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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>Clostridioides difficile</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>Tg(lyZ: KalTA4)bz17/Tg(4xUAS-E1b:EGFP)hzm3</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>Tg(mpeg1.1: KalTA4)bz16/Tg(4xUAS-E1b:EGFP)hzm3</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 ...

development-larval-zebrafish-infection-model-for-clostridioides

Research

JoVE Journal

Developmental Biology

6.1K 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

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration have been thoroughly analyzed in vitro. However, in vivo cell migration somehow differs from in vitro migration, and has proven more difficult to analyze, being less accessible to direct observation and manipulation. This protocol uses the migration of the prospective prechordal plate in the early zebrafish embryo as a model system to study the function of candidate genes in cell migration. Prechordal plate progenitors form a group of cells which, during gastrulation, undergoes a directed migration from the embryonic organizer to the animal pole of the embryo. The proposed protocol uses cell transplantation to create mosaic embryos. This offers the combined advantages of labeling isolated cells, which is key to good imaging, and of limiting gain/loss of function effects to the observed cells, hence ensuring cell-autonomous effects. We describe here how we assessed the function of the TORC2 component Sin1 in cell migration, but the protocol can be used to analyze the function of any candidate gene in controlling cell migration in vivo.

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Combining cell transplantation, cytoskeletal labeling and loss/gain of function approaches, this protocol describes how the migrating zebrafish prospective prechordal plate can be used to analyze the function of a candidate gene in in vivo cell migration.

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration ...

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

Development of an Ethanol-induced Fibrotic Liver Model in Zebrafish to Study Progenitor Cell-mediated Hepatocyte Regeneration

Sustained liver fibrosis with continuation of extracellular matrix (ECM) protein build-up results in the loss of cellular competency of the liver, leading to cirrhosis with hepatocellular dysfunction. Among multiple hepatic insults, alcohol abuse can lead to significant health problems including liver failure and hepatocellular carcinoma. Nonetheless, the identity of endogenous cellular sources that regenerate hepatocytes in response to alcohol has not been properly investigated. Moreover, few studies have effectively modeled hepatocyte regeneration upon alcohol-induced injury. We recently reported on establishing an ethanol (EtOH)-induced fibrotic liver model in zebrafish in which hepatic progenitor cells (HPCs) gave rise to hepatocytes upon near-complete hepatocyte loss in the presence of fibrogenic stimulus. Furthermore, through chemical screens using this model, we identified multiple small molecules that enhance hepatocyte regeneration. Here we describe in detail the procedures to develop an EtOH-induced fibrotic liver model and to perform chemical screens using this model in zebrafish. This protocol will be a critical tool to delineate the molecular and cellular mechanisms of how hepatocyte regenerates in the fibrotic liver. Furthermore, these methods will facilitate potential discovery of novel therapeutic strategies for chronic liver disease in vivo.

Sustained fibrosis with deposition of excessive extracellular matrix proteins leads to cirrhosis. Alcohol abuse is one of the main causes of severe liver disease. We established an ethanol-induced zebrafish fibrotic liver model to study the mechanisms and strategies of promoting hepatocyte regeneration upon alcohol-induced injury.

Sustained liver fibrosis with continuation of extracellular matrix (ECM) protein build-up results in the loss of cellular competency of the liver, leading ...

A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. In particular, posterior body elongation is an essential process in embryonic development by which multiple tissue deformations act together to direct the formation of a large part of the body axis. In order to observe this process by long-term time-lapse imaging it is necessary to utilize a mounting technique that allows sufficient support to maintain samples in the correct orientation during transfer to the microscope and acquisition. In addition, the mounting must also provide sufficient freedom of movement for the outgrowth of the posterior body region without affecting its normal development. Finally, there must be a certain degree in versatility of the mounting method to allow imaging on diverse imaging set-ups. Here, we present a mounting technique for imaging the development of posterior body elongation in the zebrafish D. rerio. This technique involves mounting embryos such that the head and yolk sac regions are almost entirely included in agarose, while leaving out the posterior body region to elongate and develop normally. We will show how this can be adapted for upright, inverted and vertical light-sheet microscopy set-ups. While this protocol focuses on mounting embryos for imaging for the posterior body, it could easily be adapted for the live imaging of multiple aspects of zebrafish development.

Here, we present a versatile mounting method that allows for the long-term time-lapse imaging of the posterior body development of live zebrafish embryos without perturbing normal development.

