We thank the MDI Biological Laboratory animal core service facility for zebrafish maintenance. Research reported in this publication was supported by Institutional Development Awards (IDeA) from the National Institute of General Medical Sciences of the National Institutes of Health under grant numbers P20GM104318 (for COBRE) and P20GM103423 (INBRE) and Department of Defense &#8211; USAMRAA (W81XWH-BAA-1) grant.

Zebrafish (nacre strain) were bred and raised according to established protocols. All efforts were made to minimize suffering, using 0.4 mM Tricaine for anesthesia and 1 mM Tricaine for euthanasia. Zebrafish embryos and larvae were handled in strict accordance with good animal practice as approved by the appropriate committee (MDI Biological Laboratory animal core IACUC number 13-20). This study was approved by the National Human Genome Research Institute Animal Care and Use Committee, MDIBL Institutional Assurance # A-3562-01 under protocol # 14-09.
Note: The imaging procedure that captures fin regeneration in larval zebrafish is summarized in the following steps:
1. Raising of Zebrafish to Larval Stages
<ol>
	<li>Collect the eggs and place approximately 50 eggs into a 100 x 25 mm Petri dish containing 0.03% Instant Ocean salt in deionized water supplemented with 0.00004% methylene blue. Incubate O/N in a 28.5 &#176;C incubator.</li>
	<li>The next morning remove the dead embryos with a glass pipette and rinse the eggs in a strainer with 0.03% Instant Ocean salt in deionized water (termed embryo medium). 
	Note: Medium such as Ringers 18, Hanks 18, E2 19, E3 20, and Danieau 21 may be preferred.</li>
	<li>sup&#62;Add fresh embryo medium to the dish. If using pigmented strains, optionally add 0.2 mM 1-phenyl-2-thiourea (PTU), as PTU will prevent melanogenesis and thus pigmentation of the larvae. Let the embryos further develop in the incubator until 2 days post fertilization or any other desired larval stage.</li>
</ol>
2. Preparation of the Imaging Chamber
<ol>
	<li>Method 1: Imaging chambers made from PVC or Teflon tubing (Figure 1) 
	Note: This method is similar to Concha and Adams (1998) 22.
	<ol>
		<li>Acquire drinking-water grade plastic or Teflon tubing from a hardware store with a 25 mm outer and a 20 mm inner diameter. Cut the tubing to make rings of approximately 10 mm thickness with an even surface on each side. Use &#62; 200 grit sandpaper to smoothen the edges.</li>
		<li>Clean the rings with warm water and 70% ethanol and let them air dry.</li>
		<li>With a pipette tip, apply silicon grease to one half of a ring and attach the ring to a 75 mm x 25 mm glass cover slip. Alternatively, use a 3 ml syringe filled with silicon grease instead of a pipette tip. 
		Note: Because it is difficult to insert the silicon grease into the 3 ml syringe, add the silicon grease to a 30 ml syringe first and use this for filling the 3 ml syringe.</li>
	</ol>
	</li>
	<li>Method 2: Preparation of a Petri dish as an imaging chamber.
	<ol>
		<li>Acquire 35 or 60 mm diameter Petri dishes with a glass coverslip attached to the lid (Figure 2A). Alternatively, as shown in Figure 3 of Distel &#38; Koester (2007) 23, drill an opening small enough to hold a coverslip into the lid of a small Petri dish and apply silicon grease to the outside with a 3 ml syringe. Using a clean pipette tip, carefully attach a round or square coverslip of desired thickness to the outside (Figure 2B).</li>
	</ol>
	</li>
	<li>To ensure that the agarose being used for mounting stays firmly attached during the imaging procedure, attach a fine plastic mesh inside the ring. First, cut the mesh made of window screen obtained from a hardware store, into the size of the inner ring diameter using fine scissors with an angle. Then cut a small rectangle &#62; twice the size of the larva into the middle of the mesh (Figure 1, 2).</li>
	<li>Apply four small dots of non-toxic silicon grease to the interface between the cover slip and the chamber ring (Figure 1A).</li>
	<li>Use forceps to attach the mesh firmly to the bottom of the glass coverslip.</li>
</ol>
3. Mounting and Imaging of the Pre-injured Larva (this Step is Optional)
Note: This step is suitable for comparisons between the amputated and regenerated fin length, as the amputation plane after fin regeneration is not recognizable in zebrafish larvae.
<ol>
	<li>Prepare a 0.5-1.2% low-melt agarose solution in embryo medium for immobilization of the specimen.</li>
	<li>Heat the agarose in a microwave and transfer the liquid agarose into 1.5 ml tubes that are pre-warmed to 42 &#176;C in a heating block.</li>
	<li>Let the hot agarose cool down to 42 &#176;C, which can be maintained over several weeks at this temperature. Try to avoid pipetting the larvae into agarose above 42 &#176;C, as this will be detrimental to the animals.</li>
	<li>Anesthetize several larvae using 10 ml of 0.4 mM Tricaine (pH 7) in embryo medium in a 60 mm diameter Petri dish. Prepare the Tricaine according to the Zebrafish Book Recipes 18.
	<ol>
		<li>Alternatively, use 10 ml of a 1:1000 dilution of 99% 2-Phenoxyethanol in a 60 mm diameter Petri dish. Poke the larvae with a capped microloader pipette tip to assess their touch response before proceeding. Use only non-responsive larvae.</li>
