We are grateful to Timo Fritsch for excellent animal care. We thank the members of the K&#246;ster and Steinert labs for support and helpful discussions. We thank Dr. Dandan Han for critical reading the manuscript. We gratefully acknowledge funding by the Federal State of Lower Saxony, Nieders&#228;chsisches Vorab (VWZN2889).

All animal work described here was performed in accordance with legal regulations (EU-Directive 2010/63, license AZ 325.1.53/56.1-TUBS and license AZ 33.9-42502-04-14/1418).
1. Preparation of Low Melting Agarose, Gel Plate, and Microinjection/Microgavage Needles
<ol>
	<li>Dissolve 0.08 g of low melting agarose (Table of Materials, agarose A2576) in 10 mL of 30% Danieau&#39;s medium (0.12 mM MgSO4, 0.18 mM Ca [NO3]2), 0.21 mM KCl, 1.5 mM HEPES (pH = 7.2), and 17.4 mM NaCl, stored at room temperature (RT) to obtain a 0.8% solution. 
	NOTE: Higher or lower concentrations of agarose can be used. However, the required time to solidify varies for different brands of agarose, even at the same concentration.</li>
	<li>Prepare microinjection and microgavage needles from glass capillaries (Table of Materials).
	<ol>
		<li>Use a micropipette puller with the following settings (note that units are specific to the puller used here; see Table of Materials): microinjection needles (air pressure = 500; heat = 400; pull = 125; velocity = 75; time = 150); and microgavage needles (air pressure = 500; heat = 400; pull = 100; velocity = 75; time = 150). Use a microloader tip to load 3 &#181;L of nuclease-free H2O into the pulled needle.</li>
		<li>Introduce the needle into the injector and fasten it properly. Adjust the needle to a suitable angle for injection. Set the injection pressure between 600&#8211;900 hPa for microinjection needle and 200&#8211;300 hPa for gavage needle.</li>
		<li>Place a drop of mineral oil onto the black circle of the calibration slide. Use fine forceps to clip the tip of the needle. Inject one drop into the mineral oil to measure the size of the droplet.</li>
		<li>For microinjection, adjust the injection time to obtain a droplet with a diameter of 0.10&#8211;0.12 mm, which equals a volume of 0.5&#8211;1.0 nL. For microgavage, obtain a droplet with a diameter of 0.18&#8211;0.20 mm, which equals a volume of 3&#8211;5 nL.</li>
	</ol>
	</li>
	<li>Prepare a 1.5 % agarose plate with agarose (Table of Materials, 8050) in a 10 cm Petri dish in 30% Danieau&#39;s medium using a plastic mold as the microgavage mold. Store at 4 &#176;C to prevent desiccation until needed. Warm to RT or 28 &#176;C prior to the experiment.</li>
</ol>
2. Preparation and Labeling of C. difficile and Spores with Fluorescent Dye
<ol>
	<li>Prepare a 1 mM stock solution of a fluorescent dye (Table of Materials). Because the dye is sold in 50 &#181;g aliquots of powder, add 69 &#181;L of DMSO to the vial to obtain a 1 mM stock concentration.</li>
	<li>Prepare a 100 &#181;M working solution of the fluorescent dye by adding 2 &#181;L of 1 mM stock solution to 18 &#181;L of DMSO in a centrifuge tube, and mix well.</li>
	<li>Culture C. difficile (R20291, a ribotype 027 strain) by inoculating 10 mL of BHIS liquid medium with two to three colonies from a plate in an anaerobic hood without shaking overnight. BHIS is BHI supplemented with 0.5% (w/v) yeast extract and 0.1% (w/v) L-cysteine. Dissolve 1 g of L-cysteine in 10 mL of ddH2O and sterilize by filtration, then add to the autoclaved medium to obtain a final concentration of 1 g/L. Plates are prepared by adding 15 g/L agar-agar to the medium before autoclaving. Selective culturing of C. difficile is done by using chromID C. difficile plates (Table of Materials).</li>
	<li>Staining C. difficile with the fluorescent dye
	<ol>
		<li>Harvest C. difficile at an OD600 of 1.0&#8211;1.2 and wash 1x with 1 mL of 1x PBS (5,000 x g for 3 min at RT). Resuspend in 1 mL of 1 x PBS.</li>