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. ...

Dissection of Larval Zebrafish Gonadal Tissue

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex determination system, where several genes distributed throughout the genome collectively determine the sex identities of individual fish. Currently, the genes involved in regulating gonad development and how they work remain elusive. Normally, isolating gonadal tissue is the first step to examine the sex developmental processes. Here, we present a procedure to isolate gonadal tissue from 17 dpf (days post fertilization) and 25 dpf zebrafish larvae. The isolated gonadal tissue may be subsequently examined by morphology and gene expression profiling.

Here, we present a protocol for isolating gonadal tissue of larval zebrafish, which will facilitate investigations of zebrafish sex differentiation and maintenance.

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

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 ...

Electroretinogram Recording in Larval Zebrafish using A Novel Cone-Shaped Sponge-tip Electrode

The zebrafish (Danio rerio) is commonly used as a vertebrate model in developmental studies and is particularly suitable for visual neuroscience. For functional measurements of visual performance, electroretinography (ERG) is an ideal non-invasive method, which has been well established in higher vertebrate species. This approach is increasingly being used for examining the visual function in zebrafish, including during the early developmental larval stages. However, the most commonly used recording electrode for larval zebrafish ERG to date is the glass micropipette electrode, which requires specialized equipment for its manufacture, presenting a challenge for laboratories with limited resources. Here, we present a larval zebrafish ERG protocol using a cone-shaped sponge-tip electrode. The novel electrode is easier to manufacture and handle, more economical, and less likely to damage the larval eye than the glass micropipette. Like previously published ERG methods, the current protocol can assess outer retinal function through photoreceptor and bipolar cell responses, the a- and b-wave, respectively. The protocol can clearly illustrate the refinement of visual function throughout the early development of zebrafish larvae, supporting the utility, sensitivity, and reliability of the novel electrode. The simplified electrode is particularly useful when establishing a new ERG system or modifying existing small-animal ERG apparatus for zebrafish measurement, aiding researchers in the visual neurosciences to use the zebrafish model organism.

Here, we present a protocol that simplifies the measurement of light evoked electroretinogram responses from larval zebrafish. A novel cone-shaped sponge-tip electrode can help to make the study of visual development in larval zebrafish using the electroretinogram ERG easier to achieve with reliable outcomes and lower cost.

The zebrafish (Danio rerio) is commonly used as a vertebrate model in developmental studies and is particularly suitable for visual neuroscience. For ...

Quantifying Liver Size in Larval Zebrafish Using Brightfield Microscopy

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval liver size in zebrafish HCC models provides a means to rapidly assess the effects of drugs and other manipulations on an oncogene-related phenotype. Here we show how to fix zebrafish larvae, dissect the tissues surrounding the liver, photograph livers using bright-field microscopy, measure liver area, and analyze results. This protocol enables rapid, precise quantification of liver size. As this method involves measuring liver area, it may underestimate differences in liver volume, and complementary methodologies are required to differentiate between changes in cell size and changes in cell number. The dissection technique described herein is an excellent tool to visualize the liver, gut, and pancreas in their natural positions for myriad downstream applications including immunofluorescence staining and in situ hybridization. The described strategy for quantifying larval liver size is applicable to many aspects of liver development and regeneration.

Here we demonstrate a method for quantifying liver size in larval zebrafish, providing a way to assess the effects of genetic and pharmacologic manipulations on liver growth and development.

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval ...

A Simple and Effective Transplantation Device for Zebrafish Embryos

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex developmental processes. Zebrafish embryos are ideally suited for these manipulations since they are easily accessible, relatively large in size, and transparent. However, previously developed devices for cell removal and transplantation are cumbersome to use or expensive to purchase. In contrast, the transplantation device presented here is economical, easy to assemble, and simple to use. In this protocol, we first introduce the handling of the transplantation device as well as its assembly from commercially and widely available parts. We then present three applications for its use: generation of ectopic clones to study signal dispersal from localized sources, extirpation of cells to produce size-reduced embryos, and germline transplantation to generate maternal-zygotic mutants. Finally, we show that the tool can also be used for embryological manipulations in other species such as the Japanese rice fish medaka.

Embryological manipulations such as extirpation and transplantation of cells are important tools to study early development. This protocol describes a simple and effective transplantation device to perform these manipulations in zebrafish embryos.

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex ...