The authors would like to acknowledge primary project funding from the Morgridge Institute for Research and the Laboratory for Optical and Computational Instrumentation. We also acknowledge funding from NIH# R01GM102924 (AH and KWE). KH, JMS, RS, AH and KWE conceived and designed the study. KH and JMS performed all experiments with support from DL, KP and RS. KH, JS, RS, AH and KWE contributed to the writing of the manuscript.

Note: The base zWEDGI design was formulated for zebrafish larvae that are 2 to 4 days post-fertilization (dpf) and follow the guidelines of the University of Wisconsin-Madison Research Animals Resource Center.
1. Design and 3D Printing of Molds
<ol>
	<li>Model the PDMS component of the device with desired geometries and attributes in a 3D modeling software5. Create an assembly of a blank mold and the PDMS part and generate a negative mold for the PDMS part by creating a cavity in the mold corresponding to the PDSM part. Save the mold as an .stl file for 3D printing (Figure 2, Step 1.1). 
	NOTE: A stereolithography format (.stl) file of the mold design presented here (Figure 1) is available for download at https://morgridge.org/designs/.</li>
	<li>Print molds using a photopolymer 3D printer (Figure 2, Step 1.2). Make multiple molds in a single printing, if possible, so more than one device can be molded with a single batch of PDMS. 
	NOTE: The example shown was printed in Hi-Res mode with a 0.075 mm beam diameter and 0.05 mm layer thickness, using photopolymer resin5.
	<ol>
		<li>Clean molds using a fine brush, denatured alcohol in a spray bottle, and compressed air to gently scrub and remove uncured resin. Remove any material from the microchannel regions.</li>
		<li>Post-cure the molds in a UV post cure apparatus for 60 min on each side as uncured resign is toxic to zebrafish larvae10.</li>
	</ol>
	</li>
	<li>Sand the cavity side of the mold with 200 grit sandpaper on a flat surface until all the sealing surfaces are in contact with the sandpaper (loading channel geometries and mold perimeter). Lightly sand with 400 and 600 grit sand paper, progressively, to produce smooth flush surfaces across all geometry facings (Figure 2, Step 1.3). Measure the depth of the cavity after sanding with a dial indicator to verify that it is close to the designed depth.
	<ol>
		<li>Clean molds and cover discs (1 &#190; inch diameter x &#188; inch thick borosilicate glass or acrylic; one glass cover per mold) by placing in an ultrasonic cleaner filled with water for 30 min or by flushing under running water.</li>
		<li>Blow dry with compressed air and clean both the molds and covers with isopropyl alcohol and filtered compressed air. Use a clean bench as a place to fabricate the devices to minimize contamination from airborne debris. Leave the cleaned molds and glass covers in the clean bench or a covered petri dish until needed.</li>
	</ol>
	</li>
</ol>
2. PDMS Fabrication of zWEDGI Device
<ol>
	<li>Make the PDMS by pouring 184 silicone elastomer polydimethylsiloxane (PDMS) at a ratio of 5:1 (base to activator) into a plastic cup. Mix well for 2 min with a wooden craft stick, stirring the gel over onto itself, like kneading bread. For 5 molds, use 10 g of base and 2 g of activator.
	<ol>
		<li>De-gas mixture in a vacuum desiccator for 25 - 45 min until all bubbles are gone.</li>
		<li>Fill the cavity of each of the 3D printed molds with approximately 0.75 mL of PDMS using a 10mL syringe until the mold slightly overflows with a meniscus (Figure 2, Step 2.1).</li>
		<li>De-gas the filled molds for 45 min to remove additional bubbles that may have formed when filling.</li>
	</ol>
	</li>
	<li>Apply a glass cover disc on top of the PDMS-filled mold, pressing the disc down at an angle to prevent bubbles from being trapped (Figure 2, Step 2.2). Allow excess PDMS to be expelled as the glass disc cover is applied.</li>
	<li>Use small ratchet clamp to hold the cover disc tightly to the mold. 
	NOTE: This creates a flat surface once the PDMS is cured and produces through holes where the 3D printed geometries are flush with the glass cover slip. Alternatively, to increase the number of devices that can be cured at one time, build a multi-clamp device (e.g. Figure 2, Step 2.3).</li>