		<li>Add 3 &#181;L of working solution of the fluorescent dye into 1 mL of bacteria suspension. Incubate the sample for 15 min at RT in the dark. Wash the stained C. difficile once with 1 mL PBS to remove residual dye and resuspend in 1x PBS to an OD600 of 1.0 (1.0 OD600 is approximately equivalent to 108 cfu/mL).</li>
	</ol>
	</li>
</ol>
3. Injection of Stained C. difficile into Zebrafish Larvae
<ol>
	<li>Anesthetize 20&#8211;30 zebrafish larvae at 5 days post-fertilization (referred to here as 5 dpf) with 0.02%&#8211;0.04% tricaine (tricaine powder is dissolved in double-distilled water and adjusted to pH = 7 with 1 M Tris-HCL solution) in 30% Danieau&#39;s&#39;s medium ~10 min before injection. Transfer the anesthetized larvae to a fresh 10 cm Petri dish and remove any excess 30% Danieau&#39;s&#39;s medium.</li>
	<li>Place a drop of 0.8% low melting agarose onto the zebrafish larvae to cover. Gently adjust the larvae to a lateral position. Place the Petri dish on ice for 30&#8211;60 s to allow the low melting agarose to solidify. Add 30% Danieau&#39;s medium containing 0.02%&#8211;0.04% tricaine to cover the agarose.</li>
	<li>Prepare the injection solution. Add 1 &#181;L of 0.5% phenol red in PBS solution into 9 &#181;L of the dye-stained C. difficile inoculum to visualize the injection process.</li>
	<li>Load a calibrated microinjection needle with the injection solution using a microloader. Mount the loaded needle onto a micromanipulator and position it under a stereomicroscope.</li>
	<li>Adjust the injection pressure between 600&#8211;900 hPa. Set the injection time to 0.1&#8211;0.3 s to obtain 0.5&#8211;1.0 nL. Set the needle in the micromanipulator at a ~45&#176; angle pointing toward the embedded larvae.</li>
	<li>Place the needle tip above the gastrointestinal tract close to the urogenital pore. Pierce through agarose then the muscle with the needle tip, then insert it into the intestinal lumen and inject 0.5&#8211;1.0 nL of C. difficile. Use a fluorescence microscope to monitor the injected larvae and pick up the properly injected larvae for confocal imaging.</li>
</ol>
4. Generation of Gnotobiotic Zebrafish Larvae
<ol>
	<li>Use the well-established natural breeding method to generate gnotobiotic zebrafish embryos, including: in vitro fertilization, washing with antibiotic-containing medium (1 &#181;g/mL amphotericin B, 10 &#181;g/mL kanamycin, and 20 &#181;g/mL ampicillin), washing with 0.1% wt/vol polyvinyl pyrrolidone-iodine (PVP-I) solution, and incubation of the embryos in a cell culture hood12.</li>
	<li>Maintain all gnotobiotic zebrafish larvae under gnotobiotic conditions until 5 dpf or just before the gavage. After the gavage, zebrafish larvae will be transferred into a standard incubator but with sterile 30% Danieau&#39;s medium.</li>
</ol>
5. Gavage of Zebrafish Larvae
<ol>
	<li>Calibrate the microgavage needle as described in step 1.2.</li>
	<li>Measure the diameter of the tip of the needle by placing the needle on a calibration slide with one drop of mineral oil. Ensure that the tip is 30&#8211;40 &#181;m in diameter, blunt, and smooth. Discard the sharp or rough needles. 
	NOTE: Sharp edges of needles can be blunted by quick flaming.</li>
	<li>Prepare the gavage solution as described in step 3.3.</li>
	<li>Load and mount the needle onto a micromanipulator as described in step 3.4. Adjust the micromanipulator to position the needle at a 45&#176; angle.</li>
	<li>Anesthetize the zebrafish larvae referred to in step 3.1. When the larvae stop moving, transfer them to the groove of a microgavage mold using a Pasteur pipette.</li>
	<li>Place a drop of 0.8% low melting agarose onto the zebrafish larvae to cover. Gently adjust the larvae with heads facing upright at 45&#176; angles in the groove and tails against the wall of the groove. Ensure that the angles of the heads are approximately the same so that they are aligned with the angle of the gavage needles. Place the microgavage mold on ice for 30&#8211;60 s, allowing the low melting agarose to solidify in order to stabilize the positions of the larvae.</li>
