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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M., Akerlund, J., Mittge, E., Guillemin, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intestinal+alkaline+phosphatase+detoxifies+lipopolysaccharide+and+prevents+inflammation+in+zebrafish+in+response+to+the+gut+microbiota.">Intestinal alkaline phosphatase detoxifies lipopolysaccharide and prevents inflammation in zebrafish in response to the gut microbiota.</a> Cell Host Microbe. 2, 371-382 (2007).</li><li>Camp, J. G., Jazwa, A. L., Trent, C. M., Rawls, J. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intronic+cis-regulatory+modules+mediate+tissue-specific+and+microbial+control+of+angptl4/fiaf+transcription.">Intronic cis-regulatory modules mediate tissue-specific and microbial control of angptl4/fiaf transcription.</a> PLoS Genetics. , (2012).</li><li>Cheesman, S. E., Neal, J. T., Mittge, E., Seredick, B. M., Guillemin, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epithelial+cell+proliferation+in+the+developing+zebrafish+intestine+is+regulated+by+the+Wnt+pathway+and+microbial+signaling+via+Myd88.">Epithelial cell proliferation in the developing zebrafish intestine is regulated by the Wnt pathway and microbial signaling via Myd88.</a> Proc. Natl. Acad. Sci. U.S.A. 108, 4570-4577 (2011).</li><li>Field, H. A., Kelley, K. A., Martell, L., Goldstein, A. M., Serluca, F. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+gastrointestinal+physiology+using+a+novel+intestinal+transit+assay+in+zebrafish.">Analysis of gastrointestinal physiology using a novel intestinal transit assay in zebrafish.</a> Neurogastroenterol. Motil. 21, 304-312 (2009).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Zebrafish Book. , (2000).</li><li>Pham, L. N., Kanther, M., Semova, I., Rawls, J. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+generating+and+colonizing+gnotobiotic+zebrafish.">Methods for generating and colonizing gnotobiotic zebrafish.</a> Nat. Protoc. 3, 1862-1875 (2008).</li><li>Shen, L., Weber, C. R., Raleigh, D. R., Yu, D., Turner, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tight+junction+pore+and+leak+pathways:+a+dynamic+duo.">Tight junction pore and leak pathways: a dynamic duo.</a> Annu. Rev. Physiol. 73, 283-309 (2011).</li><li>Van Itallie, C. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+density+of+small+tight+junction+pores+varies+among+cell+types+and+is+increased+by+expression+of+claudin-2.">The density of small tight junction pores varies among cell types and is increased by expression of claudin-2.</a> J. Cell. Sci. 121, 298-305 (2008).</li><li>Van Itallie, C. M., Anderson, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measuring+size-dependent+permeability+of+the+tight+junction+using+PEG+profiling.">Measuring size-dependent permeability of the tight junction using PEG profiling.</a> Methods Mol. Biol. 762, 1-11 (2011).</li><li>Watson, C. J., Rowland, M., Warhurst, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+modeling+of+tight+junctions+in+intestinal+cell+monolayers+using+polyethylene+glycol+oligomers.">Functional modeling of tight junctions in intestinal cell monolayers using polyethylene glycol oligomers.</a> Am. J. Physiol. Cell Physiol. 281, 388-397 (2001).</li><li>Rodgers, L. S., Fanning, A. 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Biol. 207, 4085-4094 (2004).</li><li>Rich, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kit-like+immunoreactivity+in+the+zebrafish+gastrointestinal+tract+reveals+putative.">Kit-like immunoreactivity in the zebrafish gastrointestinal tract reveals putative.</a> 236, 903-911 (2007).</li><li>Holmberg, A., Olsson, C., Hennig, G. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=TTX-sensitive+and+TTX-insensitive+control+of+spontaneous+gut+motility+in+the+developing+zebrafish+(Danio+rerio)+larvae.">TTX-sensitive and TTX-insensitive control of spontaneous gut motility in the developing zebrafish (Danio rerio) larvae.</a> J. Exp. Biol. 210, 1084-1091 (2007).</li><li>Hennig, G. W., Costa, M., Chen, B. N., Brookes, S. 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Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+and+molecular+analyses+of+vascular+tube+and+lumen+formation+in+zebrafish.">Cellular and molecular analyses of vascular tube and lumen formation in zebrafish.</a> Development. 132, 5199-5209 (2005).</li><li>Berghmans, S., Hunt, J., Roach, A., Goldsmith, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+offer+the+potential+for+a+primary+screen+to+identify+a+wide+variety+of+potential+anticonvulsants.">Zebrafish offer the potential for a primary screen to identify a wide variety of potential anticonvulsants.</a> Epilepsy Res. 75, 18-28 (2007).</li></ol>

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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16.8K 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.

Video: Microgavage of Zebrafish Larvae

Dissection of Imaginal Discs from 3rd Instar Drosophila Larvae

This protocol demonstrates the dissection technique used for isolating imaginal discs from drosophila larvae at the 3rd instar stage. Methods for fixing the tissue after saying and removing the wing, leg, and eye discs are demonstrated directly on a microscope slide for subsequent visualization.

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

Protocols for Oral Infection of Lepidopteran Larvae with Baculovirus

Baculoviruses are widely used both as protein expression vectors and as insect pest control agents. This video shows how lepidopteran larvae can be infected with polyhedra by droplet feeding and diet plug-based bioassays. This accompanying Springer Protocols section provides an overview of the baculovirus lifecycle and use of baculoviruses as insecticidal agents, including discussion of the pros and cons for use of baculoviruses as insecticides, and progress made in genetic enhancement of baculoviruses for improved insecticidal efficacy.

In this video, we demonstrate oral infection techniques of lepidopteran larvae with baculovirus in order to determine insecticidal efficiency.

