune system in host-pathogen ...

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 ...

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 ...

A Zebrafish Embryo Model for In Vivo Visualization and Intravital Analysis of Biomaterial-associated Staphylococcus aureus Infection

Biomaterial-associated infection (BAI) is a major cause of the failure of biomaterials/medical devices. Staphylococcus aureus is one of the major pathogens in BAI. Current experimental BAI mammalian animal models such as mouse models are costly and time-consuming, and therefore not suitable for high throughput analysis. Thus, novel animal models as complementary systems for investigating BAI in vivo are desired. In the present study, we aimed to develop a zebrafish embryo model for in vivo visualization and intravital analysis of bacterial infection in the presence of biomaterials based on fluorescence microscopy. In addition, the provoked macrophage response was studied. To this end, we used fluorescent protein-expressing S. aureus and transgenic zebrafish embryos expressing fluorescent proteins in their macrophages and developed a procedure to inject bacteria alone or together with microspheres into the muscle tissue of embryos. To monitor bacterial infection progression in live embryos over time, we devised a simple but reliable method of microscopic scoring of fluorescent bacteria. The results from microscopic scoring showed that all embryos with more than 20 colony-forming units (CFU) of bacteria yielded a positive fluorescent signal of bacteria. To study the potential effects of biomaterials on infection, we determined the CFU numbers of S. aureus with and without 10 &#181;m polystyrene microspheres (PS10) as model biomaterials in the embryos. Moreover, we used the ObjectJ project file &#34;Zebrafish-Immunotest&#34; operating in ImageJ to quantify the fluorescence intensity of S. aureus infection with and without PS10 over time. Results from both methods showed higher numbers of S. aureus in infected embryos with microspheres than in embryos without microspheres, indicating an increased infection susceptibility in the presence of the biomaterial. Thus, the present study shows the potential of the zebrafish embryo model to study BAI with the methods developed here.

A Zebrafish Embryo Model for In Vivo Visualization and Intravital Analysis of Biomaterial-associated Staphylococcus aureus Infection

The present study describes a zebrafish embryo model for in vivo visualization and intravital analysis of biomaterial-associated infection over time based on fluorescence microscopy. This model is a promising system complementing mammalian animal models such as mouse models for studying biomaterial-associated infections in vivo.

Biomaterial-associated infection (BAI) is a major cause of the failure of biomaterials/medical devices. Staphylococcus aureus is one of the major ...

Bioengineering

Photoconversion-Based Leukocyte Tracking in Bacterial-Infected Transgenic Zebrafish

Source: Harvie, E. A. et al., <a href="https://app.jove.com/v/52788">Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection.</a>&#160;J. Vis. Exp. (2015)
This video demonstrates the tracking of leukocyte recruitment using transgenic zebrafish larvae expressing photoconvertible green fluorescent protein. Pre-injecting red fluorescent dye-labeled bacteria into the otic vesicle and inducing photoconversion enables observation of leukocytes&#39; migration, phagocytosis, and dispersal throughout the body, providing valuable insights into immune response dynamics.

1. Preparing Microinjection Needles

	Prepare thin wall glass capillary injection needles (1.0 OD/0.75 ID) using a micropipette puller device with the ...

Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection

Summary

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Live Imaging of Innate Immune and Preneoplastic Cell Interactions Using an Inducible Gal4/UAS Expression System in Larval Zebrafish Skin

Infection of Zebrafish Embryos with Intracellular Bacterial Pathogens

Non-invasive Imaging of Disseminated Candidiasis in Zebrafish Larvae

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

Quantification of the Respiratory Burst Response as an Indicator of Innate Immune Health in Zebrafish

Deciphering and Imaging Pathogenesis and Cording of Mycobacterium abscessus in Zebrafish Embryos

Development of a Larval Zebrafish Infection Model for Clostridioides difficile

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

A Zebrafish Embryo Model for In Vivo Visualization and Intravital Analysis of Biomaterial-associated Staphylococcus aureus Infection

Photoconversion-Based Leukocyte Tracking in Bacterial-Infected Transgenic Zebrafish