	<li>Adjust the injection pressure between 200&#8211;300 hPa. Set the injection time to 0.1&#8211;0.3 s to obtain an injection volume of 3&#8211;5 nL of C. difficile.</li>
	<li>Gently operate the needle through the agarose then into the mouth of zebrafish larvae, through the esophagus. Once the tip of the needle is inside the anterior intestinal bulb, press the injection pedal to release the bacteria. Fill the lumen of the intestine with the delivered volume. Do not let it overflow from the esophagus or cloaca. Gently withdraw the needle from the mouth of the zebrafish.</li>
	<li>Following gavage, rescue the infected zebrafish larvae from the agarose with a flexible microloader tip by first cutting the agarose away then by lifting the larvae. Transfer these larvae into sterile 30% Danieau&#39;s medium. Rinse the larvae in sterile medium twice. Transfer the larvae to a fresh 10 cm Petri dish. The larvae will be maintained for up to 11 dpf.</li>
</ol>
6. Confocal Microscopy Analysis of Injected Zebrafish Larvae
<ol>
	<li>Anesthetize zebrafish larvae referred to step 3.1. Make a hole in the bottom of a 35 mm Petri dish with a glass slide attached to the hole, referred as the imaging chamber. Transfer embryos to the bottom of the imaging chamber with an adequate amount of 30% Danieau&#39;s medium.</li>
	<li>Add 200&#8211;300 &#956;L of 1% low melting agarose to cover the anesthetized larvae. Since an inverted confocal microscope is used, place the infected region of the larvae against the glass slide as closely as possible.</li>
	<li>Let the agarose solidify on ice for 30&#8211;60 s. Submerge the agarose with 30% Danieau&#39;s containing 0.02%&#8211;0.04% tricaine.</li>
	<li>Image the larvae with a confocal laser scanning microscope (Table of Materials).</li>
</ol>
7. Dissection of Larval Zebrafish Intestine to Recover Viable C. difficile
<ol>
	<li>Isolate gastrointestinal tracts from larvae to analyze bacterial load. Start by euthanizing zebrafish larvae with 0.4 % tricaine.</li>
	<li>Rinse the zebrafish briefly with sterile 1x PBS and transfer them to a fresh agarose plate.</li>
	<li>Dissection of zebrafish
	<ol>
		<li>Insert a needle into the dorsal trunk of zebrafish larvae close to the head to immobilize the zebrafish. Remove the head behind the gills with a lancet.</li>
		<li>Insert the second needle into the middle of the dorsal trunk. Insert the third needle into the abdomen of the zebrafish and pull the intestine out of the body cavity. 
		NOTE: Extreme care is needed to isolate the intact intestine. If it is difficult to do so, perform additional pulls to separate the rest of the intestine from the remaining internal organs.</li>
		<li>Use a microinjection needle to transfer 10&#8211;15 intestines into a 1.5 mL tube containing 200 &#181;L sterile of 1x PBS.</li>
		<li>Homogenize the intestines with a pestle to disrupt the tissue and prepare homogenates. Ensure the pestle reaches the bottom of the tube to disrupt all intestines completely.</li>
	</ol>
	</li>
	<li>Incubate the homogenates in C. difficile rearing medium containing D-cycloserine and cefoxitin, with or without taurocholate (TCA, a germinant of C. difficile spores) in an anaerobic chamber.</li>
	<li>Incubate the plate anaerobically for 48 h at 37 &#176;C.</li>
	<li>Use bacterial culture for 16S rRNA-PCR.
	<ol>
		<li>Resuspend a colony in 50 &#181;L of H2O and boil it at 95 &#176;C for 15 min. Pellet the lysed debris by centrifugation (14,000 rpm for 2 min at RT) and use 2 &#181;L of the supernatant as a template in 25 &#181;L of PCR-reaction using C. difficile-specific primers (Cdiff16Sfw: 5&#39; GTG AGC CAG TAC AGG 3&#39;; Cdiff16Srev: 5&#39; TTA AGG AGA TGT CAT TGG 3&#39;).</li>
		<li>When bacteria from liquid culture are used, harvest 1 mL of culture and wash once with 1 mL of PBS (14,000 rpm for 2 min at RT). Resuspend the pellet in 100 &#181;L of H2Odd and treat as done above. To further characterize bacterial colonies, streak on a BHIS- or chromID-plate (Table of Materials).</li>
	</ol>
	</li>
</ol>