Baculoviruses are widely used both as protein expression vectors and as insect pest control agents. This video shows how lepidopteran larvae can be ...

Protocols for Microapplicator-assisted Infection of Lepidopteran Larvae with Baculovirus

Baculoviruses are widely used both as protein expression vectors and as insect pest control agents. . This video shows how lepidopteran larvae can be infected with microapplicator techniques in the gut with baculovirus polyhedra and in the hemolymph with budded virus. This accompanying Springer Protocols section provides an overview of the baculovirus lifecycle and use of baculoviruses as insecticidal agents.  Formulation and application of baculoviruses for pest control purposes are described elsewhere.

In this video, we demonstrate two microapplicator techniques used to infect of lepidopteran larvae with baculovirus in order to determine insecticidal efficiency.

Baculoviruses are widely used both as protein expression vectors and as insect pest control agents. . This video shows how lepidopteran larvae can be ...

Zebrafish Brain Ventricle Injection

Proper brain ventricle formation during embryonic brain development is required for normal brain function.  Brain ventricles are the highly conserved cavities within the brain that are filled with cerebrospinal fluid.  In zebrafish, after neural tube formation, the neuroepithelium undergoes a series of constrictions and folds while it fills with fluid resulting in brain ventricle formation.  In order to understand the process of ventricle formation, and the neuroepithelial shape changes that occur at the same time, we needed a way to visualize the ventricle space in comparison to the brain tissue.  However, the nature of transparent zebrafish embryos makes it difficult to differentiate the tissue from the ventricle space.  Therefore, we developed a brain ventricle injection technique where the ventricle space is filled with a fluorescent dye and imaged by brightfield and fluorescent microscopy.  The brightfield and the fluorescent images are then processed and superimposed in Photoshop.  This technique allows for visualization of the ventricle space with the fluorescent dye, in comparison to the shape of the neuroepithelium in the brightfield image.  Brain ventricle injection in zebrafish can be employed from 18 hours post fertilization through early larval stages.  We have used this technique extensively in our studies of brain ventricle formation and morphogenesis as well as in characterizing brain morphogenesis mutants (1-3).

After neural tube formation, the neuroepithelium constricts and folds while the tube fills with embryonic cerebrospinal fluid (eCSF) to form the embryonic brain ventricles. We developed this ventricle injection technique to better visualize the fluid filled space in contrast to the neuroepithelial shape in a live embryo.

Proper brain ventricle formation during embryonic brain development is required for normal brain function.  Brain ventricles are the highly conserved ...

Intravenous Microinjections of Zebrafish Larvae to Study Acute Kidney Injury

In this video article we describe a zebrafish model of AKI using gentamicin as the nephrotoxicant.  The technique consists of intravenous microinjections on 2 dpf zebrafish. This technique represents an efficient and rapid method to deliver soluble substances into the bloodstream of zebrafish larvae, allowing for the injection of 15-20 fish per hour. In addition to AKI studies, this microinjection technique can also be used for other types of experimental studies such as angiography.  We provide a detailed protocol of the technique from equipment required to visual measures of decreased kidney function. In addition, we also demonstrate the process of fixation, whole mount immunohistochemistry with a kidney tubule marker, plastic embedding and sectioning of the larval zebrafish. We demonstrate that zebrafish larvae injected with gentamicin show morphological features consistent with AKI: edema, loss of cell polarity in proximal tubular epithelial cells, and morphological disruption of the tubule.

We describe a technique of microinjecting the aminoglycoside, gentamicin, into 2 days post-fetilization (dpf) zebrafish larvae to induce acute kidney injury (AKI). We also describe a method for whole mount immunohistochemistry, plastic embedding and sectioning of zebrafish larvae to visualize the AKI mediated damage.

In this video article we describe a zebrafish model of AKI using gentamicin as the nephrotoxicant.  The technique consists of intravenous microinjections ...

Intraperitoneal Injection into Adult Zebrafish

A convenient method for chemically treating zebrafish is to introduce the reagent into the tank water, where it will be taken up by the fish. However, this method makes it difficult to know how much reagent is absorbed or taken up per fish. Some experimental questions, particularly those related to metabolic studies, may be better addressed by delivering a defined quantity to each fish, based on weight. Here we present a method for intraperitoneal (IP) injection into adult zebrafish. Injection is into the abdominal cavity, posterior to the pelvic girdle. This procedure is adapted from veterinary methods used for larger fish. It is safe, as we have observed zero mortality. Additionally, we have seen bleeding at the injection site in only 5 out of 127 injections, and in each of those cases the bleeding was brief, lasting several seconds, and the quantity of blood lost was small. Success with this procedure requires gentle handling of the fish through several steps including fasting, weighing, anesthetizing, injection, and recovery. Precautions are required to minimize stress throughout the procedure. Our precautions include using a small injection volume and a 35G needle. We use Cortland salt solution as the vehicle, which is osmotically balanced for freshwater fish. Aeration of the gills is maintained during the injection procedure by first bringing the fish into a surgical plane of anesthesia, which allows slow operculum movements, and second, by holding the fish in a trough within a water-saturated sponge during the injection itself. We demonstrate the utility of IP injection by injecting glucose and monitoring the rise in blood glucose level and its subsequent return to normal. As stress is known to increase blood glucose in teleost fish, we compare blood glucose levels in vehicle-injected and non-injected adults and show that the procedure does not cause a significant rise in blood glucose.

We demonstrate intraperitoneal injection into adult zebrafish. We use a 10 &mu;l NanoFil microsyringe controlled by a Micro4 controller and UltraMicroPump III. This demonstration includes the use of cold water as an anesthetic.

A convenient method for chemically treating zebrafish is to introduce the reagent into the tank water, where it will be taken up by the fish. However, ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...