We appreciate the support provided by William Archer and the Notre Dame Integrated Imaging Facility. Special thanks are directed to the Freimann Life Sciences technicians for their continuous help and their care and husbandry of the zebrafish. This study was supported by grants from the National Eye Institute of NIH to DRH (R01-EY018417, R01-EY024519) and the Center for Zebrafish Research, University of Notre Dame, Notre Dame, IN.

Note: Zebrafish were raised and maintained in the Notre Dame Zebrafish facility in the Freimann Life Sciences Center. The methods described in this manuscript are approved by the University of Notre Dame Animal Care and Use Committee and are in compliance with the statement for the use of animals in vision research by the Association for Research in Vision and Ophthalmology.
1. Solutions
<ol>
	<li>Prepare 70% ethanol to sterilize the tissue culture hood and any equipment/reagents that are transferred into the tissue culture hood.</li>
	<li>Add 2 mL of 2-phenoxyethanol to 1 L of system water (1:500 2-phenoxyethanol).</li>
	<li>Prepare 0.1 mM NaHCO3, pH 8.0. Use a 10 mL&#160;syringe and a 0.2 &#181;m pore-size syringe filter to sterilize the solution in a sterilized tissue culture hood. 
	NOTE: NaHCO3 is used to efficiently spread the cell and tissue adhesive across the coverslip surface of fluorodishes (see step 3.2).</li>
	<li>Prepare 1.0 M CaCl2 and 1.0 M MgCl2. Sterilize with a 10 mL syringe and 0.2 &#181;m pore-size syringe filter in a sterilized tissue culture hood.</li>
	<li>To prepare Hank&#39;s-balanced Salt Solution (HBSS), add sterile CaCl2 and sterile MgCl2 to 1x HBSS without Ca2+/Mg2+, without phenol red at a final concentration of 1 mM each. Work in a sterile environment.</li>
	<li>To prepare culture medium, mix 50% 1x Minimum Essential Medium (MEM) without phenol red, 25% HBSS containing CaCl2 and MgCl2 (see section 1.4), 25% Horse Serum (HS), 10 units/mL penicillin, and 10 &#181;g/mL streptomycin. Work in a sterile environment. 
	NOTE: Avoid medium containing phenol red as it autofluoresces, thereby affecting signal to noise ratios.23</li>
	<li>Prepare 10 mL&#160;of 1% low melting point agarose in 1x MEM without phenol red. Melt agarose/1x MEM using a microwave. Prepare small volumes of agarose (e.g., 10 mL)&#160;as repetitive reheating will change ion concentrations due to fluid evaporation.</li>
	<li>Prior to culturing, sterilize 1.5 mL microcentrifuge tubes by autoclaving.</li>
</ol>
2. Light-damage Paradigm
<ol>
	<li>Dark-adaptation
	<ol>
		<li>Place 2 - 3&#160;Tg[gfap:nGFP]mi2004 zebrafish (or other transgenic zebrafish of interest) at 6 - 14 months of age into an environment devoid of light for 14 d. For further detail see reference2, 6, 24, 25.</li>
	</ol>
	</li>
	<li>Position the tank containing 2 - 3 dark-adapted transgenic zebrafish in system water between two fluorescent lamps that emit light of 2,800 lux2, 6, 24, 25.</li>
	<li>Expose zebrafish to constant intense light for 35 h. During the light exposure, assure that the water temperature is maintained between 31 - 33 &#176;C.</li>
</ol>
3. Preparation for Culturing (On the Day of Retinal Isolation)
<ol>
	<li>Using 70% ethanol, sterilize the tissue culture hood and the components/tools that are transferred into the tissue culture hood (e.g., flasks containing MEM and HBSS, pipettes, sterile pipette tips, etc.).</li>
	<li>Preparation of fluorodishes
	<ol>
		<li>Coat one fluorodish for each retina with cell and tissue adhesive (see Table of Materials).</li>
		<li>Dilute the cell and tissue adhesive in 0.1 mM NaHCO3 to a final concentration of 70 &#181;g/mL, add 50 &#181;L to each fluorodish and spread the solution across the center of the fluorodish with a 200 &#181;L pipette tip (approximately 1 - 1.2 cm diameter).</li>
		<li>Incubate the coated fluorodishes for 1 - 3 h at RT&#160;(alternatively, incubate O/N at 4 &#176;C).</li>
		<li>Remove the solution and rinse 3x with 500 &#181;L of 1x MEM for each wash. To avoid the coated fluorodishes from drying out, maintain them in MEM until retinal explants are mounted.</li>
	</ol>
	</li>
	<li>Prepare culture medium as described in step 1.6. The culture medium can be stored for up to 1 week at 4 &#176;C.</li>
</ol>