	</ol>
	</li>
	<li>Transfer one larva into the 42 &#176;C agarose solution using a glass Pasteur pipet. Do not transfer excessive liquid into the agarose, otherwise the agarose will be too diluted and not solidify.</li>
	<li>Discard the remaining liquid from the pipet and transfer the larva with one drop of agarose into a small Petri dish (35 or 60 mm diameter). Position the larva on its side for imaging of the tail fin.</li>
	<li>Allow the agarose to solidify. Assess the agarose with a plastic pipette tip; the tip will immerse into the agarose if it is too liquid. Once the agarose has solidified, a small indentation will be visible upon touching with a pipette tip. Following solidification, add Tricaine solution and proceed to the stereomicroscope that will be used later for time-lapse imaging. 
	Note: This step may also be performed by placing the anesthetized larva on a Petri dish coated with 1.5% agarose in embryo medium.</li>
	<li>Use a stereomicroscope with time-lapse software. Select an appropriate objective and magnification, which will be used later for time-lapse imaging. Here, use a 3.5 x, 16 mm working distance objective lens on a stereomicroscope. As desired, utilize alternative microscopes and objective lenses but choose a proper magnification to account for potential xy-drift and growth of the fin during the imaging procedure.</li>
	<li>Select the camera detection mode on the microscope.</li>
	<li>In the software, select the &#39;live&#39; mode to view the larva on the screen.</li>
	<li>Open the &#39;Properties&#39; window to automatically detect the brightness.</li>
	<li>Manually adjust the contrast at the microscope trans-illumination base.</li>
	<li>Move the larva out of the field of view and select the Shading Correction feature to minimize background noise.</li>
	<li>Position the larva back into the field of view and take a snapshot. Save the image.</li>
	<li>Remove the agarose from the larva by first scraping the agarose off the head. This way the larva can be slipped out of the agarose by gently pulling the head away from the remaining agarose with a capped microloader pipet tip or an insect pin.</li>
	<li>With a glass Pasteur pipet transfer the larva into fresh Tricaine solution.</li>
</ol>
4. Amputation Assay
<ol>
	<li>Prepare a 1.5% agarose solution using embryo medium and pour a thin layer into a Petri dish. Let the agarose solidify.</li>
	<li>Under a stereomicroscope, place the larva sideward onto the solidified agarose and amputate the tail fin with a 23 G&#160;syringe needle with slight pressure (Figure 3A).</li>
</ol>
5. Mounting the Larva for Time-lapse Imaging
<ol>
	<li>Proceed as described in steps 3.1 &#8211; 3.5 (Figure 3B).</li>
	<li>Transfer a drop of 0.5-1.2% liquid agarose at 42 &#176;C containing the larva into the imaging chamber ring (step 2.1), orient the larva and let the agarose solidify. Fill the ring with Tricaine solution. Alternatively, if the Petri dish chamber (step 2.2) is utilized, mount the larva onto the lid coverslip and fill the lid with Tricaine solution.</li>
	<li>To allow proper wound healing or tissue regeneration to occur, carefully scrape off the agarose surrounding the distal tail fin using a capped microloader pipette tip or an insect pin. Try not to injure the fin repeatedly (Figure 3C).</li>
	<li>Decant the Tricaine solution containing the removed agarose and fill the chamber ring with fresh Tricaine solution.</li>
	<li>Apply silicon grease to the top of the chamber ring and attach a 75 mm x 25 mm glass slide. Try to avoid air pockets in the chamber, as they will interfere with brightfield imaging and desiccate the larva over time.</li>
	<li>If using a Petri dish as an imaging chamber, apply silicon grease to the top rim of the bottom chamber and fill the bottom chamber with Tricaine solution. Carefully decant the Tricaine solution in the lid and turn the lid over to immerse the larva into the Tricaine solution of the bottom chamber at a slight angle to avoid air pockets. The chamber will be sealed due to the silicon grease.</li>
</ol>
6. Time-lapse Imaging
<ol>
	<li>Assemble a heated incubation chamber as described in 23,24 (Figure 4).Place the incubation chamber around the microscope and turn on the heat. Adjust the temperature to 28 &#176;C for about 10 - 20 min or until the temperature has stabilized.</li>
	<li>Open the front of the heated incubation chamber and place the imaging chamber onto the microscope stand with the coverslip facing upward toward the objective.</li>
	<li>Position the larval fin in a way that 2/3 of the field of view remains unoccupied. This ensures the capture of the growth and regeneration of the fin over the course of the imaging procedure without having to reposition the larva.
	<ol>
		<li>Adjust the mounted larva to 28 &#176;C for ~ 30 min prior to starting the time-lapse recording to avoid changes in brightfield intensity or potential shifts of the agarose. Alternatively, utilize pre-warmed buffer to start imaging after shorter adjustment time.</li>
	</ol>
	</li>
	<li>To set up the time-lapse recording, open the 6D Multidimensional Acquisition window in the Axiovision software and select the z-stack and time-lapse option.</li>
