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 border="0" cellpadding="0" cellspacing="0">
	<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 1/2 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&#8211;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 mm 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 mm 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

Non-invasive Imaging of Leukocyte Homing and Migration in vivo

Two-photon (2P) microscopy is a high resolution imaging technique that has been broadly adapted by biologists. The value of 2P microscopy is that it provides rich spatiotemporal information regarding cell behaviors within intact tissues and in live mice. Leukocyte recruitment plays a significant role in host defense against infection and when unchecked, can contribute to inflammatory and autoimmune diseases. Studying leukocyte recruitment in vivo is technically challenging since cells are moving rapidly within vessels located deep within light scattering tissues. To date, most intravital imaging studies require surgical preparation to expose the blood vessels and tissues. To avoid the tissue damage and inflammation induced by surgery itself, here, we describe a non-invasive single-cell imaging approach that can be used to study leukocyte trafficking in the mouse footpad and phalanges. We discuss the technical aspects of our 2P imaging preparation and walk the reader through a typical experiment from initial set up to execution and data collection.

Non-invasive Imaging of Leukocyte Homing and Migration in vivo

Here, we describe a non-invasive two-photon (2P) microscopy approach to study leukocyte homing in the mouse footpad. We discuss the technical aspects of our tissue imaging preparation and walk the reader through a typical experiment from initial set up to execution and data collection.

Two-photon (2P) microscopy is a high resolution imaging technique that has been broadly adapted by biologists.  The value of 2P microscopy is that it ...

Using Bioluminescent Imaging to Investigate Synergism Between Streptococcus pneumoniae and Influenza A Virus in Infant Mice

During the 1918 influenza virus pandemic, which killed approximately 50 million people worldwide, the majority of fatalities were not the result of infection with influenza virus alone. Instead, most individuals are thought to have succumbed to a secondary bacterial infection, predominately caused by the bacterium Streptococcus pneumoniae (the pneumococcus). The synergistic relationship between infections caused by influenza virus and the pneumococcus has subsequently been observed during the 1957 Asian influenza virus pandemic, as well as during seasonal outbreaks of the virus (reviewed in 1, 2). Here, we describe a protocol used to investigate the mechanism(s) that may be involved in increased morbidity as a result of concurrent influenza A virus and S. pneumoniae infection. We have developed an infant murine model to reliably and reproducibly demonstrate the effects of influenza virus infection of mice colonised with S. pneumoniae. Using this protocol, we have provided the first insight into the kinetics of pneumococcal transmission between co-housed, neonatal mice using in vivo imaging 3.

Using Bioluminescent Imaging to Investigate Synergism Between Streptococcus pneumoniae and Influenza A Virus in Infant Mice

A concurrent infection with influenza A virus is one of the factors implicated in the induction of invasive pneumococcal disease during asymptomatic Streptococcus pneumoniae carriage. Here we describe a mixed infection method using infant mice to investigate the synergism between these two respiratory pathogens.

During the 1918 influenza virus pandemic, which killed approximately 50 million people worldwide, the majority of fatalities were not the result of ...

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 ...

Non-invasive Optical Imaging of the Lymphatic Vasculature of a Mouse

The lymphatic vascular system is an important component of the circulatory system that maintains fluid homeostasis, provides immune surveillance, and mediates fat absorption in the gut. Yet despite its critical function, there is comparatively little understanding of how the lymphatic system adapts to serve these functions in health and disease1. Recently, we have demonstrated the ability to dynamically image lymphatic architecture and lymph "pumping" action in normal human subjects as well as in persons suffering lymphatic dysfunction using trace administration of a near-infrared fluorescent (NIRF) dye and a custom, Gen III-intensified imaging system2-4. NIRF imaging showed dramatic changes in lymphatic architecture and function with human disease. It remains unclear how these changes occur and new animal models are being developed to elucidate their genetic and molecular basis. In this protocol, we present NIRF lymphatic, small animal imaging5,6 using indocyanine green (ICG), a dye that has been used for 50 years in humans7, and a NIRF dye-labeled cyclic albumin binding domain (cABD-IRDye800) peptide that preferentially binds mouse and human albumin8. Approximately 5.5 times brighter than ICG, cABD-IRDye800 has a similar lymphatic clearance profile and can be injected in smaller doses than ICG to achieve sufficient NIRF signals for imaging8. Because both cABD-IRDye800 and ICG bind to albumin in the interstitial space8, they both may depict active protein transport into and within the lymphatics. Intradermal (ID) injections (5-50 &mu;l) of ICG (645 &mu;M) or cABD-IRDye800 (200 &mu;M) in saline are administered to the dorsal aspect of each hind paw and/or the left and right side of the base of the tail of an isoflurane-anesthetized mouse. The resulting dye concentration in the animal is 83-1,250 &mu;g/kg for ICG or 113-1,700 &mu;g/kg for cABD-IRDye800. Immediately following injections, functional lymphatic imaging is conducted for up to 1 hr using a customized, small animal NIRF imaging system. Whole animal spatial resolution can depict fluorescent lymphatic vessels of 100 microns or less, and images of structures up to 3 cm in depth can be acquired9. Images are acquired using V++ software and analyzed using ImageJ or MATLAB software. During analysis, consecutive regions of interest (ROIs) encompassing the entire vessel diameter are drawn along a given lymph vessel. The dimensions for each ROI are kept constant for a given vessel and NIRF intensity is measured for each ROI to quantitatively assess "packets" of lymph moving through vessels.