	<li>Cure the PDMS device in the clamped molds at 100 oC in an oven for 90 min (Figure 2, Step 2.4). Remove the molds from oven and allow to cool until they can be easily handled. If the clamping device is metal, remove the mold and cover disc assembly from the clamp to prevent the metal from continuing to cure the PDMS due to residual heat. 
	NOTE: The device is easiest to remove from the mold while still slightly warm and not fully cured. However, once the mold is removed from the oven and the cover disc is removed, the device can sit for a couple of days before proceeding to demolding. If the device is demolded shortly after removing from the curing oven, place it in a covered petri dish to minimize contaminants while allowing it to fully cure.</li>
</ol>
3. Plasma-bonding zWEDGI to Glass Dish
<ol>
	<li>Clamp the mold containing the PDMS device in a bench vice so that the mold&#39;s geometries are facing up, parallel to the working station bench.
	<ol>
		<li>Start to remove the PDMS device by releasing the PDMS pull tab from the mold using flat-tipped tweezers. Work around the perimeter of the mold with the tweezers (like removing a cake out of a pan (Figure 2, Step 3.1)).</li>
		<li>Use filtered compressed air and tweezers to gently pull the device out of the mold by holding onto the pull tab and blowing air under the device. Work slowly, allowing the air to help separate the PDMS from the mold, taking special care with the tips of the tweezers around the thin tunnel sections.</li>
	</ol>
	</li>
	<li>Place the PDMS device upside down onto the inside of the cover of a glass bottomed dish so that the restraining tunnel wedges are touching the plastic (Figure 2, Step 3.2).</li>
	<li>Place the dish cover with upside-down zWEDGI and the corresponding glass bottomed dish into a plasma cleaner with the inner glass facing upward (Figure 2, Step 3.3).
	<ol>
		<li>Evacuate the plasma cleaning chamber until the pressure reaches 500 mTorr.</li>
		<li>Set radio frequency (RF) power to high. Expose the device and the dish to RF frequency for approximately 2 min. Slowly de-pressurize the chamber and return the device and the dish to a clean room hood.</li>
	</ol>
	</li>
	<li>Remove the PDMS device from dish cover. Flip over the PDMS zWEDGI to the correct orientation on the glass by positioning it carefully onto the center well of the glass bottom dish (Figure 2, Step 3.4)
	<ol>
		<li>Using the back end of the tweezers, lightly press down on the PDMS device to ensure air bubbles are not trapped beneath. Apply pressure around the minute geometries of the micro channels and smooth the PDMS out to the edges (Figure 5C). 
		NOTE: Complete contact between the PDMS and the glass ensures better adherence from plasma bonding. The zWEDGI device can be plasma bonded onto other glass bottomed dish formats, such as a glass bottomed 6-well plate (Figure 2C).</li>
		<li>Place a cover over the device dish before removing from the clean room hood. 
		NOTE: Once the devices have been fixed to glass, the protocol can be paused until they are ready for use.</li>
	</ol>
	</li>
</ol>
4. Channel Preparation and Loading Larvae
Note: General zebrafish husbandry was conducted per The Zebrafish Book, available online at http://zfin.org/zf_info/zfbook/zfbk.html. Adult zebrafish and embryos were maintained as described previously1. The wild type AB strain was used. Follow the institution&#8217;s Animal Care Protocol for specifics regarding requirements for imaging live larvae.
<ol>
	<li>Rinse the channels with a minimum of&#160;100 &#181;L 70% ethanol per channel, using a micropipette to rinse through the restraining tunnel. Remove ethanol and rinse 2 or 3 times with double distilled water. Allow to air dry.</li>
	<li>Fill the channels with skim milk (at a concentration of 1% diluted in water) for 10 min at room temperature to minimize adherence of larvae to the glass coverslip of the dish. Then, gently submerge the device several times in double distilled water to rinse. Allow to air dry upside down. 
	NOTE: Protocol can be paused here. This preparation of zWEDGI device can be done several days prior to use. Store covered at room temperature.</li>
	<li>Prepare 1% low melting point (LMP) agarose by combining 100 &#181;L 2% melted LMP agarose with 100 &#181;L 2x&#160;Tricaine (0.4 mg/mL Tricaine - ethyl 3-aminobenzoate) in E3 buffer11 pre-warmed to 38 oC. Maintain the 1% agarose/tricaine solution at 38 oC in a hot block to prevent gelling. 
	NOTE: For multiphoton microscopy 11, use either un-pigmented zebrafish variants (such as casper12) or maintain larvae in E3 containing 0.2 mM N-phenylthiourea to prevent pigment formation11to minimize absorption of the near infrared wavelengths by the pigment. 
	Caution: N-phenylthiourea is toxic. Follow the institution&#39;s rules for disposal.</li>
	<li>Anesthetize larvae in E3 buffer containing 0.2 mg/mL tricaine (Tricaine/E3)11. Wait until larvae are motionless and non-responsive to a touch stimulus.</li>
	<li>Pre-wet the channels with a few microliters of Tricaine/E3.
	<ol>
		<li>Pick up a single larva using a wide orifice pipette tip. Deposit the larvae into the loading channel (Figure 3A). Using a pipette tip or similar tool, orient the larva in the loading chamber such that the dorsal side faces the straighter edge of the loading chamber and the tail faces toward the restraining tunnel (Figure 3B).</li>
		<li>Carefully withdraw fluid from the wounding chamber, allowing the larva to flow into the restraining tunnel (Figure 3C). Remove most of the liquid while maintaining moisture around the larva (Figure 3D). This process can be assisted by tilting the dish slightly toward the wounding chamber.</li>
	</ol>
	</li>
	<li>Place 1% LMP agarose in Tricaine/E3 (~38 oC) over the larva&#39;s head, filling the loading chamber (Figure 3E). Allow agarose to solidify with the larva in the proper position. Add Tricaine/E3 to the wounding chamber as needed to maintain hydration. Repeat this loading process for the remaining 2 channels.</li>
	<li>Using a syringe needle, carefully remove any agarose that seeped through the restraining tunnel into the wounding chamber (Figure 3F). Add additional tricaine/E3 either just over the agarose (for wounding, short term imaging, or wound treatment isolation) or to fill the culture dish (for long-term imaging). Replace the culture dish lid to prevent evaporation. Larvae can be imaged at this point (unwounded) or wounded.</li>
</ol>
5. Wounding and Imaging Larvae
<ol>
	<li>Using a sterile scalpel blade, transect the tail fin posterior to the notochord11 in the wounding chamber (Figure 3G, H). Add additional tricaine/E3 if needed and replace the culture dish lid. 
	NOTE: Alternatively, depending of the developmental time window of interest, larvae can be wounded, permitted to recover in E3 and maintained in an incubator (28.5 oC) until the desired imaging time, at which point they can be loaded into channels as described above.</li>
	<li>Install the zWEDGI device with anesthetized larvae onto an inverted microscope in a stage insert that will accommodate the 60 mm glass bottom dish. Locate the tail of the larva in the upper-most channel, rotating the dish as needed to get the tail in the desired position. Image as required for the specific experiment. 
	Note: The zWEDGI is broadly applicable for high-resolution light microscopy, including widefield fluorescence and laser scanning microscopy. There are a number of parameter considerations when imaging zebrafish larvae, but specific imaging parameters are instrument, sample and experiment dependent. Here, the larval tail was imaged on a custom build multiphoton microscope5,11 utilizing the following parameters: 40X long working distance water immersion objective, 890 nm laser excitation, 445/20 nm emission filter and 512 x 512 resolution.</li>
</ol>
6. End of Experiment
<ol>
	<li>Remove the zWEDGI dish from microscope. Euthanize the larvae by placing the zWEDGI either on an ice water bath or at 4 oC for at least 20 min and assess for absence of heartbeat and circulation. 
	NOTE: Because the larvae are maintained in separate compartments, the larvae can be individually recovered by gently pulling on the agar with forceps or pipette. The agar can be removed from the head region and the larva used for additional procedures, such as fixation for antibody staining or processed for RNA or protein extraction.</li>
	<li>Following removal of the larvae and agar, clean the zWEDGI with ethanol and distilled water, as described in step 4.1 and set upside down to air dry. Store covered in a cool, dry location. Re-coat with skim milk (step 4.2) as needed prior to next use. 
	NOTE: zWEDGIs can be re-used multiple times, until the PDMS begins to come away from the glass.</li>
</ol>

<ol>
	<li>Yoo, S. K., Freisinger, C. M., LeBert, D. C., Huttenlocher, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+redox,+Src+family+kinase,+and+calcium+signaling+integrate+wound+responses+and+tissue+regeneration+in+zebrafish.">Early redox, Src family kinase, and calcium signaling integrate wound responses and tissue regeneration in zebrafish.</a> J. Cell Biology. 199 (2), 225-234 (2012).</li><li>Kawakami, A., Fukazawa, T., Takeda, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+fin+primordia+of+zebrafish+larvae+regenerate+by+a+similar+growth+control+mechanism+with+adult+regeneration.">Early fin primordia of zebrafish larvae regenerate by a similar growth control mechanism with adult regeneration.</a> Dev. Dynam. 231 (4), 693-699 (2004).</li><li>Konantz, J., Antos, C. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reverse+genetic+morpholino+approach+using+cardiac+ventricular+injection+to+transfect+multiple+difficult-to-target+tissues+in+the+zebrafish+larva.">Reverse genetic morpholino approach using cardiac ventricular injection to transfect multiple difficult-to-target tissues in the zebrafish larva.</a> JoVE. (88), (2014).</li><li>Hall, C., Flores, M. F., Kamei, M., Crosier, K., Crosier, P., Sampath, K., Roy, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+Imaging+Innate+Immune+Cell+Behavior+During+Normal+Development,+Wound+Healing+and+Infection.">Live Imaging Innate Immune Cell Behavior During Normal Development, Wound Healing and Infection.</a> Live Imaging in Zebrafish: Insights into Development and Disease. , (2010).</li><li>Huemer, K., Squirrell, J. M., Swader, R., LeBert, D. C., Huttenlocher, A., Eliceiri, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=zWEDGI:+Wounding+and+Entrapment+Device+for+Imaging+Live+Zebrafish+Larvae.">zWEDGI: Wounding and Entrapment Device for Imaging Live Zebrafish Larvae.</a> Zebrafish. , (2016).</li><li>Lisse, T. S., Brochu, E. A., Rieger, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Capturing+tissue+repair+in+zebrafish+larvae+with+time-lapse+brightfield+stereomicroscopy.">Capturing tissue repair in zebrafish larvae with time-lapse brightfield stereomicroscopy.</a> JoVE. (95), (2015).</li><li>Kamei, M., Isogai, S., Pan, W., Weinstein, B. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+blood+vessels+in+the+zebrafish.">Imaging blood vessels in the zebrafish.</a> Methods Cell Biol. 100, 27-54 (2010).</li><li>Graeden, E., Sive, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+the+zebrafish+embryonic+brain+by+confocal+microscopy.">Live imaging of the zebrafish embryonic brain by confocal microscopy.</a> JoVE. (26), (2009).</li><li>Petzold, A. M., Bedell, V. 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=SCORE+imaging:+specimen+in+a+corrected+optical+rotational+enclosure.">SCORE imaging: specimen in a corrected optical rotational enclosure.</a> Zebrafish. 7 (2), 149-154 (2010).</li><li>Macdonald, N. P., Zhu, F., 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=Assessment+of+biocompatibility+of+3D+printed+photopolymers+using+zebrafish+embryo+toxicity+assays.">Assessment of biocompatibility of 3D printed photopolymers using zebrafish embryo toxicity assays.</a> Lab Chip. 16 (2), 291-297 (2016).</li><li>LeBert, D. C., Squirrell, J. M., Huttenlocher, A., Eliceiri, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Second+harmonic+generation+microscopy+in+zebrafish.">Second harmonic generation microscopy in zebrafish.</a> Methods Cell Biol. 133, 55-68 (2016).</li><li>White, R. M., Sessa, 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=Transparent+Adult+Zebrafish+as+a+Tool+for+In+Vivo+Transplantation+Analysis.">Transparent Adult Zebrafish as a Tool for In Vivo Transplantation Analysis.</a> Cell Stem Cell. 2 (2), 183-189 (2008).</li><li>LeBert, D. C., Squirrell, J. 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=Matrix+metalloproteinase+9+modulates+collagen+matrices+and+wound+repair.">Matrix metalloproteinase 9 modulates collagen matrices and wound repair.</a> Development. 142 (12), 2136-2146 (2015).</li><li>Campagnola, P. J., Millard, A. C., Terasaki, M., Hoppe, P. E., Malone, C. J., Mohler, W. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+high-resolution+second-harmonic+generation+imaging+of+endogenous+structural+proteins+in+biological+tissues.">Three-dimensional high-resolution second-harmonic generation imaging of endogenous structural proteins in biological tissues.</a> Biophys. J. 82 (1 Pt 1), 493-508 (2002).</li><li>Schindelin, J., Arganda-Carreras, I., 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=Fiji:+an+open-source+platform+for+biological-image+analysis.">Fiji: an open-source platform for biological-image analysis.</a> Nat. Methods. 9 (7), 676-682 (2012).</li></ol>