	<li>Optional) In the z-stack tab and Slice Mode, select the slice thickness and then select the Start/Stop mode.</li>
	<li>Define the upper and lower position of the stack.</li>
	<li>In the time-lapse tab, select the interval and the duration of the movie and then start the movie by pressing the start button. We found that 30 min intervals are sufficient and this interval does not generate excessive data; however shorter intervals may be used.</li>
	<li>Check the position and z-stack dimensions during the first hour if the larva was not pre-adjusted. If necessary, reposition the larva again after a day, as the larva may have shifted.</li>
	<li>Save the file at the end of the time-lapse recording and proceed with the post-processing and quantifications using available image analysis software, such as Imaris 25 or the open source software packages Image J 26 and Fiji 27.</li>
</ol>
7. Data Analysis
<ol>
	<li>Determining the fin length.
	<ol>
		<li>Open the time-lapse movie in the imaging software and save the files in the proprietary file format to enhance software performance.</li>
		<li>Select the orthogonal view to display individual sections as projected stacks. For drift correction, under the Fiji menu, select Plugins, then Registration, and &#39;Correct 3D drift&#39;. This will open a Fiji window and perform the drift corrections. Correct rotational drift in the Imaris Spots function. Alternatively, install the StackReg and TurboReg plugins in Image J and import to Fiji. Choose the desired transformation algorithm in the StackReg plugin.</li>
		<li>To measure distances (e.g., wound diameter or fin length), select the &#39;Add New Measurement Points&#39; option in the upper left tool bar.
		<ol>
			<li>Under &#39;Configure list of visible Statistics Values&#39; in the bottom left menu, select the statistics values to be displayed.</li>
			<li>In the &#39;Line Mode&#39; under the settings tab select &#39;Pairs (AB, CD ...)&#39;.</li>
			<li>In &#39;Labels Properties&#39; select &#39;Name&#39; and &#39;Distance&#39;, to display the distance between points A and B next to the measurement line.</li>
			<li>Switch to the &#39;Edit&#39; mode and hold down the shift button to select the first point at the end of the notochord. Then use the same configuration to select the second point at the distal fin margin.</li>
			<li>Under the &#39;Statistics&#39; tab, select the disk button (Export All) at the lower right to display and export the distance in the image.</li>
			<li>Repeat the measurements at selected times by moving the slider below the image to the right. Instead of creating new Measurement Points, the previous ones can be repositioned by first selecting the point with the left mouse button, then simultaneously pressing the shift and the left mouse button at the new position.</li>
		</ol>
		</li>
		<li>Alternatively to the Measurement Points option, utilize the slice viewer to measure distances. In the slice view mode, scroll to a desired position and click on the first and second position with the left mouse button. The distance will be displayed. This option however does not allow for data export.</li>
	</ol>
	</li>
	<li>Determining fin length and area in ImageJ.
	<ol>
		<li>Open the time-lapse movie in ImageJ using a plugin that recognizes the .zvi file format. Alternatively, load in a QuickTime file or tiff sequence. 
		Note: If using an uncompressed tiff file format, the image dimensions do not need to be specified.</li>
		<li>If opening a different file format without the file information, select &#39;Set Scale&#39; under the &#39;Analyze&#39; menu to first define the image distance and unit. In the &#39;Set Scale menu&#39;, type in the &#39;Distance in pixels&#39;, below type in the &#39;Known distance&#39; for the pixel value (this can be obtained by measuring the number of pixels on a scale bar that was added to the image, then click Measure to obtain the result), and the &#39;Unit of length&#39; (typically &#39;&#181;m&#39;). Then click OK.</li>
		<li>For fin area measurements select the &#39;Freehand Selection&#39; tool in the tool bar and outline the fin area by holding the left mouse button down while drawing along the outline of the fin. For fin length measurements, select the &#39;Straight&#39; line tool and draw a line between the desired points to be measured.</li>
		<li>Click the &#39;Measure&#39; option under &#39;Analyze&#39; to display the area and length of the fin. Repeat this step as often as necessary for multiple time points of the movie.</li>
	</ol>
	</li>
	<li>Using statistics software the data can be graphically displayed.</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+method+of+PSF+generation+for+3D+brightfield+deconvolution.">A method of PSF generation for 3D brightfield deconvolution.</a> J Microsc. 237, 192-199 (2010).</li><li>Distel, M., Babaryka, A., K&#246;ster, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multicolor+in+vivo+time-lapse+imaging+at+cellular+resolution+by+stereomicroscopy.">Multicolor in vivo time-lapse imaging at cellular resolution by stereomicroscopy.</a> Dev Dyn. 235, 1100-1106 (2006).</li></ol>

The authors have nothing to disclose.

The presented method allows for observing wound healing and tissue regeneration in living zebrafish larvae with in vivo time-lapse imaging on a brightfield stereomicroscope, using a comparatively simple set-up. This procedure requires certain important aspects that we have tested, which will optimize the outcome: 1) Low agarose concentrations (~0.5%) will minimize growth impediments of the continually growing larval zebrafish, 2) Removal of the agarose around the fin is important not to obscure the healing process, 3) Trapping the agarose in a plastic mesh retains the agarose and animal in a stable position throughout the procedure, and 4) A proper temperature-controlled environment, which is essential for larval viability. We have adapted a heated incubation chamber 23,24, which utilizes bubble wrap that is taped onto cardboard, and a wired dome heater to control the temperature and proper air circulation with minimal fluctuations during the imaging procedure. This simple and cost-efficient chamber can be prepared to fit any microscope. A similar heated incubation chamber has been also utilized for imaging mice and chick development 24,29.
We suggest that pre-amputated larvae are mounted for a pre-amputation image, demounted for amputation, and remounted for time-lapse imaging. Although it is feasible to perform these steps in a single step in the final imaging chamber, in our experience we found that amputating the tail fin on a glass coverslip is not optimal, as it tears the tissue and does not result in a clean cut. The agarose-based amputation method using a syringe needle was originally described by Kawakami and colleagues (2004) 16 and is also, in our experience, ideal to perform the amputations. Thus, the rather complicated series of steps that we presented is well justified and ensures an optimal regeneration outcome.
We showed that larval zebrafish at 2 dpf can be imaged up to 1.5 days in agarose and Tricaine solution. We used pH-optimized Tricaine (pH7) solution prepared with Instant Ocean salt, which does not interfere with the specimen&#8217;s health for the presented imaging period. We previously however also demonstrated that using Tricaine in Danieau medium permits time-lapse imaging of 2.5 dpf larval zebrafish on a confocal microscope for at least 2 days 30. Thus, optimal buffer conditions can extend larval health and the length of imaging. Alternatively, lower Tricaine concentrations may be used for anesthesia, or 2-phenoxyethanol, which we found is well tolerated during larval and adult stages at 28 &#176;C for at least 60 hr.
To avoid defects in fin regeneration, we removed the agarose from the tail fin prior to imaging. Our data shows that within 1.5 days the fin has regenerated to about 60%. This regeneration rate is consistent with a previous study defining 3 days as an average time for tail fin regeneration in zebrafish larvae up to 6 dpf 16. Alternative methods to agarose could however be utilized to mount the fish for imaging. For example, thin plasma clots 31 or fluorinated ethylene propylene (FEP) tubes coated with methylcellulose and filled with very low agarose concentrations (0.1%) have been recommended for light sheet microscopy 32 and may be suitable for our presented method. However, we do not recommend methylcellulose and 0.1% agarose, as they require that the specimen are mounted at the bottom of the chamber due to the lack of solidification of these media. Very high concentrations of methylcellulose will moreover generate air pockets based on our experience, and these may interfere with the imaging procedure. If these media are preferred with using the bottom chamber, it is important that an appropriate working distance between the objective lens and the specimen is present. It should be noted that methylcellulose as a mounting medium is recommended only for up to 1 day, as it may interfere with larval health 32.
Mounting the specimen in the lid may result in a slow gravitational downward drift. It is therefore recommended to image multiple sections at each time point, which can either be projected into a single plane or only images that are in the focal plane may be extracted for assembling the final movie. Imaging the specimen at the bottom chamber could be an alternative methodology to avoid potential downward drift. Plasma clots could be useful to avoid drift, as the plasma will stick to the outer enveloping layer (EVL, periderm) 31 and therefore may stabilize the specimen. This however needs to be tested, as well as how long larval zebrafish can be maintained in plasma clots without interfering with larval health or fin regeneration.
Our movie was assembled utilizing individual sections (26 &#181;m) of a recorded z-stack, which covered the full thickness of the fin (~10 &#181;m) and which accounted for potential z-drift of the fin during the imaging procedure. In order to retain 3-D information, it is also possible to project z-stacks into single images. Because this may result in blurriness of the image, brightfield deconvolution may be desired. Software, such as Deconvolve or Autoquant X3 could be utilized for this purpose. Alternatively, mathematical algorithms (described in Tadrous33) can be applied for obtaining a point-spread function of high signal-to-noise ratio (SNR). Obtaining a high SNR represents one of the major hurdles in brightfield deconvolution. Although this method requires high contrast and thin sample thickness, it would be appropriate for imaging of the tail fin due to its reduced width.
A clear advantage of the presented imaging method is that it is rapidly adaptable to any stereomicroscope equipped with a CCD camera and time-lapse software and offers a low-cost alternative to more expensive confocal imaging systems. While this method does not utilize fluorescence for cell detection, it can be extended for such applications by utilizing an automated system for shutter control and post-imaging deconvolution software 34. This would enable users to further observe wound repair and regeneration processes with single cell or subcellular resolution over longer time periods.
The optical clarity and ease with which embryonic and larval zebrafish can be handled, and the adaptability of this method to any stereomicroscope makes it suitable for teaching basic vertebrate biology in a classroom setting. This method can provide students with a better understanding of the basic biological processes underlying tissue repair and regeneration. Other biological processes that have been captured with a similar method are zebrafish embryonic development 23,34 and cardiac function (unpublished). This method also offers the possibility for monitoring wound repair and regeneration in larvae that have been genetically and pharmacologically manipulated.

The ability of an organism to orchestrate tissue repair processes after injury is crucial for its survival 1. While all animals have the capacity to heal their wounds, the extent to which tissues regenerate differs greatly among species. Vertebrate species such as zebrafish, salamanders and frog tadpoles have the remarkable ability to regenerate lost tissues, including their appendages, portions of their eyes, heart, and central nervous system 2-4. Mammalian species, such as the African spiny mouse and rabbits, are capable of regenerating holes in their pinnae 5-7, and humans and mice regenerate portions of their liver as well as their digit tips during fetal and juvenile stages 8-12. Although it is not well understood yet why and how certain species regenerate tissues more effectively than others, the presence of similar genetic pathways suggests that these mechanisms may lie dormant in species without great regeneration potential 13,14. Thus elucidating tissue repair and regeneration mechanisms in species with satisfactory regeneration outcomes will benefit regeneration in humans.
We have chosen the larval zebrafish tail fin as a paradigm to demonstrate its regeneration with time-lapse brightfield stereomicroscopy. The zebrafish larval tail fin is anatomically simple as compared to the more complex adult structures, consisting of a two-layered, infolded epithelium with somatosensory axons innervating the skin that surrounds medially located mesenchymal cells 15. Despite the anatomical differences, larval tail fin regeneration is somewhat comparable to adult fin regeneration in terms of the molecular signatures and the outgrowth responses 16,17. As compared to the adult fin, imaging larval tail fin regeneration has however several advantages: 1) larval fin regeneration is completed within just 2-3 days 16, 2) larvae can be mounted in low-melt agarose, and 3) larvae do not require feeding until ~ 5 days post fertilization (dpf) due to the presence of the yolk sac. This makes zebrafish larvae ideal for observing tissue repair dynamics in vivo.
The presented method enables the capture of detailed dynamics underlying the early processes of fin regeneration. Many studies have utilized fluorescence-based confocal microscopy to study cellular and subcellular biological processes in embryonic and larval zebrafish. Sophisticated confocal imaging setups are however often not accessible to everyone and highly expensive as compared to other imaging techniques. In contrast, the presented methodology utilizes a Discovery V12 stereomicroscope equipped with Axiovision software and a time-lapse module, thus providing a more affordable alternative to expensive imaging equipment to examine tissue behaviors. We demonstrate that this method can be utilized for imaging tissue regeneration with high temporal resolution at a minimal cost. The implications for this method could extend beyond basic biology to advance mammalian regeneration studies using organ cultures, for therapeutic development through pharmacological and genetic screens, and it can serve as a teaching tool in a classroom setting.