Recently developed imaging techniques using near-infrared fluorescence (NIRF) may help elucidate the role the lymphatic system plays in cancer metastasis, immune response, wound repair, and other lymphatic-associated diseases.

The lymphatic  vascular system is an important component of the circulatory system that  maintains fluid homeostasis, provides immune surveillance, and ...

Quantitative Measurement of the Immune Response and Sleep in Drosophila

A complex interaction between the immune response and host behavior has been described in a wide range of species. Excess sleep, in particular, is known to occur as a response to infection in mammals 1 and has also recently been described in Drosophila melanogaster2. It is generally accepted that sleep is beneficial to the host during an infection and that it is important for the maintenance of a robust immune system3,4. However, experimental evidence that supports this hypothesis is limited4, and the function of excess sleep during an immune response remains unclear. We have used a multidisciplinary approach to address this complex problem, and have conducted studies in the simple genetic model system, the fruitfly Drosophila melanogaster. We use a standard assay for measuring locomotor behavior and sleep in flies, and demonstrate how this assay is used to measure behavior in flies infected with a pathogenic strain of bacteria. This assay is also useful for monitoring the duration of survival in individual flies during an infection. Additional measures of immune function include the ability of flies to clear an infection and the activation of NF&kappa;B, a key transcription factor that is central to the innate immune response in Drosophila. Both survival outcome and bacterial clearance during infection together are indicators of resistance and tolerance to infection. Resistance refers to the ability of flies to clear an infection, while tolerance is defined as the ability of the host to limit damage from an infection and thereby survive despite high levels of pathogen within the system5. Real-time monitoring of NF&kappa;B activity during infection provides insight into a molecular mechanism of survival during infection. The use of Drosophila in these straightforward assays facilitates the genetic and molecular analyses of sleep and the immune response and how these two complex systems are reciprocally influenced.

Quantitative Measurement of the Immune Response and Sleep in Drosophila

To understand a link between the immune response and behavior, we describe a method to measure locomotor behavior in Drosophila during bacterial infection as well as the ability of flies to mount an immune response by monitoring survival, bacterial load, and real-time activity of a key regulator of innate immunity, NF&kappa;B.

A complex interaction between the immune response and  host behavior has been described in a wide range of species. Excess sleep, in  particular, is known ...

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). ...

Dissecting Innate Immune Signaling in Viral Evasion of Cytokine Production

In response to a viral infection, the host innate immune response is activated to up-regulate gene expression and production of antiviral cytokines. Conversely, viruses have evolved intricate strategies to evade and exploit host immune signaling for survival and propagation. Viral immune evasion, entailing host defense and viral evasion, provides one of the most fascinating and dynamic interfaces to discern the host-virus interaction. These studies advance our understanding in innate immune regulation and pave our way to develop novel antiviral therapies.
Murine &#947;HV68 is a natural pathogen of murine rodents. &#947;HV68 infection of mice provides a tractable small animal model to examine the antiviral response to human KSHV and EBV of which perturbation of in vivo virus-host interactions is not applicable. Here we describe a protocol to determine the antiviral cytokine production. This protocol can be adapted to other viruses and signaling pathways.
Recently, we have discovered that &#947;HV68 hijacks MAVS and IKK&#946;, key innate immune signaling components downstream of the cytosolic RIG-I and MDA5, to abrogate NF&#922;B activation and antiviral cytokine production. Specifically, &#947;HV68 infection activates IKK&#946; and that activated IKK&#946; phosphorylates RelA to accelerate RelA degradation. As such, &#947;HV68 efficiently uncouples NF&#922;B activation from its upstream activated IKK&#946;, negating antiviral cytokine gene expression. This study elucidates an intricate strategy whereby the upstream innate immune activation is intercepted by a viral pathogen to nullify the immediate downstream transcriptional activation and evade antiviral cytokine production.

We describe a protocol to measure the antiviral cytokine production in mice infected with a model herpesvirus, murine gamma herpesvirus 68 (&#947;HV68) that is closely-related to human Kaposi&#8217;s sarcoma-associated herpesvirus (KSHV) and Epstein-Barr virus (EBV). Utilizing genetically modified mouse strains and mouse embryonic fibroblasts (MEFs), we assessed the antiviral cytokine production both in vivo and ex vivo. &#8220;Reconstituting&#8221; the expression of innate immune components in knockout embryonic fibroblasts by lentiviral transduction, we further pinpoint specific innate immune molecules and dissect the key signaling events that differentially regulate the antiviral cytokine production.