The authors have nothing to disclose.

The purpose of the zWEDGI device is to capture 3D time lapse imaging by stabilizing and orienting the fish within the small working distance of a high-resolution microscope objective. While meeting these design specifications, it is also an improvement over traditional agar-based preparation for live imaging. There are three critical steps (below) in the fabrication of the zWEDGI, which, if not done correctly, can result in defective devices:
PDMS preparation (Figure 5A</s...

The regenerative capacity of zebrafish larvae Danio rerio make them an ideal model organism for examining wound response as well as healing and regrowth1,2,3,4. Access to an array of transgenic zebrafish lines and zebrafish&#39;s anatomical transparency further enhance their utility for in vivo studies of wound response events as well as longer-term regenerative processes4. Study of these biological processes using high-resolution time-lapse fluorescence microscopy therefore demands a live imaging zebrafish device that allows for high stability and minimal movement of the zebrafish larva while maintaining viability. It is key that the device allows for effective wounding while healing and regeneration occur unaffected by the device.
The standard live imaging stabilization method of embedding the larva in agarose during live imaging restricts growth and wound regeneration5 and may increase death rates since larvae begin to show sign of cardiac stress and tissue necrosis after four hours4. Therefore, removal of agarose from regions of interest is often necessary to allow normal development and regeneration6, exposing the larvae to potential damage as the agarose is cut away. Furthermore, with the agarose embedding technique, the user must orient the larvae in the short time before the agarose solidifies5,6,7. Rapidly manipulating the larva not only requires skill of the user, it also risks damage to the larva. Although methods to stabilize the larva for live imaging have been described to circumvent these drawbacks, such as ridged agar wells3 or divets8, the use of silicone vacuum grease to create an imaging chamber with PVC piping or other materials6, and rotational tubing9, many of these methods are labor intensive, messy, often non-reusable and don&#39;t allow for environmental manipulation (drug treatments, wounding etc.) after the fish has been mounted.
Therefore, the zWEDGI device (Figure 1) was designed to overcome some of the drawbacks of agar mounting for long-term live imaging of zebrafish larvae while permitting manipulation of the specimen. The zWEDGI consists of three semi-open compartmentalized chambers (Figure 1A) to allow for loading, restraint, wounding and imaging of 2 to 4 days post-fertilization zebrafish larvae. The device is fabricated from Polydimethylsiloxane (PDMS) and placed onto the cover slip of a 60 mm glass bottom imaging dish. The design presented here was intended for wound healing studies, however the use of a modular design and standard fabrication technologies make the zWEDGI design modifiable and amenable to a variety of experimental procedures, especially for procedures that require minimal restraint with experimental manipulation and long-term imaging.

