We thank members of the Rawls laboratory for helpful suggestions on content and Dr. Alan Fanning for valuable discussions on tight junction size permeability and disruption methods. We also thank Dr. Michael Chua and Dr. Neal Kramarcy of the Michael Hooker Microscopy Facility for confocal microscope support. This work was supported by National Institutes of Health grants T32 DK007737-15 (J.L.C. trainee), F32 DK094592 (to J.L.C.), and R01 DK081426 (to J.F.R.).

We use published protocols for standard zebrafish husbandry and maintenance to obtain larvae for microgavage27. We primarily use natural breeding to propagate fish, and they are kept on a 14 hr light/10 hr dark cycle. This protocol has been optimized for microgavage of zebrafish larvae at 6-7 days post-fertilization (dpf). We anticipate that these methods will be generally applicable to younger and older developmental stages with minor modifications. Unless otherwise noted, we used the wild-type TL zebrafish strain at 6 dpf, and the fish were not fed. All experiments were performed in system water acquired from our conventional zebrafish aquaculture facility; however, any appropriate zebrafish media (e.g. embryo medium or gnotobiotic zebrafish medium (GZM)28) can be used. In this protocol, we refer to 'zebrafish media' to represent the user's media of choice.
1. Preparation of Methylcellulose, Gavage Mold, and Microgavage Needle Fabrication
The day before or early the day of the experiment, prepare the methylcellulose, agarose mold for larvae stabilization, and the microgavage needles.
<ol>
	<li>Dissolve 3 g of methylcellulose in 100 ml of sterile, deionized water or zebrafish media (depending on your application) to make a 3% solution. Add a magnetic stir bar to the bottle, heat water to close to boiling, and place on a stir plate at high heat. While stirring, slowly sprinkle in methylcellulose to ensure it does not clump. Slowly decrease the speed of stirring and temperature of the stir plate throughout the day until it is room temperature.</li>
	<li>Continue stirring slowly overnight until the methylcellulose is completely dissolved. Solution can be stored at room temperature for several months in a sealed container.</li>
	<li>Prepare a 3% agarose gel mold in a 10 cm Petri dish using a plastic cast (Adaptive Science Tools, TU-1) according to the manufacturer's instructions. Seal with parafilm and store at 4 &deg;C until needed to prevent desiccation. Warm up to room temperature or 28 &deg;C prior to experiment. Agarose molds can generally be re-used several times.</li>
	<li>Pull several gavage needles using borosilicate glass capillaries, and a Flaming Brown Micropipette Puller fitted with a wide-trough heat filament (FT330B). The programs we use for needle fabrication are as follows:</li>
</ol>
<table border="1">
	<tbody>
		<tr>
			<td>Needle type</td>
			<td>Pressure</td>
			<td>Heat</td>
			<td>Pull</td>
			<td>Velocity</td>
			<td>Time</td>
		</tr>
		<tr>
			<td>embryo</td>
			<td>500</td>
			<td>315</td>
			<td>100</td>
			<td>50</td>
			<td>200</td>
		</tr>
		<tr>
			<td>gavage</td>
			<td>500</td>
			<td>315</td>
			<td>125</td>
			<td>75</td>
			<td>200</td>
		</tr>
	</tbody>
</table>
Since filaments can vary between instruments, record the ramp time (active heat time). Our ramp time ranges between 8.57-8.79 ms for this program. This program creates shorter, stockier needles than traditional microinjection needles (Figure 1). These settings may need to be optimized for use in different micropipette pullers.
2. Clipping, Loading, and Calibration of the Microgavage Needles
Note*: The needle clipping is a critical step in this protocol. Also, the methods for needle loading and calibration will vary depending on the type of microinjection unit being used. We have optimized this protocol using Drummond II microinjectors, but we anticipate that this protocol will also be applicable to other microinjection instruments with minor modification. Since injection rigs and needle pullers will vary between labs, it will be important to measure/calibrate the injection volumes (see section 2.8).
<ol>
	<li>The needles are clipped under a stereomicroscope. We use a Leica S6E stereomicroscope set to 2.5x magnification. At this setting, our eyepiece graticle with 100 divisions is 2 mm in length.</li>
	<li>Align the tip of the needle with the end of the graticle ruler (division mark 100), and clip the needle using fine-tipped watchmaker forceps at ~1 mm from the tip (division mark 48-50) (Figure 1A).</li>
	<li>The needle should be ~27-30 &mu;m in diameter and blunt. Examine the needle tips at 100x magnification. Sharp or jagged needles lead to higher probability of damage during gavage and should be avoided if possible.</li>
	<li>Alternatively, when available, a microforge can be used to better control the needle clip point and to fire-polish the needle tips to remove sharp edges (Figure 1B).</li>
	<li>Prepare the gavage solution. For most applications, we prepare the solution with addition of a 1:10 dilution of a 0.5% phenol red solution in Dulbecco's phosphate buffered saline (DPBS) (Sigma-Aldrich) to ensure that the solution can be seen under the stereomicroscope to monitor proper functioning of the microinjector and placement of the gavage solution in the anterior bulb. Loading dyes other than phenol red have not been evaluated but could be used.</li>
	<li>Prepare the gavage needle by filling with mineral oil using a 1 ml syringe with a 25G 5/8-needle. Mount onto the Nanoject II microinjection unit.</li>
	<li>Wrap a plastic 10 cm Petri dish with parafilm to provide a flat, waterproof surface and aliquot ~2 &mu;l of the gavage solution onto the parafilm. Manually backfill the gavage needle being careful to avoid creating air bubbles in the mineral oil.</li>
	<li>To test microinjector function and calibrate the ejection volume, fill a 3 cm Petri dish with mineral oil and inject into the oil until the droplets are consistent in size. Measure the diameter of the droplets using the graticle or a measurement function on the stereomicroscope, if available. Calculate the droplet volume (Equation: V=4/3&pi;r3 ). Volume consistency between needles is plotted (Figure 1C).</li>
	<li>Rinse the tip of the needle by repeated submersions in clean zebrafish media to remove residual mineral oil before starting gavage. Mineral oil is very dense and may cause damage to epithelial tissues if not washed off of the needle tip.</li>
</ol>
3. Anesthetizing, Mounting, and Gavage of Zebrafish Larvae
<ol>
	<li>Warm the agarose mold at room temperature or 28 &deg;C prior to experiment.</li>
	<li>Prepare the agarose mold for gavage by covering 3 grooves with 3% methylcellulose. Use an amount of methylcellulose sufficient to cover the grooves and hold the fish, but not so scant that it dries out quickly.</li>
	<li>In a clean 6-well plate, aliquot zebrafish larvae for each experimental group into separate wells, each containing 3.5-4.0 ml of zebrafish media.</li>
	<li>Prepare 3x tricaine solution (0.05% w/v) in the appropriate zebrafish medium. Anesthetize fish one well at a time by mixing an equal volume of 3x tricaine into the well containing larvae for a final concentration of 1.5x (0.025%), and swirl gently.</li>
	<li>As soon as the zebrafish stop moving, remove them from the well using a wide bore glass Pasteur pipette and pipette pump, and place them one by one onto the methylcellulose with their heads on the 45&deg; angle of the groove and their tails laid toward the 90&deg; angle of the groove. Gently press them into the methylcellulose with a blunt dissection probe to stabilize their position.</li>
	<li>Adjust the Nanoject II microinjection rig such that the needle is tilted at a shallow angle that is approximately parallel to the 45&deg; angle of the agarose mold. Make sure that the extension range of the needle will be sufficient to reach past the bottom of the mold well so that fewer adjustments have to be made once fish are in place.</li>
	<li>Set the microinjection unit to release 4.6 nl (maximum) on the slow inject setting (23 nl/sec). At these settings, the delivered volume should just fill the anterior bulb of the intestine and not leak out of the esophagus or cloaca. Smaller volumes may be used depending on the purpose of the experiment.</li>
	<li>Use one hand to make minor adjustments of the plate containing the immobilized larvae while simultaneously using the other to control the manipulator of the microinjection unit.</li>
	<li>Gently maneuver the gavage needle into the mouth of anesthetized fish, through the esophagus, and slightly depress the esophageal sphincter to introduce the tip of the gavage needle just inside the anterior intestinal bulb (Figure 2, step 1-2, a&amp;b).</li>
	<li>Once inside the anterior bulb, gently depress the foot pedal or inject button to administer the material. Retract the needle quickly and smoothly trying not to release significant quantities of material into the esophagus (Figure 2, step 3-4).</li>
	<li>Following gavage, use a wide-bore glass Pasteur pipette and pipette pump to gently move the zebrafish larvae to fresh media. Fill the pipette with water to about 1 inch above the neck bend, make a small water pocket in the methylcellulose by the larva's head to release it, lift the head gently into the mouth of the pipette, and draw the larva out of the methylcellulose. Gently expel fish and all water from the pipette into a dish of fresh media.</li>
	<li>Rinse the larvae in fresh media by pipetting up and down gently or splashing with media several times to remove methylcellulose and revive the larvae from anesthesia. Transfer larvae into a Petri dish or 6-well plate until needed for subsequent imaging and analysis.</li>
</ol>
4. Dextran Assay to Test Gavage Safety and Intestinal Barrier Integrity
Microgavage can be used to deliver diverse materials into the intestine. As an example, we provide a protocol below for gavaging a 10 kD dextran conjugated to Texas Red to analyze barrier integrity of the intestinal epithelium. At least two potential routes of paracellular permeability exist, with differences in selectivity based on charge and size of the solutes, termed the "pore" or "leak" pathways29. Since this dextran is too large to be absorbed across the intestinal epithelium through the paracellular pore pathway (size limitation ~4&Aring;, 10kD dextran= ~23&Aring;)9,30-32, it should be retained within the lumen if intestinal barrier integrity is not compromised. Although 10 kD dextran could be transported via the paracellular leak pathway, this pathway has lower capacity and slower kinetics29,33, and, therefore, is not likely to occur within the time-frame of this assay. A 'positive control' treatment to disrupt barrier integrity is provided by inclusion of EDTA which disrupts epithelial tight junctions34,35. This dextran gavage assay can be used as a quality control test to demonstrate that the gavage procedure is being performed safely without unintended tissue damage. Additionally, this assay can be used to test intestinal barrier integrity as a function of genotype or treatment.
<ol>
	<li>Prepare the following gavage solutions: 1% dextran/1x PBS/0.05% phenol red and 1% dextran/20 mM EDTA/1x PBS/0.05% phenol red (positive control). The 5% dextran stock in ddH2O is stored in 50 &mu;l aliquots at -20 &deg;C to reduce the number of freeze-thaw cycles. Also, prepare a mock gavage solution of 1x PBS/0.05% phenol red.</li>
</ol>
<table border="1">
	<tbody>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>+ control (EDTA)</td>
			<td>no EDTA</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>stock</td>
			<td>final</td>
			<td>dilution factor</td>
			<td>10 &mu;l</td>
			<td>10 &mu;l</td>
		</tr>
		<tr>
			<td>EDTA, pH 8</td>
			<td>200 mM</td>
			<td>20 mM</td>
			<td>10</td>
			<td>1</td>
			<td>0</td>
		</tr>
		<tr>
			<td>Dextran</td>
			<td>5%</td>
			<td>1%</td>
			<td>5</td>
			<td>2</td>
			<td>2</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>10x</td>
			<td>1x</td>
			<td>10</td>
			<td>1</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Phenol Red</td>
			<td>0.50%</td>
			<td>0.05%</td>
			<td>10</td>
			<td>1</td>
			<td>1</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>subtotal:</td>
			<td>5</td>
			<td>4</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>ddH2O:</td>
			<td>5</td>
			<td>6</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Total:</td>
			<td>10</td>
			<td>10</td>
		</tr>
	</tbody>
</table>
<ol start="2">
	<li>Use the protocol in section 3 to gavage groups of 10-20 zebrafish larvae for each experimental condition. Repeat this on duplicate or triplicate groups if possible.</li>
	<li>After gavage, recover larvae from anesthesia and allow them to swim freely in fresh media.</li>
	<li>At 18-20 min post-gavage, re-anesthetize the larvae as in step 3.3. Position the larvae in 3% methylcellulose on top of a 3% agarose block (this reduces background glare during imaging).</li>
	<li>Image the larvae with a fluorescence stereoscope (e.g., Leica M205C) and a Texas Red filter set at a magnification that displays the larvae from the snout to immediately posterior to the end of the cloaca.</li>
	<li>Acquire all images at the same exposure settings so that they can be quantified and compared later. Fluorescence will be intense in the intestinal lumen of all dextran-injected larvae, but barrier function is evaluated by the level of dextran that appears in extraintestinal tissues (i.e., the trunk and blood vessels) (Figure 5). Adjust the exposure to settings that enable visualization of trunk fluorescence even if this over-exposes lumen fluorescence.</li>
	<li>Stagger the gavage times so that each group spends approximately the same amount of time in recovery after gavage and is imaged at the same approximate time post-gavage. For this experiment, each group takes approximately 1 hr for one person to complete. Fish are anesthetized at 18-20 min post-gavage, and imaging takes ~1-2 min per fish.</li>
	<li>Use an image analysis program such as ImageJ to quantify the relative mean fluorescence intensity in a region of the trunk just above the intestine. Normalize these values by subtracting a background measurement taken in a region of the image outside of the fish.</li>
	<li>Graph and analyze data from image analysis using GraphPad Prism or similar statistical software.</li>
</ol>
5. Fluorescent Microsphere Assay to Test Intestinal Motility
Gavage provides enhanced control over timing and amount of material delivery to the intestine which makes it an ideal alternative method for intestinal motility studies. Previously published techniques for measuring intestinal motility in zebrafish larvae include direct observation of intestinal contractions and waves by microscopy1,21,36, or feeding larvae with food blended with yellow-green fluorescent polystyrene microspheres26. In some studies, quantitation of intestinal motility patterns has been enhanced by immersing larvae in media containing food dye37 or a specialized video analysis technique called spatiotemporal mapping38,39. Here we provide a proof-of-concept study to show that it is possible to gavage fluorescent microspheres into the intestinal anterior bulb, and tracking of microsphere movement can be utilized to assess intestinal motility (Figure 6). The primary difference between this method and the approach of Field and colleagues26 is that the beads are introduced alone and cannot be incorporated into food due to the gavage needle diameter limits.
<ol>
	<li>Maintain zebrafish larvae in 10 cm plates with 35 ml media until the experiment is performed at 7 dpf. Begin feeding larvae once each morning starting at 5 dpf with ~0.6 mg powdered food (larvae diet, as previously described40; administered using a sterile, 1 mm inoculating loop).</li>
	<li>Gently shake the bottle of Fluoresbrite YG 2.0 &mu;m polystyrene microspheres (~2.5% aqueous suspension) to mix the suspension. Aliquot a drop from the bottle onto parafilm so that a specific volume can be measured by pipetting.</li>
	<li>In an eppendorf tube, centrifuge the beads at low speed in a benchtop microfuge, remove the supernatant, and replace with 1x PBS at the original volume. Repeat this process to wash the microspheres 2 times.</li>
	<li>Prepare a gavage suspension containing 0.25% microspheres (1:10 dilution of stock suspension)/1x PBS/0.05% phenol red. There are ~2.6x103 microspheres in the 4.6 nl gavage volume administered per larvae.</li>
	<li>Use the protocol in section 3 to gavage the desired number of larvae. Microspheres are somewhat buoyant; therefore, gavage should be performed as rapidly as possible to maximize consistency of the suspension.</li>
	<li>In this experiment, 15 larvae were gavaged and then maintained in single wells of a 12-well plate following recovery from anesthesia in order to follow transit in individuals. However, larger groups can also be gavaged and transit assessed as a population rather than individually. Refer to Field et al. for standard practices26.</li>
	<li>Assess intestinal motility by scoring the most rostral (or anterior) location of the fluorescent microspheres in live larvae at different times using a fluorescent stereomicroscope. Briefly anesthetize the larvae in 1x (0.017%) tricaine, and score the zones of microsphere location or capture images to score later.</li>
	<li>Graph and compare the percent of total larvae with microspheres in a certain intestinal region over time.</li>
</ol>

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No conflicts of interest declared.

In this work, we describe a novel protocol for direct delivery of materials to the larval zebrafish intestine by microgavage. There are several critical steps throughout the procedure that should be kept in mind. First, the zebrafish larvae should be in good health before the gavage experiment to prevent death unrelated to treatment. Another important factor is the quality of the gavage needle. One of the most difficult parts of the protocol is clipping the needle at the appropriate location without creating sharp, jagge...

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Finquel (Tricaine methanesulfonate)</td>
			<td>Argent Chemical Laboratories</td>
			<td>MS-222</td>
			<td>Larvae anesthesia</td>
		</tr>
		<tr>
			<td>Phenol Red Solution</td>
			<td>Sigma-Aldrich</td>
			<td>P0290-100 ml</td>
			<td>0.5% in DPBS, cell-culture tested</td>
		</tr>
		<tr>
			<td>Mineral oil, light, white (high purity)</td>
			<td>Amresco</td>
			<td>J217-500 ml</td>
			<td>For needle backfill and volume testing</td>
		</tr>
		<tr>
			<td>TxRed-Dextran, 10,000 MW, lysine fixable</td>
			<td>Invitrogen/ Molecular Probes</td>
			<td>D-1863</td>
			<td>1% suspension in 1x PBS/phenol red</td>
		</tr>
		<tr>
			<td>FM4-64FX</td>
			<td>Invitrogen/ Molecular Probes</td>
			<td>F34653</td>
			<td>5 mM stock in water</td>
		</tr>
		<tr>
			<td>Methylcellulose</td>
			<td>MP Biochemicals</td>
			<td>0215549590</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Borosilicate glass capillaries</td>
			<td>Drummond Scientific</td>
			<td>3-000-203-G/X</td>
			<td>OD 1.14 mm, ID 0.53 mm, 3.5 in length</td>
		</tr>
		<tr>
			<td>Plastic form for mold making</td>
			<td>Adaptive Science Tools</td>
			<td>TU-1</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Nanoject II microinjection unit</td>
			<td>Drummond Scientific</td>
			<td>3-000-204</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Flaming Brown Micropipette Puller, 3.0 mm wide-trough filament</td>
			<td>Sutter Instrument Co.</td>
			<td>P-97, FT330B</td>
			<td>Needle fabrication</td>
		</tr>
		<tr>
			<td>Student Dumont #5 forceps</td>
			<td>Fine Science Tools</td>
			<td>91150-20</td>
			<td>0.1 x 0.06 mm, needle clipping</td>
		</tr>
		<tr>
			<td>Eyepiece Graticle, 5 mm - 100 divisions</td>
			<td>Leica</td>
			<td>10394771</td>
			<td>Needle clipping</td>
		</tr>
		<tr>
			<td>Microforge</td>
			<td>Narishige</td>
			<td>MF-900</td>
			<td>Needle clipping and fire-polishing</td>
		</tr>
		<tr>
			<td>Tuberculin SlipTip syringe needle</td>
			<td>Becton Dickinson</td>
			<td>309626</td>
			<td>1 ml, 25G 5/8-needle</td>
		</tr>
		<tr>
			<td>Fluoresbrite YG Microspheres (2.0 &mu;m)</td>
			<td>Polysciences, Inc.</td>
			<td>18338</td>
			<td>Intestinal motility analysis</td>
		</tr>
		<tr>
			<td>Dissecting Microscope</td>
			<td>Leica</td>
			<td>S6E</td>
			<td>Gavage procedure</td>
		</tr>
		<tr>
			<td>Fluorescence Stereomicroscope</td>
			<td>Leica</td>
			<td>M205C</td>
			<td>Dextran barrier assay</td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Zeiss</td>
			<td>LSM510</td>
			<td>Imaging of dextran in circulation</td>
		</tr>
	</tbody>
</table>

microgavage of zebrafish larvae

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe interactions17-25. Experimental approaches for studying intestinal biology often require the in vivo introduction of selected materials into the lumen of the intestine. In the larval zebrafish model, this is typically accomplished by immersing fish in a solution of the selected material, or by injection through the abdominal wall. Using the immersion method, it is difficult to accurately monitor or control the route or timing of material delivery to the intestine. For this reason, immersion exposure can cause unintended toxicity and other effects on extraintestinal tissues, limiting the potential range of material amounts that can be delivered into the intestine. Also, the amount of material ingested during immersion exposure can vary significantly between individual larvae26. Although these problems are not encountered during direct injection through the abdominal wall, proper injection is difficult and causes tissue damage which could influence experimental results. We introduce a method for microgavage of zebrafish larvae. The goal of this method is to provide a safe, effective, and consistent way to deliver material directly to the lumen of the anterior intestine in larval zebrafish with controlled timing. Microgavage utilizes standard embryo microinjection and stereomicroscopy equipment common to most laboratories that perform zebrafish research. Once fish are properly positioned in methylcellulose, gavage can be performed quickly at a rate of approximately 7-10 fish/ min, and post-gavage survival approaches 100% depending on the gavaged material. We also show that microgavage can permit loading of the intestinal lumen with high concentrations of materials that are lethal to fish when exposed by immersion. To demonstrate the utility of this method, we present a fluorescent dextran microgavage assay that can be used to quantify transit from the intestinal lumen to extraintestinal spaces. This test can be used to verify proper execution of the microgavage procedure, and also provides a novel zebrafish assay to examine intestinal epithelial barrier integrity under different experimental conditions (e.g. genetic manipulation, drug treatment, or exposure to environmental factors). Furthermore, we show how gavage can be used to evaluate intestinal motility by gavaging fluorescent microspheres and monitoring their subsequent transit. Microgavage can be applied to deliver diverse materials such as live microorganisms, secreted microbial factors/toxins, pharmacological agents, and physiological probes. With these capabilities, the larval zebrafish microgavage method has the potential to enhance a broad range of research fields using the zebrafish model system.

When gavage is performed properly, the delivered material should be entirely contained within the anterior intestine with little to no residual material in the esophagus (Figure 3). A delivery volume should be chosen that can be accommodated within the anterior bulb and does not leak out through cloaca or esophagus. If the volume of material or pressure of delivery is too high, then the physical force of gavage may lead to damage of or leakage through the epithelium. It may also push out or alter other c...

Watch this Scientific Journal Video about Microgavage of Zebrafish Larvae at JoVE.com

Microgavage of Zebrafish Larvae

We present a novel method for microgavage of larval zebrafish utilizing standard embryo microinjection and stereomicroscopy equipment. We demonstrate that microgavage is a safe and efficient technique useful for delivering controlled amounts of diverse materials specifically into the larval zebrafish intestinal lumen.

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe ...

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16.6K Views.  University of North Carolina at Chapel Hill. We present a novel method for microgavage of larval zebrafish utilizing standard embryo microinjection and stereomicroscopy equipment. We demonstrate that microgavage is a safe and efficient technique useful for delivering controlled amounts of diverse materials specifically into the larval zebrafish intestinal lumen.