In response to a viral infection, the host innate immune response is activated to up-regulate gene expression and production of antiviral cytokines. ...

Using RNA-interference to Investigate the Innate Immune Response in Mouse Macrophages

Macrophages are key phagocytic innate immune cells. When macrophages encounter a pathogen, they produce antimicrobial proteins and compounds to kill the pathogen, produce various cytokines and chemokines to recruit and stimulate other immune cells, and present antigens to stimulate the adaptive immune response. Thus, being able to efficiently manipulate macrophages with techniques such as RNA-interference (RNAi) is critical to our ability to investigate this important innate immune cell. However, macrophages can be technically challenging to transfect and can exhibit inefficient RNAi-induced gene knockdown. In this protocol, we describe methods to efficiently transfect two mouse macrophage cell lines (RAW264.7 and J774A.1) with siRNA using the Amaxa Nucleofector 96-well Shuttle System and describe procedures to maximize the effect of siRNA on gene knockdown. Moreover, the described methods are adapted to work in 96-well format, allowing for medium and high-throughput studies. To demonstrate the utility of this approach, we describe experiments that utilize RNAi to inhibit genes that regulate lipopolysaccharide (LPS)-induced cytokine production.

In this protocol, we describe methods to efficiently transfect murine macrophage cell lines with siRNAs using the Amaxa Nucleofector 96-well Shuttle System, stimulate the macrophages with lipopolysaccharide, and monitor the effects on inflammatory cytokine production.

Macrophages are key phagocytic innate immune cells.  When macrophages encounter a pathogen, they produce antimicrobial proteins and compounds to kill the ...

Non-Invasive Model of Neuropathogenic Escherichia coli Infection in the Neonatal Rat

Investigation of the interactions between animal host and bacterial pathogen is only meaningful if the infection model employed replicates the principal features of the natural infection. This protocol describes procedures for the establishment and evaluation of systemic infection due to neuropathogenic Escherichia coli K1 in the neonatal rat. Colonization of the gastrointestinal tract leads to dissemination of the pathogen along the gut-lymph-blood-brain course of infection and the model displays strong age dependency. A strain of E. coli O18:K1 with enhanced virulence for the neonatal rat produces exceptionally high rates of colonization, translocation to the blood compartment and invasion of the meninges following transit through the choroid plexus. As in the human host, penetration of the central nervous system is accompanied by local inflammation and an invariably lethal outcome. The model is of proven utility for studies of the mechanism of pathogenesis, for evaluation of therapeutic interventions and for assessment of bacterial virulence.

Non-Invasive Model of Neuropathogenic Escherichia coli Infection in the Neonatal Rat

Here, a procedure is described for the establishment of systemic infection in the neonatal rat with cultures of Escherichia coli K1. This non-invasive procedure permits colonization of the gastrointestinal tract, translocation of the pathogen to the systemic circulation, and invasion of the central nervous system at the choroid plexus.

Investigation of the interactions between animal host and bacterial pathogen is only meaningful if the infection model employed replicates the principal ...

In Vitro Analysis of Myd88-mediated Cellular Immune Response to West Nile Virus Mutant Strain Infection

An attenuated West Nile virus (WNV), a nonstructural (NS) 4B-P38G mutant, induced higher innate cytokine and T cell responses than the wild-type WNV in mice. Recently, myeloid differentiation factor 88 (MyD88) signaling was shown to be important for initial T cell priming and memory T cell development during WNV NS4B-P38G mutant infection. In this study, two flow cytometry-based methods &#8211; an in vitro T cell priming assay and an intracellular cytokine staining (ICS) &#8211; were utilized to assess dendritic cells (DCs) and T cell functions. In the T cell priming assay, cell proliferation was analyzed by flow cytometry following co-culture of DCs from both groups of mice with carboxyfluorescein succinimidyl ester (CFSE) - labeled CD4+ T cells of OTII transgenic mice. This approach provided an accurate determination of the percentage of proliferating CD4+ T cells with significantly improved overall sensitivity than the traditional assays with radioactive reagents. A microcentrifuge tube system was used in both cell culture and cytokine staining procedures of the ICS protocol. Compared to the traditional tissue culture plate-based system, this modified procedure was easier to perform at biosafety level (BL) 3 facilities. Moreover, WNV- infected cells were treated with paraformaldehyde in both assays, which enabled further analysis outside BL3 facilities. Overall, these in vitro immunological assays can be used to efficiently assess cell-mediated immune responses during WNV infection.

In Vitro Analysis of Myd88-mediated Cellular Immune Response to West Nile Virus Mutant Strain Infection

Two flow cytometry-based methods &#8211; an in vitro T cell priming assay and intracellular cytokine staining were utilized to measure antigen presenting capacity of dendritic cells and antigen-specific T cell responses to a West Nile virus mutant infection in mice.

An attenuated West Nile virus (WNV), a nonstructural (NS) 4B-P38G mutant, induced higher innate cytokine and T cell responses than the wild-type WNV in ...