<table><tbody><tr><td>&lt;strong&gt;Fabricate molds&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Solidworks Professional Accedemic Research 3D modeling software</td><td>Dassault Systemes</td><td>SPX0117-01</td><td>Fisher Unitech</td></tr><tr><td>Viper Si2 SLA 3D printer</td><td>3D Systems Inc.</td><td>23200-902</td><td>3D Systems Inc.</td></tr><tr><td>Accura 60 photopolymer resin</td><td>3D Systems Inc.</td><td>24075-902</td><td>3D Systems Inc.</td></tr><tr><td>denatured alcohol</td><td>Sunnyside</td><td>5613735</td><td>Menards</td></tr><tr><td>UV post cure apparatus</td><td>3D Systems Inc.</td><td>23363-101-00</td><td>3D Systems Inc.</td></tr><tr><td>TouchNTuff nitrile gloves</td><td>Ansell</td><td>92-600</td><td>McMaster Carr</td></tr><tr><td>220B, 400B, 600 grit T414 blue-bak sandpaper&amp;nbsp;</td><td>Norton</td><td>66261139359, 54, 52</td><td>MSC</td></tr><tr><td>borosilicate glass disc, 2&quot; diameter</td><td>McMaster-Carr</td><td>MIL-G-47033</td><td>McMaster-Carr</td></tr><tr><td>ultrasonicator cleaner</td><td>Branson</td><td>1510R-MTH</td><td></td></tr><tr><td>isopropyl rubbing alcohol 70%</td><td>Hydrox</td><td>54845T43</td><td>McMaster-Carr</td></tr><tr><td>10oz clear plastic cup</td><td>WNA Masterpiece</td><td>557405</td><td>Amazon</td></tr><tr><td>6&quot;craft stick</td><td>Perfect Stix</td><td>Craft WTD-500</td><td>Amazon</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Fabricate zWEDGI PDMS device&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Sylgard 184 silicon elastomeric kit&amp;nbsp;</td><td>Dow-Corning</td><td>4019862</td><td>Ellworth Adhesives&amp;nbsp;</td></tr><tr><td>10mL syringe</td><td>Becton Dickinson</td><td>305219</td><td>Vitality Medical Inc</td></tr><tr><td>desiccator</td><td>Bel-Art Scienceware</td><td>F42027-0000</td><td>Amazon</td></tr><tr><td>4 in ratcheting bar clamp</td><td>Pittsburgh</td><td>68974</td><td>Harbor Freight</td></tr><tr><td>lab oven</td><td>Quincy Lab Inc.</td><td>20GC</td><td>Global Industrial</td></tr><tr><td>tweezer set</td><td>Aven</td><td>549825</td><td>McMaster-Carr</td></tr><tr><td>compressed air filtered nozzle</td><td>Innotech</td><td>TA-N2-2000FT</td><td>Cleanroom Supply</td></tr><tr><td>vacuum bench vise</td><td>Wilton Tool Group</td><td>63500</td><td>MSC Industrial</td></tr><tr><td>55mm glass bottom dish; 30mm micro-well #1.5 cover glass</td><td>Cellvis</td><td>D60-30-1.5-N</td><td>Cellvis</td></tr><tr><td>plasma cleaner</td><td>Harrick Plasma</td><td>PDC-001</td><td>Harrick Plasma</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Loading Larvae&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Pipetteman, P200</td><td>Gilson</td><td>F123601</td><td></td></tr><tr><td>100% ethanol (diluted to 70% with water prior to use)</td><td>Pharmco-aaper</td><td>111000200</td><td></td></tr><tr><td>Transfer pipette</td><td>Fisherbrand</td><td>13-711-5A</td><td>Fisher Scientific</td></tr><tr><td>powdered skim milk</td><td></td><td>2902887</td><td>MP Biomedicals</td></tr><tr><td>double distilled water</td><td></td><td></td><td></td></tr><tr><td>N-phenylthiorurea</td><td>Sigma-Aldrich</td><td>P7629</td><td>Sigma-Aldrich</td></tr><tr><td>tricaine (ethyl 3-aminobenzoate)</td><td></td><td>C-FINQ-UE</td><td>Western Chemical</td></tr><tr><td>low melting point agarose</td><td>Sigma-Aldrich</td><td>A0701</td><td>Sigma-Aldrich</td></tr><tr><td>heat block (dry bath incubator)</td><td>Fisher Scientific</td><td>11-718-2</td><td>Fisher Scientific</td></tr><tr><td>E3 buffer&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>large orifice pipette tip, 200 uL</td><td>Fisherbrand</td><td>02-707-134</td><td>Fisher Scientific</td></tr><tr><td>General purpose pipette tip, 200 uL</td><td>Fisherbrand</td><td>21-197-8E</td><td>Fisher Scientific</td></tr><tr><td>#15 scalpel blade&amp;nbsp;</td><td>Feather</td><td>2976</td><td>Amazon</td></tr><tr><td>25G syringe needle</td><td>BD&amp;nbsp;</td><td>BD305122</td><td>Fisher Scientific</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Imaging&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>inverted microscope</td><td></td><td></td><td></td></tr><tr><td>Imaris imaging software</td><td>Bitplane</td><td></td><td></td></tr></tbody></table>

long-term live imaging device for improved experimental manipulation of zebrafish larvae

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for live high-resolution microscopic imaging of dynamic biological phenomena in space and time with cellular resolution. However, the traditional method of agarose encapsulation for live imaging can impede larval development and tissue regrowth. Therefore, this manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which was designed and fabricated as a functionally compartmentalized device to orient larvae for high-resolution microscopy while permitting caudal fin transection within the device and subsequent unrestrained tail development and re-growth. This device allows for wounding and long-term imaging while maintaining viability. Given that the zWEDGI mold is 3D printed, the customizability of its geometries make it easily modified for diverse zebrafish imaging applications. Furthermore, the zWEDGI offers numerous benefits, such as access to the larva during experimentation for wounding or for the application of reagents, paralleled orientation of multiple larvae for streamlined imaging, and reusability of the device.

The zWEDGI PDMS microfluidic device is a functionally compartmentalized device designed to accommodate four main functions (listed below) associated with live imaging of caudal fin wounding healing and regrowth in the zebrafish larvae. PDMS was chosen for zWEDGI fabrication because it is not only readily available and an industry standard for biocompatibility, but also works well in molds. Additionally, PDMS makes the device reusable and void of hard or sharp edges once the device is form...

Watch this Scientific Journal Video about Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae at JoVE.com

Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae

This manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which is a compartmentalized device designed to orient and restrain zebrafish larvae. The design permits tail transection and long-term collection of high-resolution fluorescent microscopy images of wound healing and regeneration.

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for ...

long-term-live-imaging-device-for-improved-experimental-manipulation

Research

JoVE Journal

Bioengineering

8.7K Views.  University of Wisconsin-Madison. Dit manuscript beschrijft de zWEDGI (zebrafish Wounding en Entrapment apparaat voor groei en Imaging), die is een verzuilde apparaat ontworpen bedwingen zebrafish larven te oriënteren. Het ontwerp staat staart transect en op lange termijn collectie van hoge-resolutie fluorescentie microscopie beelden van wondgenezing en regeneratie.

Video: Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae

Live Imaging of Innate Immune and Preneoplastic Cell Interactions Using an Inducible Gal4/UAS Expression System in Larval Zebrafish Skin

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS expression system1 in zebrafish larvae, and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells. The KalTA4-ERT2/UAS system is both inducible and reversible which allows us to induce cell transformation with precise temporal/spatial resolution in vivo. This provides us with a unique opportunity to live image how individual preneoplastic cells interact with host tissues as soon as they emerge, then follow their progression as well as regression. Recent studies in zebrafish larvae have shown a trophic function of innate immunity in the earliest stages of tumorigenesis2,3. Our inducible system would allow us to live image the onset of cellular transformation and the subsequent host response, which may lead to important insights on the underlying mechanisms for the regulation of oncogenic trophic inflammatory responses. We also discuss how one might adapt our protocol to achieve temporal and spatial control of ectopic gene expression in any tissue of interest.

Studying the earliest events of preneoplastic cell progression and innate immune cell interaction is pivotal to understand and treat cancer. Here we describe a method to conditionally induce epithelial cell transformations and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells in zebrafish larvae.

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS ...

Developmental Biology

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. ...

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

Biology

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning fundamental developmental biology through to large-scale genetic and chemical screens. However, many experiments, especially those related to infection and xenograft models, rely on microinjection and imaging of embryos and larvae, which are laborious techniques that require skill and expertise. To improve the precision and throughput of current microinjection techniques, we developed a series of microstructured devices to orient and stabilize zebrafish embryos at 2 days post fertilization (dpf) in ventral, dorsal, or lateral orientation prior to the procedure. To aid in the imaging of embryos, we also designed a simple device with channels that orient 4 zebrafish laterally in parallel against a glass cover slip. Together, the tools that we present here demonstrate the effectiveness of photolithographic approaches to generate useful devices for the optimization of zebrafish techniques.

Microinjection of zebrafish embryos and larvae is a crucial but challenging technique used in many zebrafish models. Here, we present a range of microscale tools to aid in the stabilization and orientation of zebrafish for both microinjection and imaging.

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning ...

Automated High-throughput Behavioral Analyses in Zebrafish Larvae

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system measures spontaneous behaviors and the response to an aversive stimulus, which is shown to the larvae via a PowerPoint presentation. The recorded images are analyzed with an ImageJ macro, which automatically splits the color channels, subtracts the background, and applies a threshold to identify individual larvae placement in the lanes. We can then import the coordinates into an Excel sheet to quantify swim speed, preference for edge or side of the lane, resting behavior, thigmotaxis, distance between larvae, and avoidance behavior. Subtle changes in behavior are easily detected using our system, making it useful for behavioral analyses after exposure to environmental toxicants or pharmaceuticals.

Our laboratory developed a novel high-throughput automated imaging system that is useful for the detection of several different behaviors in 7-day-old zebrafish larvae. The system can be used for detecting subtle changes in behavior after the larvae have been exposed to environmental toxicants or pharmaceuticals.

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system ...

Neuroscience

A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues or cells. A difficulty with imaging whole embryo development is that zebrafish embryos grow substantially in length. When mounted as regularly done in 0.3-1% low melt agarose, the agarose imposes growth restriction, leading to distortions in the soft embryo body. Yet, to perform confocal time-lapse microscopy, the embryo must be immobilized. This article describes a layered mounting method for zebrafish embryos that restrict the motility of the embryos while allowing for the unrestricted growth. The mounting is performed in layers of agarose at different concentrations. To demonstrate the usability of this method, whole embryo vascular, neuronal and muscle development was imaged in transgenic fish for 55 consecutive hours. This mounting method can be used for easy, low-cost imaging of whole zebrafish embryos using inverted microscopes without requirements of molds or special equipment.

This article describes a method to mount fragile zebrafish embryos for extended time-lapse confocal microscopy. This low-cost method is easy to perform using regular glass-bottom microscopy dishes for imaging on any inverted microscope. The mounting is performed in layers of agarose at different concentrations.

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues ...

Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish with fluorescently-marked plasma membranes can be scanned in a confocal laser scanning microscope in order to determine the volume of muscle tissue and the number of muscle fibers present. Efficient approaches for the measurement of cell number and size in live animals over time are described and validated against more arduous segmentation methods. Methods are described that permit the control of muscle electrical, and thus contractile, activity. Loss of skeletal muscle contractile activity greatly reduced muscle growth. In larvae, a protocol is described that allows reintroduction of patterned electrical-evoked contractile activity. The described methods minimize the effect of inter-individual variability and will permit analysis of the effect of electrical, genetic, drug, or environmental stimuli on a variety of cellular and physiological growth parameters in the context of the living organism. Long-term follow-up of the measured effects of a defined early-life intervention on individuals can subsequently be performed.

Optical clarity is a major advantage for cell biological and physiological work in zebrafish. Robust methods for measurement of cell growth in individual animals are described that permit novel insights into how growth of skeletal muscle and neighboring tissues are integrated with whole body growth.

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish ...

Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function. Our studies have found that dying cells send signals to their live neighbors to form and contract a ring of actin and myosin that ejects it out from the epithelial sheet while closing any gaps that might have resulted from its exit, a process termed cell extrusion1. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe a method to induce and image extrusion in the larval zebrafish epidermis. To visualize extrusion, we inject a red fluorescent protein labeled probe for F-actin into one-cell stage transgenic zebrafish embryos expressing green fluorescent protein in the epidermis and induce apoptosis by addition of G418 to larvae. We then use time-lapse imaging on a spinning disc confocal microscope to observe actin dynamics and epithelial cell behaviors during the process of apoptotic cell extrusion. This approach allows us to investigate the extrusion process in live epithelia and will provide an avenue to study disease states caused by the failure to eliminate apoptotic cells.

Dying cells are extruded from epithelial tissues by concerted contraction of neighboring cells without disrupting barrier function. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe methods to induce and image extrusion in the larval zebrafish epidermis at cellular resolution.

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function.  Our studies have found ...

Analysis of Zebrafish Cardiac Contraction with Virtual Reality Interaction

4D Light-sheet Imaging of Zebrafish Cardiac Contraction

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for gaining insights into structural and functional changes in response to injuries. However, obtaining high-resolution and high-speed 4-dimensional (4D, 3D spatial + 1D temporal) images of the zebrafish heart to assess cardiac architecture and contractility remains challenging. In this context, an in-house light-sheet microscope (LSM) and customized computational analysis are&#160;used to overcome these technical limitations. This strategy, involving LSM system construction, retrospective synchronization, single cell tracking, and user-directed analysis, enables one to investigate the micro-structure and contractile function across the entire heart at the single-cell resolution in the transgenic Tg(myl7:nucGFP) zebrafish larvae. Additionally, we are able to further incorporate microinjection of small molecule compounds to induce cardiac injury in a precise and controlled manner. Overall, this framework allows one to track physiological and pathophysiological changes, as well as the regional mechanics at the single-cell level during cardiac morphogenesis and regeneration.

Author Spotlight: High-Resolution 4D Light-Sheet Imaging and Virtual Reality in Zebrafish for Single-Cell Analysis of Heart Function

This protocol utilizes light-sheet imaging to investigate cardiac contractile function in zebrafish larvae and gain insights into cardiac mechanics through cell tracking and interactive analysis.

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for ...

In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at cellular and subcellular resolution. Advances in molecular and imaging technologies have allowed us to gain numerous detailed insights into cellular and molecular mechanisms of brain development in the transparent zebrafish embryo. Recently, processes of refinement of neuronal connectivity that occur at later larval stages several weeks after fertilization, which are for example control of social behavior, decision making or motivation-driven behavior, have moved into focus of research. At these stages, pigmentation of the zebrafish skin interferes with light penetration into brain tissue, and solutions for embryonic stages, e.g., pharmacological inhibition of pigmentation, are not feasible anymore.
Therefore, a minimally invasive surgical solution for microscopy access to the brain of awake zebrafish is provided that is derived from electrophysiological&#160;approaches. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, exposing underlying neurons and axonal tracts without damage. This allows for recording neuronal morphology, including synaptic structures and their molecular contents, and the observation of physiological changes such as Ca2+ transients or intracellular transport events. In addition, interrogation of these processes by means of pharmacological inhibition or optogenetic manipulation is feasible. This brain exposure approach provides information about structural and physiological changes in neurons as well as the correlation and interdependence of these events in live brain tissue in the range of minutes or hours. The technique is suitable for in vivo brain imaging of zebrafish larvae up to 30 days post fertilization, the latest developmental stage tested so far. It, thus, provides access to such important questions as synaptic refinement and scaling, axonal and dendritic transport, synaptic targeting of cytoskeletal cargo or local activity-dependent expression. Therefore, a broad use for this mounting and imaging approach can be anticipated.

Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at ...

Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae

Samenvatting

Meer video's verkennen