<table><tbody><tr><td colspan="4">&lt;strong&gt;Reagents&lt;/strong&gt;</td></tr><tr><td>Bullseye Agarose</td><td>MidSci</td><td>BE-GCA500</td><td></td></tr><tr><td>Low-melt agarose</td><td>Fisher BioReagents</td><td>BP1360-100</td><td></td></tr><tr><td>1-phenyl-2-thiourea</td><td>Alfa Aesar</td><td>L06690</td><td></td></tr><tr><td>Instant Ocean Aquarium Salt</td><td>Pet store</td><td></td><td></td></tr><tr><td>Methylene Blue (0.1% solution)</td><td>Sigma</td><td>M9140</td><td></td></tr><tr><td>Tricaine (Ethyl 3-aminobenzoate methanesulfonate)</td><td>Sigma-Aldrich</td><td>E10505</td><td></td></tr><tr><td>2-Phenoxyethanol</td><td>Sigma-Aldrich</td><td>77699</td><td></td></tr><tr><td>Petri Dish 35 x 15 mm</td><td>BD Falcon</td><td>351008</td><td></td></tr><tr><td>Petri Dish 60 x 15 mm</td><td>BD Falcon</td><td>351007</td><td></td></tr><tr><td>Petri Dish 100 x 25 mm</td><td>BD Falcon</td><td>351013</td><td></td></tr><tr><td>5.75 inch boroschillate glass pipets</td><td>Fisher</td><td></td><td></td></tr><tr><td>35 mm Glass Top Glass Bottom Dish (Glass: 0.085-0.115mm)</td><td>MatTek Corporation</td><td>D35-20-0-TOP</td><td></td></tr><tr><td>Superfrost/Plus microscope slides</td><td>Fisherbrand</td><td>12-550-15</td><td></td></tr><tr><td>Glass coverslips</td><td>Electron Microscopy Services</td><td>72191-75</td><td></td></tr><tr><td>Glass coverslips</td><td>Warner Instruments</td><td>CS-18R15</td><td></td></tr><tr><td>Phifer Phiferglass Insect Screen Charcoal - 48&quot;</td><td>Home Depot</td><td></td><td></td></tr><tr><td>High vacuum grease</td><td>Dow Corning</td><td></td><td></td></tr><tr><td>Microloader pipette tips 20 &amp;micro;l</td><td>Eppendorf</td><td>930001007</td><td></td></tr><tr><td>Fine Scissors - Sharply Angled Up</td><td>Fine Science Tools</td><td>14037-10</td><td></td></tr><tr><td>3 ml Luer-Lok&amp;trade; disposable syringe&amp;nbsp;</td><td>BD</td><td>309657</td><td></td></tr><tr><td>60 ml Luer-Lok&amp;trade; disposable syringe</td><td>BD</td><td>309653</td><td></td></tr><tr><td>23 G syringe needles</td><td>BD</td><td>305145</td><td></td></tr><tr><td>Dumont #5 Forceps</td><td>Fine Science Tools</td><td>11295-00</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Equipment&lt;/strong&gt;</td></tr><tr><td>LabDoctor Mini Dry Bath</td><td>MidSci</td><td></td><td></td></tr><tr><td>Discovery.V12 compound microscope&amp;nbsp;</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Plan Apo S 3.5X objective</td><td>Zeiss</td><td></td><td></td></tr><tr><td>AxioCam MRm&amp;nbsp;</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Axiovision software, Release 4.8.2SP1 (12-2011)</td><td>Zeiss</td><td></td><td></td></tr></tbody></table>

capturing tissue repair in zebrafish larvae with time-lapse brightfield stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