&#160; &#160;

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The authors have nothing to disclose.

The presented methods modify and extend an existing approach to infect zebrafish larvae by performing both injection and microgavage10,14. It also demonstrates an approach to study anaerobic pathogens with zebrafish larvae22. In addition, the protocol facilitates the analysis of innate immune cell responses in vivo upon CDI and upon colonization of C. difficile in zebrafish. The method is reproducible and easy to conduct in a rout...

C. difficile is a gram-positive, spore-forming, anaerobic, and toxin-producing bacillus that is the leading cause of severe infections in the gastrointestinal tract1. Typical symptoms of CDI include diarrhea, abdominal pain, and fatal pseudomembranous colitis, and it can sometimes lead to death1,2. Evidence has shown that host immune responses play a critical role in both the progression and outcome of this disease3. In addition to the immune response, the indigenous gut microbiota is crucial for the onset and pathogenesis of CDI4. In the past decade, both the number of cases and the mortality rate of CDI have increased significantly due to the emergence of a hypervirulent strain of C. difficile (BI/NAP1/027)5,6. A better understanding of underlying immune mechanisms and the role of microbiota during CDI will help lead to new therapeutic developments and advances, enabling better control of this epidemic.
Several animal models, such as the hamster and mouse, have been developed to provide insight into the immune defense against C. difficile7,8. However, the role of innate immune cells is still poorly understood, particularly since innate immune cell behavior is mainly derived from histological analysis or cultured cells in vitro. Therefore, establishing a transparent zebrafish model to reveal the innate immune response to C. difficile inside of a living vertebrate organism will facilitate such studies. Zebrafish larvae have a functional innate immune system, but they lack the adaptive immune system until 4&#8211;6 weeks after fertilization9. This unique feature makes zebrafish larvae an excellent model to study the isolated response and function of innate immune cells in CDI.
This report describes new methods using zebrafish larvae to study the interactions between C. difficile and innate immune cells, such as macrophages and neutrophils. First, a localized microinjection protocol that includes C. difficile inoculum and staining is presented. Using in vivo confocal time-lapse imaging, the response of neutrophils and macrophages towards the infection site is recorded, and the phagocytosis of bacteria by neutrophils and macrophages is observed. However, it has been reported that the injection itself causes tissue damage and leads to the recruitment of leukocytes independent of the bacteria10. Therefore, a noninvasive microgavage protocol to deliver C. difficile into the intestine of zebrafish larvae is subsequently described. Previous studies have demonstrated that indigenous gastrointestinal microbiota protect a host against the colonization of C. difficile11. Therefore, gnotobiotic zebrafish larvae are also used to predispose the zebrafish that are infected 12. Afterwards, intestinal dissection is performed to recover viable C. difficile and validate the duration of their presence in zebrafish intestinal tracts.

<table><tbody><tr><td>Agarose</td><td>Sigma-Aldrich</td><td>A2576</td><td>Ultra-low gelling agarose</td></tr><tr><td>Agarose low-melting (LM)</td><td>Pronadisa</td><td>8050</td><td>It is used in agarose plates</td></tr><tr><td>BacLight Red Bacterial Stain</td><td>Thermo Fisher Scientific</td><td>B35001</td><td>Fluorescent dye</td></tr><tr><td>Brain-Heart-Infusion Broth</td><td>Carl Roth GmbH</td><td>X916.1</td><td></td></tr><tr><td>Brass (wild-type)</td><td></td><td></td><td>deficient in melanin synthesis, used to generate stable transgenic lines</td></tr><tr><td>Calcium nitrate (Ca(NO3)2)</td><td>Sigma-Aldrich</td><td>C1396</td><td></td></tr><tr><td>Capillary Glass</td><td>Harvard Apparatus</td><td>30-0019</td><td>Injection needles</td></tr><tr><td>&lt;em&gt;Clostridioides difficile&lt;/em&gt;</td><td></td><td></td><td>R20291,, a ribotype 027 strain, TcdA+/TcdB+/CDT+ production</td></tr><tr><td>DMSO</td><td>Carl Roth GmbH</td><td>A994</td><td></td></tr><tr><td>FIJI</td><td>open-source platform</td><td></td><td>Image processing</td></tr><tr><td>HEPES</td><td>Carl Roth GmbH</td><td>6763</td><td></td></tr><tr><td>Horizontal needle puller</td><td>Sutter instrument Inc</td><td>P-87</td><td></td></tr><tr><td>L-cysteine</td><td>Sigma-Aldrich</td><td>168149</td><td></td></tr><tr><td>Leica Application Suite X (LAS X)</td><td>Leica</td><td></td><td>Image processing</td></tr><tr><td>Magnesium sulfate (MgSO4)</td><td>Carl Roth GmbH</td><td>P026</td><td></td></tr><tr><td>Micro injector</td><td>eppendorf</td><td>5253000017</td><td></td></tr><tr><td>Microinjection molds</td><td>Adaptive Science Tools</td><td>TU1</td><td></td></tr><tr><td>Leica SP8 confocal microscope</td><td>Leica</td><td></td><td></td></tr><tr><td>Phenol Red</td><td>Sigma-Aldrich</td><td>P0290</td><td></td></tr><tr><td>Potassium chloride (KCl)</td><td>Carl Roth GmbH</td><td>5346</td><td></td></tr><tr><td>Sodium chloride (NaCl)</td><td>Carl Roth GmbH</td><td>9265</td><td></td></tr><tr><td>Taurocholate</td><td>Carl Roth GmbH</td><td>8149</td><td></td></tr><tr><td>&lt;em&gt;Tg(lyZ: KalTA4)bz17/Tg(4xUAS-E1b:EGFP)hzm3&lt;/em&gt;</td><td></td><td></td><td>stable transgenic line in which in which the lyZ promoters drive the expression of EGFP fluorescent protein in neutrophils</td></tr><tr><td>&lt;em&gt;Tg(mpeg1.1: KalTA4)bz16/Tg(4xUAS-E1b:EGFP)&lt;sup&gt;hzm3&lt;/sup&gt;&lt;/em&gt;</td><td></td><td></td><td>stable transgenic line in which in which the mpeg1.1 drive the expression of EGFP fluorescent protein in macrophages</td></tr><tr><td>Tricaine</td><td>Sigma-Aldrich</td><td>E10521</td><td></td></tr><tr><td>Yeast extract</td><td>BD Bacto</td><td>212750</td><td></td></tr></tbody></table>

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.

C. difficile is strictly anaerobic, but the chromophore of fluorescent proteins usually requires oxygen to mature. To overcome this problem, a fluorescent dye was used to stain live C. difficile cells that were actively growing (R20291, a ribotype 027 strain; Figure 1A). Using the Gal4/UAS system, stable transgenic zebrafish lines were generated for live imaging, in which the mpeg1.1 or lyZ promoters drove the expression of EGFP fluorescen...

Watch this Scientific Journal Video about Development of a Larval Zebrafish Infection Model for Clostridioides difficile at JoVE.com

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.

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 ...

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Developmental Biology

6.2K Views.  Technische Universität Braunschweig. Presented here is a safe and effective method to infect zebrafish larvae with fluorescently labeled anaerobic C. difficile by microinjection and noninvasive microgavage.

Video: Development of a Larval Zebrafish Infection Model for Clostridioides difficile

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 ...

Immunology and Infection

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 ...

Establishment and Optimization of a High Throughput Setup to Study Staphylococcus epidermidis and Mycobacterium marinum Infection as a Model for Drug Discovery

Zebrafish are becoming a valuable tool in the preclinical phase of drug discovery screenings as a whole animal model with high throughput screening possibilities. They can be used to bridge the gap between cell based assays at earlier stages and in vivo validation in mammalian models, reducing, in this way, the number of compounds passing through to testing on the much more expensive rodent models. In this light, in the present manuscript is described a new high throughput pipeline using zebrafish as in vivo model system for the study of Staphylococcus epidermidis and Mycobacterium marinum infection. This setup allows the generation and analysis of large number of synchronous embryos homogenously infected. Moreover the flexibility of the pipeline allows the user to easily implement other platforms to improve the resolution of the analysis when needed. The combination of the zebrafish together with innovative high throughput technologies opens the field of drug testing and discovery to new possibilities not only because of the strength of using a whole animal model but also because of the large number of transgenic lines available that can be used to decipher the mode of action of new compounds.

Establishment and Optimization of a High Throughput Setup to Study Staphylococcus epidermidis and Mycobacterium marinum Infection as a Model for Drug Discovery

This video article describes the high throughput pipeline that has been successfully established to infect and analyze large numbers of zebrafish embryos providing a new powerful tool for compound testing and drug discovery using a whole animal vertebrate organism.

Zebrafish are becoming a valuable tool in the preclinical phase of drug discovery screenings as a whole animal model with high throughput screening ...