The presented technique is suitable to elucidate tissue repair dynamics in response to amputation. The movie demonstrates that amputation of the fin initially triggers a purse-string effect, characterized by contractions via actin-myosin cables that are present in the fin-fold 28 (Figure 5 A,B). Concomitantly, cells are extruded from the wound (see movie). The contraction may thus be a means to expel cells that are likely destined to undergo cell death. Our results further show that the developmental growth of the larva occurs independently of regeneration (movie), whereas fin regeneration does not initiate until about 14 hr post amputation as measured by fin length and area over the time course of 36 hr following amputation (Figure 5C, D). The total regenerative fin growth after 1.5 days was about 60% of the original fin length (Figure 5E). Taken together, these results demonstrate that amputation triggers fin contraction, extrusion of cells from the wound, and a temporally delayed regenerative response. While extruded cells are likely destined to undergo cell death, the nature of these cells needs to be further clarified.
<img alt="Figure 1" src="/files/ftp_upload/52654/52654fig1.jpg" /> 
Figure 1.&#160;Imaging chamber ring assembly
(A) Shown is a plastic ring that is attached to a coverslip with silicon grease. A plastic mesh is attached to the inside of the chamber with four small dots of silicon grease. (B) The chamber containing the mounted larva is filled with Tricaine solution and a glass slide is attached to the top. (C) A mounted 2 day-old larva (arrow) is shown at higher magnification to depict its size in relation to the mesh. <a href="https://www.jove.com/files/ftp_upload/52654/52654fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52654/52654fig2.jpg" /> 
Figure 2. Imaging chamber assembly made from Petri dishes. (A) Shown is a commercial glass top glass bottom petri dish with a plastic mesh attached to the glass coverslip. (B) Shown is a self-constructed Petri dish chamber with a hole drilled into the lid and a coverslip attached from the outside with silicon grease. The mesh and larva are mounted inside the chamber containing Tricaine solution. To seal the chamber, silicon grease is applied to the upper, outer rim of the bottom chamber and the top lid attached. <a href="https://www.jove.com/files/ftp_upload/52654/52654fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52654/52654fig3.jpg" /> 
Figure 3. Scheme of amputation and mounting of a larva for imaging. (A) For amputation, place an anesthetized larva onto an agarose-coated Petri dish and amputate the tail fin with a syringe needle. (B) For mounting, transfer the larva with a transfer pipette into a 1.5 ml tube filled with 42 &#176;C liquid agarose and pipette a drop containing the larva into the imaging chamber, orient the fish and cover the solidified agarose with embryo medium. (C) Scrape off the agarose from the tail fin using a capped microloader pipette tip or similar tool and replace embryo medium with fresh medium. (D) Image the tail fin under a stereomicroscope. <a href="https://www.jove.com/files/ftp_upload/52654/52654fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/52654/52654fig4.jpg" /> 
Figure 4. Self-constructed heated incubation chamber. (A-C) Shown is a heated incubation chamber made of cardboard, bubble wrap and Velcro. A wired dome heater (originally designed for chicken egg incubation) is attached to the chamber using aluminum tape. <a href="https://www.jove.com/files/ftp_upload/52654/52654fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/52654/52654fig5.jpg" /> 
Figure 5. Fin regeneration dynamics. (A) Scheme of the utilized tail fin amputation assay and quantification method to determine the fin length (red arrow) and area (red outline of the fin). (B) Tail fin amputation initially triggers contraction of the fin, followed by regenerative tissue outgrowth. The fin also undergoes developmental growth, as evidenced by the lateral size increase. (C) Shown is the fin length as a function of time, revealing a linear regenerative growth starting at ~14 hpa. (D) Quantification of the fin area reveals an initially decrease in size, which can be attributed to the contraction of the fin. After ~ 14 hr, the fin size increases at a linear rate. (E) Comparison of the fin length before amputation and after 36 hr&#160;shows ~ 60% regrowth. Scale bar: 100 &#181;m Abbreviations: pre-amp, pre-amputation; hours post amputation, hpa; regen, regeneration; amp, amputation <a href="https://www.jove.com/files/ftp_upload/52654/52654fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<a href="https://www.jove.com/files/ftp_upload/52654/Movie_1_H264.mov">Movie</a>. Fin regeneration over the time course of 36 hr. Shown is a tail fin of a 2.5-day old larva during the course of regeneration. Starting 30 min post amputation regenerative growth is imaged in 30 min intervals on a stereomicroscope using a 3.5x objective lens.

Watch this Scientific Journal Video about Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy at JoVE.com

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

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Research

JoVE Journal

Developmental Biology

10.8K Views.  MDI Biological Laboratory. We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

Video: Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

Zebrafish Keratocyte Explants to Study Collective Cell Migration and Reepithelialization in Cutaneous Wound Healing

Due to their unique motile properties, fish keratocytes dissociated from explant cultures have long been used to study the mechanisms of single cell migration. However, when explants are established, these cells also move collectively, maintaining many of the features which make individual keratocytes an attractive model to study migration: rapid rates of motility, extensive actin-rich lamellae with a perpendicular actin cable, and relatively constant speed and direction of migration. In early explants, the rapid interconversion of cells migrating individually with those migrating collectively allows the study of the role of cell-cell adhesions in determining the mode of migration, and emphasizes the molecular links between the two modes of migration. Cells in later explants lose their ability to migrate rapidly and collectively as an epithelial to mesenchymal transition occurs and genes associated with wound healing and inflammation are differentially expressed. Thus, keratocyte explants can serve as an in vitro model for the reepithelialization that occurs during cutaneous wound healing and can represent a unique system to study mechanisms of collective cell migration in the context of a defined program of gene expression changes. A variety of mutant and transgenic zebrafish lines are available, which allows explants to be established from fish with different genetic backgrounds. This allows the role of different proteins within these processes to be uniquely addressed. The protocols outlined here describe an easy and effective method for establishing these explant cultures for use in a variety of assays related to collective cell migration.

Zebrafish keratocytes migrate in cell sheets from explants and provide an in vitro model for the study of the mechanisms of collective cell migration in the context of epithelial wound healing. These protocols detail an effective way to establish primary explant cultures for use in collective cell migration assays.

Due to their unique motile properties, fish keratocytes dissociated from explant cultures have long been used to study the mechanisms of single cell ...

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

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

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

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

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

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

Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, the muscle stem cell is activated and undergoes cell proliferation to generate a pool of myoblasts, which subsequently differentiate to form new muscle fibers. In many muscle wasting conditions, including muscular dystrophy and ageing, this process is impaired resulting in the inability of muscle to regenerate. The process of muscle regeneration in zebrafish is highly conserved with mammalian systems providing an excellent system to study muscle stem cell function and regeneration, in muscle wasting conditions such as muscular dystrophy. Here, we present a method to examine muscle regeneration in zebrafish models of muscle disease. The first step involves the use of a genotyping platform that allows the determination of the genotype of the larvae prior to eliciting an injury. Having determined the genotype, the muscle is injured using a needle stab, following which polarizing light microscopy is used to determine the extent of muscle regeneration. We therefore provide a high throughput pipeline which allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle regeneration is driven by tissue resident muscle stem cells, which are impaired in many muscle diseases such as muscular dystrophy, and this results in the inability of muscle to regenerate. Here, we describe a protocol that allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, ...

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.

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 ...

Bioengineering

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 Chemokine Receptors in Zebrafish Neutrophils During Wound Responses

Leukocyte guidance by chemical gradients is essential for immune responses. Neutrophils are the first cells to be recruited to sites of tissue damage where they execute crucial antimicrobial functions. Their trafficking to these loci is orchestrated by several inflammatory chemoattractants, including chemokines. At the molecular level, chemoattractant signaling is regulated by the intracellular trafficking of the corresponding receptors. However, it remains unclear how subcellular changes in chemokine receptors affect leukocyte migration dynamics at the cell and tissue level. Here we describe a methodology for live imaging and quantitative analysis of chemokine receptor dynamics in neutrophils during inflammatory responses to tissue damage. These tools have revealed that differential chemokine receptor trafficking in zebrafish neutrophils coordinates neutrophil clustering and dispersal at sites of tissue damage. This has implications for our understanding of how inflammatory responses are self-resolved. The described tools could be used to understand neutrophil migration patterns in a variety of physiological and pathological settings and the methodology could be expanded to other signaling receptors.

Here we describe protocols to perform live imaging and quantitative analysis of chemoattractant receptor dynamics in zebrafish neutrophils

Leukocyte guidance by chemical gradients is essential for immune responses. Neutrophils are the first cells to be recruited to sites of tissue damage ...

Immunology and Infection

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 ...

Low-cost Polyethylene Terephthalate Lamination Microfluidics Designs for Multiplexed Zebrafish Imaging

Zebrafish embryos are transparent and thus uniquely suited for noninvasive intravital imaging of fundamental processes, such as wound healing and immune cell migration. Microfluidic devices are used for entrapment to support long-term imaging of multicellular organisms, including zebrafish. However, the fabrication of these devices using soft lithography requires specialized facilities and competency in 3D printing, which may not be accessible to every lab. Our adaptation of a previously developed low-cost polyethylene terephthalate lamination method for constructing microfluidic devices increases accessibility by enabling design fabrication and iteration for a fraction of the technical investment of conventional techniques. We use a device made with this method, the Rotational Assistant for Danio Imaging of Subsequent Healing (RADISH), to accommodate drug treatment, manual wounding, and long-term imaging of up to four embryos in the same field of view. With this new design, we successfully capture gross morphological characteristics of the calcium&#160;signal around laser ablation and manual transection wounds for multiple embryos in the 2 h immediately following injury, as well as neutrophil recruitment to the wound edge for 24 h.

An innovative method for fabricating microfluidic devices using polyethylene terephthalate (PET) lamination significantly reduces the cost and complexity of entrapping and imaging multiple live zebrafish embryos.

Zebrafish embryos are transparent and thus uniquely suited for noninvasive intravital imaging of fundamental processes, such as wound healing and immune ...

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

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