Modeling Mucosal Candidiasis in Larval Zebrafish by Swimbladder Injection

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

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

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

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.

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.

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

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 ...

Cefoperazone-treated Mouse Model of Clinically-relevant Clostridium difficile Strain R20291

Clostridium difficile is an anaerobic, gram-positive, spore-forming enteric pathogen that is associated with increasing morbidity and mortality and consequently poses an urgent threat to public health. Recurrence of a C. difficile infection (CDI) after successful treatment with antibiotics is high, occurring in 20-30% of patients, thus necessitating the discovery of novel therapeutics against this pathogen. Current animal models of CDI result in high mortality rates and thus do not approximate the chronic, insidious disease manifestations seen in humans with CDI. To evaluate therapeutics against C. difficile, a mouse model approximating human disease utilizing a clinically-relevant strain is needed. This protocol outlines the cefoperazone mouse model of CDI using a clinically-relevant and genetically-tractable strain, R20291. Techniques for clinical disease monitoring, C. difficile bacterial enumeration, toxin cytotoxicity, and histopathological changes throughout CDI in a mouse model are detailed in the protocol. Compared to other mouse models of CDI, this model is not uniformly lethal at the dose administered, allowing for the observation of a prolonged clinical course of infection concordant with the human disease. Therefore, this cefoperazone mouse model of CDI proves a valuable experimental platform to assess the effects of novel therapeutics on the amelioration of clinical disease and on the restoration of colonization resistance against C. difficile.

Cefoperazone-treated Mouse Model of Clinically-relevant Clostridium difficile Strain R20291

This protocol outlines the cefoperazone mouse model of Clostridium difficile infection (CDI) using a clinically-relevant and genetically-tractable strain, R20291. Emphasis on clinical disease monitoring, C. difficile bacterial enumeration, toxin cytotoxicity, and histopathological changes throughout CDI in a mouse model are detailed in the protocol.

Clostridium difficile is an anaerobic, gram-positive, spore-forming enteric pathogen that is associated with increasing morbidity and mortality and ...

Quantifying Vibrio cholerae Colonization and Diarrhea in the Adult Zebrafish Model

Vibrio cholerae is best known as the infectious agent that causes the human disease cholera. Outside the human host, V. cholerae primarily exists in the aquatic environment, where it interacts with a variety of higher aquatic species. Vertebrate fish are known to be an environmental host and are a potential V. cholerae reservoir in nature. Both V. cholerae and the teleost fish species Danio rerio, commonly known as zebrafish, originate from the Indian subcontinent, suggesting a long-standing interaction in aquatic environments. Zebrafish are an ideal model organism for studying many aspects of biology, including infectious diseases. Zebrafish can be easily and rapidly colonized by V. cholerae after exposure in water. Intestinal colonization by V. cholerae leads to the production of diarrhea and the excretion of replicated V. cholerae. These excreted bacteria can then go on to colonize new fish hosts. Here, we demonstrate how to assess V. cholerae-intestinal colonization in zebrafish and how to quantify V. cholerae-induced zebrafish diarrhea. The colonization model should be useful to researchers who are studying whether genes of interest may be important for host colonization and/or for environmental survival. The quantification of zebrafish diarrhea should be useful to researchers studying any intestinal pathogen who are interested in exploring zebrafish as a model system.

Quantifying Vibrio cholerae Colonization and Diarrhea in the Adult Zebrafish Model

Zebrafish are a natural Vibrio cholerae host and can be used to recapitulate and study the entire infectious cycle from colonization to transmission. Here, we demonstrate how to assess V. cholerae colonization levels and quantify diarrhea in zebrafish.

Vibrio cholerae is best known as the infectious agent that causes the human disease cholera. Outside the human host, V. cholerae primarily exists in the ...

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

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

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

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

Development of a Larval Zebrafish Infection Model for Clostridioides difficile

Summary

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Chapters in this video

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

Establishment and Optimization of a High Throughput Setup to Study Staphylococcus epidermidis and Mycobacterium marinum Infection as a Model for Drug Discovery

Modeling Mucosal Candidiasis in Larval Zebrafish by Swimbladder Injection

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

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

Cefoperazone-treated Mouse Model of Clinically-relevant Clostridium difficile Strain R20291

Quantifying Vibrio cholerae Colonization and Diarrhea in the Adult Zebrafish Model

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