4. Isolation and Culturing of Retinal Explants
NOTE: The protocol outlined below is for the isolation of the dorsal retina, which is the retinal region that is predominantly lesioned by the described light-damage paradigm. Therefore, damage-induced proliferation and the associated event of interkinetic nuclear migration occur in the dorsal retina. However, the isolation procedure can be adjusted to yield retinal regions according to the specific requirements of the researcher/research question.
<ol>
	<li>Euthanize one light-damaged transgenic zebrafish at a time in 1:500 2-phenoxyethanol.</li>
	<li>Remove MEM from one fluorodish with a 1,000 &#181;L pipette so that only a thin film of fluid remains.</li>
	<li>Transfer the zebrafish onto a dry paper towel, remove the eye with a curved pair of Dumont forceps (forceps #5, 45&#176; angle) and transfer it onto the fluorodish.</li>
	<li>Using a stereomicroscope, orient the eye with the pupil onto the cover slip of the fluorodish so that the back of the eye with the optic nerve is visible (Figure 1D).</li>
	<li>With a pair of McPherson-Vannas scissors remove the optic nerve, cutting close to the back of the eye. In addition, remove connective tissue lining the outside of the eye.</li>
	<li>Hold the eye between its nasal and temporal side with a pair of #5 forceps while making an incision at the optic stalk by piercing with one scissor blade through the lamina cribrosa and cutting along both the nasal and temporal sides of the eye (see Figure 1E, segmented line).</li>
	<li>Using two pairs of #5 forceps, one for holding the dorsal side of the retina and the other pair to separate the ventral from the dorsal retina, pull the tissue. 
	NOTE: It is advisable to orient the dorsal retina so that the lens and the ganglion cell layer face the coverslip while the sclera is facing upwards.</li>
	<li>Remove the sclera from the dorsal retina with one pair of #5 forceps, while holding the lens that is connected to the retina with a second pair of #5 forceps (Figure 1H, I).</li>
	<li>To remove the lens, use McPherson-Vannas scissors and cut behind the lens without damaging the retina (Figure 1I, J). Sometimes, the lens separates in step 4.7. In this case, remove the sclera carefully with forceps by holding the retina at a cut edge with a second pair of #5 forceps.</li>
	<li>Remove the vitreous while removing the lens without damaging the retina. Flatten the retina with the ganglion cell layer facing the cover slip of the fluorodish (Figure 1L, M).</li>
	<li>Surround the retina with 10 &#181;L of 1% low melting point agarose and let the agarose solidify.</li>
	<li>Watch that the liquid agarose does not lift the retina as even a slight elevation might affect the ability to focus deeply into the tissue. If lifting is observed, use a pipette to remove agarose. Let residual agarose set before attempting to add more.</li>
	<li>Repeat step 4.12 several times before adding 1% low melting point agarose to cover the entire fluorodish. Once the agarose has solidified, add 1.5 mL culture medium.</li>
	<li>Maintain the retinal explant culture in a 5% CO2/air environment set at 32 &#176;C for approximately 12 h to allow the retina to recover from the stress incurred by the isolation procedure. Set the temperature to 32 &#176;C to maintain retinas at the same temperature as light-treated zebrafish (see step 2.3).</li>
</ol>
5. Multiphoton Microscopy
NOTE: The experiments performed in this manuscript were optimized for a multiphoton microscope equipped with an infrared laser (see Table of Materials), a 40X Apo long distance water immersion objective (N.A. 1.15), a galvanometer scanner and an environmental chamber that contains an insert for four 35 mm Petri dishes. The images were acquired with a non-descanned detector (R-NDD).
<ol>
	<li>Prior to imaging, equilibrate the environmental chamber to achieve a 5% CO2/air atmosphere. Make sure that empty Petri dishes are inserted into the holder to avoid leakage of gas into the room.</li>
	<li>Turn on the microscopy system.</li>
	<li>Once the environmental chamber is equilibrated, add refractive index liquid onto the 40X Apo long distance water immersion objective (N.A. 1.15). 
	NOTE: The refractive index liquid with optical properties similar to water is used to avoid evaporation of water during long-term imaging.</li>
	<li>Place fluorodishes with retinal explants into the chamber. Using brightfield light, position the specimen into the light path and bring the midregion of the dorsal retina into the plane of focus.</li>
	<li>Use GFP epifluorescent light to focus on gfap:nGFP-positive M&#252;ller glia nuclei (Figure 2A, C). 
	NOTE: If explants are not mounted flat or agarose accumulated under the explant, it will be difficult to focus onto the gfap:nGFP-positive nuclei or they will fluoresce dimly.
	<ol>
		<li>Check whether moving to a different region within the same retinal explant will overcome the focusing issue. Otherwise move to a different retinal explant.</li>
	</ol>
	</li>
	<li>In the image acquisition software, open the &#39;A1 MP GUI&#39;, the &#39;TiPad&#39;, the &#39;A1 Compact GUI&#39; and the &#39;ND acquisition&#39; windows. For multiphoton imaging, ensure that the &#39;IR NDD&#39; option is chosen in the &#39;A1 Compact GUI&#39;.</li>
	<li>In the &#39;setting&#39; field, select IR-DM for the 1st dichroic mirror and choose the band pass filter 525/50 to acquire GFP fluorescence.</li>
	<li>Switch on the IR laser in the window labeled &#39;A1MP GUI&#39;. It will take a few minutes for the laser to be ready. Set the wavelength to 910 nm to excite GFP fluorescence and align the laser by clicking the &#39;Auto alignment&#39; button in the &#39;A1 MP GUI&#39; window.</li>
	<li>Ensure that room and equipment lights are switched off or covered before opening the shutter in the &#39;A1 MP GUI&#39; to avoid overexposure of the photomultiplier tube. To reduce noise levels, house the microscope in a darkened environment.</li>
	<li>Acquire images of a field of view of 300 x 300 pixels, at a zoom of two, and a pixel dwelling time of 4.8 &#181;s/pixel. Roughly set up the laser power by changing the &#39;acquisition area&#39; in the &#39;A1MP GUI&#39; and the gain in the &#39;A1 Compact GUI&#39; window.</li>
	<li>Setting up the z-stack
	<ol>
		<li>Focus on the ganglion cell layer to set the top focal plane of the z-stack in the &#39;z&#39;-subwindow within the &#39;ND acquisition&#39; window. Some gfap:nGFP-positive cells are typically located in the ganglion cell layer, which helps to identify the basal limit of the retina (Figure 2A, D).</li>
		<li>Move the focal plane through the level of the ONL (Figure 2A, B), which is characterized by the presence of dimly labeled gfap:nGFP-positive cells that are round and enlarged relative to their counterparts in the INL (Figure 2A, C).</li>
		<li>Set this plane as the bottom of the z-stack. Ensure that the entire ONL will be imaged (Figure 2A, B).</li>
		<li>When experiencing focal plane shifts that require re-adjusting during the imaging period, double-click on the middle position in the &#39;z&#39; subwindow to assign it as the &#39;home&#39; position. Change to &#39;symmetric mode defined&#39; and click &#39;relative&#39;.</li>
		<li>Set the z-step size between 0.7 to 1 &#181;m.</li>
	</ol>
	</li>
	<li>Z-intensity correction:
	<ol>
		<li>Apply z-intensity corrections to compensate for loss of pixel intensity due to light scattering when imaging in deep layers of the tissue.</li>
		<li>To set up the correction, open the &#39;z-intensity correction&#39; window. To set the &#39;z-stack range&#39;, choose &#39;From ND&#39;.</li>
		<li>Click on the bottom focal plane in the &#39;z-intensity correction&#39; window (in this case, corresponds to the ganglion cell layer) and set the laser intensity (&#39;acquisition area&#39; in the &#39;A1MP GUI&#39;) and gain (A1 Compact GUI).</li>
		<li>Click the arrow next to the &#39;z-values&#39; in the &#39;z-intensity correction&#39; window to confirm the settings that are subsequently shown under &#39;device settings&#39; in the &#39;z-intensity correction&#39; window for the chosen focal plane.</li>
		<li>Repeat the process for the middle and top planes, increasing the laser power and gain. Additional focal planes can be added if necessary. See Table 1 for specific laser and gain settings for experiments in Figures 2 - 4.</li>
		<li>Set the &#39;acquisition area&#39; in the &#39;A1 MP GUI&#39; window. Avoid selecting an acquisition area larger than 15 and a gain higher than 126 at the start of imaging to circumvent photobleaching and increased noise levels. 
		NOTE: As lasers and photomultiplier tubes differ between microscopy systems, test laser and gain settings to obtain optimal imaging conditions for the microscope set up while avoiding photobleaching.</li>
		<li>Choose &#39;relative intensity correction&#39; in the &#39;z-intensity correction&#39; window.</li>
	</ol>
	</li>
	<li>In the &#39;timeseries&#39; subwindow in the &#39;ND acquisition&#39; window, set the duration to 8 h and the interval to &#39;no delay&#39;. Then, make sure to click the &#39;Run z-correction&#39; in the &#39;z-stack&#39; sub window in the &#39;ND acquisition&#39; window button to acquire the 3-D timeseries.</li>
	<li>Maintain retinal explants at a temperature of 27 - 29 &#176;C throughout the duration of image acquisition.</li>
	<li>Throughout the image acquisition period, if necessary, readjust the power level and gain in order to maintain image quality for post-imaging analysis. For adjustments perform steps 5.11.2 - 5.11.4.</li>
	<li>If the focal plane shifts, pause or stop the run and perform step 5.11 again. 
	NOTE: If &#39;relative z-correction&#39; was chosen in step 5.12.7 and step 5.11.4 was performed it should not be necessary to readjust power and gain levels, unless extensive photobleaching occurred.</li>
</ol>
6. Analysis of Velocity
<ol>
	<li>Extract the time, setting a &#39;region of interest&#39; on the image. Use the &#39;time measurement&#39; tool and export the time values to a spreadsheet.</li>
	<li>Crop a region that contains a dividing gfap:nGFP-positive nucleus.</li>
	<li>Choose the cropped region so that it contains at least one nucleus that does not undergo INM in order to set a reference point to subsequently measure the distances that the dividing nucleus migrated in relation to the basal INL.&#160;NOTE: To choose a nucleus that remains in the basal INL, it helps to prepare a 3-D reconstruction of the timeseries.</li>
	<li>Alternatively, if a gfap:nGFP-positive cell is observed in the ONL throughout the acquisition period, use a vertical line of fixed length spanning from the ONL nucleus to the basal INL in order to identify a reference point.</li>
	<li>Using the &#39;show slices view&#39; function, generate orthogonal projections. Subsequently, change the mode from &#39;slice&#39; to &#39;maximum intensity projection&#39;.</li>
	<li>Turn off the &#39;xy&#39; view of the orthogonal maximum projection. Depending on the orientation at which the migrating/dividing nucleus is best visible, also turn off either the &#39;xz&#39;- or the &#39;yz&#39;-view.</li>
	<li>Click on the remaining image with the right mouse button and extract the &#39;xz&#39; or &#39;yz&#39;-image series with the &#39;Create new document from this view&#39; function. If necessary, rotate the image.</li>
	<li>Using the &#39;manual measurement&#39; function, draw a horizontal line across the image at the bottom level of the nucleus that remains in the basal INL and does not undergo INM (= reference point; Figure 4A - E, red horizontal line).</li>
	<li>Measure the distance between the reference line and the basal point of the migrating nucleus for the timeseries using the &#39;line measurement&#39; tool in the analysis software (see Figure 4A - E).</li>
	<li>Once the nuclear envelope breaks down, measure the basal position of the soma if it is identifiable following the diffusion of GFP into the entire cell.</li>
	<li>Using a spreadsheet software, plot the distance a nucleus migrated against the time passed (Figure 4F).
	<ol>
		<li>To determine the apical migration velocity (va), graph the distances traveled for the period before nuclear envelope breakdown occurs (Figure 4G).</li>
		<li>Select the data series within the chart, right-click the mouse and insert a linear regression curve including the corresponding function &#39;y=mx+c&#39;. The slope &#39;m&#39; in the function represents the velocity, va (Figure 4G).</li>
		<li>Repeat steps 6.11.1 and 6.11.2 for the first phase of rapid basal migration to determine the basal migration velocity, vb (Figure 4H).</li>
	</ol>
	</li>
</ol>

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R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+an+adult+rat+retinal+explant+model+for+screening+of+potential+retinal+ganglion+cell+neuroprotective+therapies.">Use of an adult rat retinal explant model for screening of potential retinal ganglion cell neuroprotective therapies.</a> Invest.Ophthalmol.Vis.Sci. 52 (6), 3309-3320 (2011).</li><li>Kustermann, S., Schmid, S., Biehlmaier, O., Kohler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Survival,+excitability,+and+transfection+of+retinal+neurons+in+an+organotypic+culture+of+mature+zebrafish+retina.">Survival, excitability, and transfection of retinal neurons in an organotypic culture of mature zebrafish retina.</a> Cell Tissue Res. 332 (2), 195-209 (2008).</li><li>Williams, P. R., Morgan, J. L., Kerschensteiner, D., Wong, R. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+imaging+of+retinal+whole+mounts.">In vitro imaging of retinal whole mounts.</a> Cold Spring Harb Protoc. 2013, (2013).</li><li>Johnson, T. V., Oglesby, E. N., Steinhart, M. R., Cone-Kimball, E., Jefferys, J., Quigley, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time-Lapse+Retinal+Ganglion+Cell+Dendritic+Field+Degeneration+Imaged+in+Organotypic+Retinal+Explant+Culture.">Time-Lapse Retinal Ganglion Cell Dendritic Field Degeneration Imaged in Organotypic Retinal Explant Culture.</a> Invest.Ophthalmol.Vis.Sci. 57 (1), 253-264 (2016).</li><li>Roehlecke, C., Schumann, U., Ader, M., Knels, L., Funk, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Influence+of+blue+light+on+photoreceptors+in+a+live+retinal+explant+system.">Influence of blue light on photoreceptors in a live retinal explant system.</a> Mol.Vis. 17, 876-884 (2011).</li><li>Coutu, D. L., Schroeder, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Probing+cellular+processes+by+long-term+live+imaging--historic+problems+and+current+solutions.">Probing cellular processes by long-term live imaging--historic problems and current solutions.</a> J.Cell.Sci. 126 (Pt 17), 3805-3815 (2013).</li><li>Thomas, J. L., Thummel, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+light+damage+paradigm+for+use+in+retinal+regeneration+studies+in+adult+zebrafish.">A novel light damage paradigm for use in retinal regeneration studies in adult zebrafish.</a> J.Vis.Exp. 80, e51017 (2013).</li><li>Conner, C., Ackerman, K. M., Lahne, M., Hobgood, J. S., Hyde, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Repressing+notch+signaling+and+expressing+TNFalpha+are+sufficient+to+mimic+retinal+regeneration+by+inducing+Muller+glial+proliferation+to+generate+committed+progenitor+cells.">Repressing notch signaling and expressing TNFalpha are sufficient to mimic retinal regeneration by inducing Muller glial proliferation to generate committed progenitor cells.</a> J.Neurosci. 34 (43), 14403-14419 (2014).</li><li>Nelson, C. M., Gorsuch, R. A., Bailey, T. J., Ackerman, K. M., Kassen, S. C., Hyde, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stat3+defines+three+populations+of+Muller+glia+and+is+required+for+initiating+maximal+muller+glia+proliferation+in+the+regenerating+zebrafish+retina.">Stat3 defines three populations of Muller glia and is required for initiating maximal muller glia proliferation in the regenerating zebrafish retina.</a> J.Comp.Neurol. 520 (18), 4294-4311 (2012).</li><li>Gorsuch, R. A., Hyde, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+Muller+glial+dependent+neuronal+regeneration+in+the+damaged+adult+zebrafish+retina.">Regulation of Muller glial dependent neuronal regeneration in the damaged adult zebrafish retina.</a> Exp.Eye Res. 123, 131-140 (2014).</li><li>Zhao, X. F., Wan, J., Powell, C., Ramachandran, R., Myers, M. G., Goldman, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Leptin+and+IL-6+family+cytokines+synergize+to+stimulate+Muller+glia+reprogramming+and+retina+regeneration.">Leptin and IL-6 family cytokines synergize to stimulate Muller glia reprogramming and retina regeneration.</a> Cell.Rep. 9 (1), 272-284 (2014).</li><li>Wan, J., Ramachandran, R., Goldman, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=HB-EGF+is+necessary+and+sufficient+for+Muller+glia+dedifferentiation+and+retina+regeneration.">HB-EGF is necessary and sufficient for Muller glia dedifferentiation and retina regeneration.</a> Dev.Cell. 22 (2), 334-347 (2012).</li><li>Lenkowski, J. R., 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=Retinal+regeneration+in+adult+zebrafish+requires+regulation+of+TGFbeta+signaling.">Retinal regeneration in adult zebrafish requires regulation of TGFbeta signaling.</a> Glia. 61 (10), 1687-1697 (2013).</li><li>Weber, I. P., Ramos, A. P., Strzyz, P. J., Leung, L. C., Young, S., Norden, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitotic+position+and+morphology+of+committed+precursor+cells+in+the+zebrafish+retina+adapt+to+architectural+changes+upon+tissue+maturation.">Mitotic position and morphology of committed precursor cells in the zebrafish retina adapt to architectural changes upon tissue maturation.</a> Cell.Rep. 7 (2), 386-397 (2014).</li><li>Magidson, V., Khodjakov, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Circumventing+photodamage+in+live-cell+microscopy.">Circumventing photodamage in live-cell microscopy.</a> Methods Cell Biol. 114, 545-560 (2013).</li><li>Bailey, T. J., Fossum, S. L., Fimbel, S. M., Montgomery, J. E., Hyde, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+inhibitor+of+phagocytosis,+O-phospho-L-serine,+suppresses+Muller+glia+proliferation+and+cone+cell+regeneration+in+the+light-damaged+zebrafish+retina.">The inhibitor of phagocytosis, O-phospho-L-serine, suppresses Muller glia proliferation and cone cell regeneration in the light-damaged zebrafish retina.</a> Exp.Eye Res. 91 (5), 601-612 (2010).</li><li>Lee, J. E., Liang, K. J., Fariss, R. N., Wong, W. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ex+vivo+dynamic+imaging+of+retinal+microglia+using+time-lapse+confocal+microscopy.">Ex vivo dynamic imaging of retinal microglia using time-lapse confocal microscopy.</a> Invest.Ophthalmol.Vis.Sci. 49 (9), 4169-4176 (2008).</li><li>Zhao, L., 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=Microglial+phagocytosis+of+living+photoreceptors+contributes+to+inherited+retinal+degeneration.">Microglial phagocytosis of living photoreceptors contributes to inherited retinal degeneration.</a> EMBO Mol.Med. 7 (9), 1179-1197 (2015).</li><li>Peri, F., Nusslein-Volhard, C. <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+neuronal+degradation+by+microglia+reveals+a+role+for+v0-ATPase+a1+in+phagosomal+fusion+in+vivo.">Live imaging of neuronal degradation by microglia reveals a role for v0-ATPase a1 in phagosomal fusion in vivo.</a> Cell. 133 (5), 916-927 (2008).</li><li>Morsch, 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=In+vivo+characterization+of+microglial+engulfment+of+dying+neurons+in+the+zebrafish+spinal+cord.">In vivo characterization of microglial engulfment of dying neurons in the zebrafish spinal cord.</a> Front.Cell.Neurosci. 9, 321 (2015).</li></ol>

The authors have nothing to disclose.

Studies investigating the mechanisms governing regeneration of the damaged adult zebrafish retina predominantly used immunocytochemical methods5,25,26,27,28,29,30. Establishing conditions to culture retinal explants and to perform live-cell imaging on phenomena, such as INM, provide us a tec...

Unlike humans, zebrafish (Danio rerio) exhibit a robust regeneration response upon cell death of retinal neurons1,2,3,4. Tumor necrosis factor &#945;, a signaling molecule that is released from dying retinal neurons induces M&#252;ller glia residing in the basal Inner Nuclear Layer (INL) of the retina, to proliferate5 and produce neuronal progenitor cells that continue to proliferate before differentiating into the neuronal cell types that died2,3,4. During the proliferative phase of the regeneration response, the nuclei of M&#252;ller glia and their derived neuronal progenitor cells undergo a repetitive migratory pattern in phase with the cell cycle (Interkinetic Nuclear Migration, INM)6,7. Nuclei positioned in the basal INL replicate their DNA before migrating to the Outer Nuclear Layer (ONL) where they divide before the arising nuclei return basally to the INL. This process was first described during neuroepithelial development using histological methods, while live-cell imaging approaches later confirmed the interpretation by Sauer8,9,10,11,12. Both histochemical and live-cell imaging approaches have been used to determine mechanisms underlying INM and its function in developing neuroepithelia including the retina9,11,12,13. However, the mechanisms governing INM in the adult regenerating retina have not been studied in much detail6,7. Live-cell imaging will be an invaluable approach to advance our knowledge of the signaling pathways that control INM in the adult regenerating retina.
Until recently, live-cell imaging of INM in the retina was limited to either live zebrafish embryos or to embryonic chick or postnatal mouse retinal explants9,10,11,12,14,15,16. While retinal explants from adult animals of a variety of species including mouse, rat and zebrafish have been utilized for different cell biological approaches17,18,19,20, live-cell imaging experiments using retinal explants have been restricted to brief periods of time and have not been executed continuously over several hours21,22. Here, we describe a detailed protocol to culture light-damaged adult zebrafish retinas to perform live-cell imaging experiments monitoring INM using multi-photon microscopy6. Live-cell imaging approaches are advantageous over immunohistochemical methods when investigating the mechanisms controlling INM, as the dynamics of INM, e.g., velocities might be affected rather than the location of mitosis, which would potentially not be detected using immunocytochemistry.
In the future, this method has also the potential to be modified to study other dynamic processes during retinal regeneration, such as phagocytosis of dying photoreceptors by M&#252;ller glia or the behavior of microglia.

<table><tbody><tr><td>Dumont forceps #5</td><td>World precision instruments</td><td>14098</td><td></td></tr><tr><td>Dumont forceps #5, 45&amp;deg; angle</td><td>World precision instruments</td><td>14101</td><td></td></tr><tr><td>McPherson-Vannas scissors</td><td>World precision instruments</td><td>501233</td><td></td></tr><tr><td>Fluordishes</td><td>World precision instruments</td><td>FD35-100</td><td></td></tr><tr><td>Stereomicroscope</td><td>Nikon&amp;nbsp;</td><td>SMZ-1B</td><td>similar type of dissection stereomicroscope will work</td></tr><tr><td>Biological Safety Cabinet class type A2</td><td>Labconco</td><td></td><td>equivalent type will work</td></tr><tr><td>tissue culture incubator</td><td>Thermoscientific</td><td>HEPA-class 100</td><td>equivalent type will work</td></tr><tr><td>Sylvania fluorescent lamps OSFP5835HOECO</td><td>Bulbtronics</td><td>31850</td><td></td></tr><tr><td>0.2 &amp;micro;m pore-size Acrodisc syringe filter</td><td>VWR</td><td>4192</td><td></td></tr><tr><td>10 mL Luer-lok syringe</td><td>VWR</td><td>BD309604</td><td></td></tr><tr><td>60 mL Luer-lok syringe</td><td>VWR</td><td>BD309653</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>FischerScientific</td><td>S233-500</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>ThermoScientific</td><td>C79-500</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>EMD Millipore</td><td>5980</td><td></td></tr><tr><td>HBSS w/o Ca&lt;sup&gt;2+&lt;/sup&gt;/Mg&lt;sup&gt;2+&lt;/sup&gt;, w/o phenol red, Gibco</td><td>ThermoScientific</td><td>14175-095</td><td></td></tr><tr><td>MEM w/o phenol red, Gibco</td><td>ThermoScientific</td><td>5100-038</td><td></td></tr><tr><td>Horse serum, heat-inactivated</td><td>ThermoScientific</td><td>26050-070</td><td></td></tr><tr><td>penicillin/streptomycin</td><td>VWR</td><td>16777-164</td><td></td></tr><tr><td>Ultrapure low melting point agarose</td><td>ThermoScientific</td><td>16520-100</td><td></td></tr><tr><td>ethanol, absolute</td><td>ThermoScientific</td><td>BP2818-4</td><td></td></tr><tr><td>2-phenoxyethanol</td><td>Sigma</td><td>77699</td><td></td></tr><tr><td>Corning Cell-Tak cell and tissue adhesive&amp;nbsp;</td><td>VWR</td><td>354240</td><td></td></tr><tr><td>refractive index liquid&amp;nbsp;</td><td>Cargille Lab</td><td>1803Y</td><td></td></tr><tr><td>Nikon A1 multiphoton microscope equipped with a MaiTai infrared laser</td><td>Nikon</td><td></td><td>equivalent system will work</td></tr><tr><td>40X Apo long-distance water immersion objective (N.A. 1.15)</td><td></td><td></td><td></td></tr><tr><td>environmental chamber equipped with insert for 35 mm petridishes</td><td>Okolab</td><td></td><td>equivalent system will work</td></tr><tr><td>NIS analysis software</td><td>Nikon</td><td></td><td></td></tr></tbody></table>

culture of adult transgenic zebrafish retinal explants for live-cell imaging by multiphoton microscopy

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The M&#252;ller glia re-enter the cell cycle and produce neuronal progenitor cells that undergo subsequent rounds of cell divisions and differentiate into the lost neuronal cell types. Both M&#252;ller glia and neuronal progenitor cell nuclei replicate their DNA and undergo mitosis in distinct locations of the retina, i.e. they migrate between the basal Inner Nuclear Layer (INL) and the Outer Nuclear Layer (ONL), respectively, in a process described as Interkinetic Nuclear Migration (INM). INM has predominantly been studied in the developing retina. To examine the dynamics of INM in the adult regenerating zebrafish retina in detail, live-cell imaging of fluorescently-labeled M&#252;ller glia/neuronal progenitor cells is required. Here, we provide the conditions to isolate and culture dorsal retinas from Tg[gfap:nGFP]mi2004 zebrafish that were exposed to constant intense light for 35 h. We also show that these retinal cultures are viable to perform live-cell imaging experiments, continuously acquiring z-stack images throughout the thickness of the retinal explant for up to 8 h using multiphoton microscopy to monitor the migratory behavior of gfap:nGFP-positive cells. In addition, we describe the details to perform post-imaging analysis to determine the velocity of apical and basal INM. To summarize, we established conditions to study the dynamics of INM in an adult model of neuronal regeneration. This will advance our understanding of this crucial cellular process and allow us to determine the mechanisms that control INM.

The isolation of the retina according to the procedure outlined in the schematic in Figure 1 allows the culturing of a flattened dorsal retina from light-damaged adult Tg[gfap:nGFP]mi2004 zebrafish over a period of at least 24 h in a 5% CO2/air environment. These flat-mounted retinal explants can be used to image focal planes at deep tissue levels. An example is M&#252;ller glia/neuronal progenitor cell nuclei labeled with GFP from the M&#25...

Watch this Scientific Journal Video about Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy at JoVE.com

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

Zebrafish retinal regeneration has mostly been studied using fixed retinas. However, dynamic processes such as interkinetic nuclear migration occur during the regenerative response and require live-cell imaging to investigate the underlying mechanisms. Here, we describe culture and imaging conditions to monitor Interkinetic Nuclear Migration (INM) in real-time using multiphoton microscopy.

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The ...

culture-adult-transgenic-zebrafish-retinal-explants-for-live-cell

Research

JoVE Journal

Developmental Biology

8.5K Views.  University of Notre Dame.  Zebravis netvlies regeneratie is meestal onderzocht met behulp van vaste netvliezen. Echter, dynamische processen zoals interkinetic nucleaire migratie optreden tijdens de regeneratieve respons en vereisen voor live-cell imaging om de onderliggende mechanismen te onderzoeken. We beschrijven hier de cultuur en beeldvorming voorwaarden te controleren Interkinetic Nuclear Migratie (INM) in real-time met behulp van multiphoton microscopie. 

Video: Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

Single-cell Photoconversion in Living Intact Zebrafish

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause disease when disrupted. Scientists have developed clever techniques to investigate characteristics and natural dynamics of these cells within intact tissue by expressing fluorescent proteins in subsets of cells. However, at times, experiments require more selected visualization of cells within the tissue, sometimes at the single-cell or population-of-cells manner. To achieve this and visualize single cells within a population of cells, scientists have utilized single-cell photoconversion of fluorescent proteins. To demonstrate this technique, we show here how to direct UV light to an Eos-expressing cell of interest in an intact, living zebrafish. We then image those photoconverted Eos+ cells 24 h later to determine how they changed in the tissue. We describe two techniques: single cell photoconversion and photoconversions of populations of cell. These techniques can be used to visualize cell-cell interactions, cell-fate and differentiation, and cell migrations, making it a technique that is applicable in numerous biological questions.

Here, we present a protocol to show how cell photoconversion is achieved through UV exposure to specific areas expressing the fluorescent protein, Eos, in living animals.

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause ...

Culture and Transfection of Zebrafish Primary Cells

Zebrafish embryos are transparent and develop rapidly outside the mother, thus allowing for excellent in vivo imaging of dynamic biological processes in an intact and developing vertebrate. However, the detailed imaging of the morphologies of distinct cell types and subcellular structures is limited in whole mounts. Therefore, we established an efficient and easy-to-use protocol to culture live primary cells from zebrafish embryos and adult tissue.
In brief, 2 dpf zebrafish embryos are dechorionated, deyolked, sterilized, and dissociated to single cells with collagenase. After a filtration step, primary cells are plated onto glass bottom dishes and cultivated for several days. Fresh cultures, as much as long term differenciated ones, can be used for high resolution confocal imaging studies. The culture contains different cell types, with striated myocytes and neurons being prominent on poly-L-lysine coating. To specifically label subcellular structures by fluorescent marker proteins, we also established an electroporation protocol which allows the transfection of plasmid DNA into different cell types, including neurons. Thus, in the presence of operator defined stimuli, complex cell behavior, and intracellular dynamics of primary zebrafish cells can be assessed with high spatial and temporal resolution. In addition, by using adult zebrafish brain, we demonstrate that the described dissociation technique, as well as the basic culturing conditions, also work for adult zebrafish tissue.

We present an efficient and easy-to-use protocol for preparing primary cell cultures of zebrafish embryos for transfection and live cell imaging as well as a protocol to prepare primary cells from adult zebrafish brain.

Zebrafish embryos are transparent and develop rapidly outside the mother, thus allowing for excellent in vivo imaging of dynamic biological processes in ...

Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their environment. Chimeric embryos, which are composed of cells of different genetic background, are great tools to study the cell-cell interactions mediated by genes of interest. The embryonic transparency of zebrafish at early developmental stages permits direct visualization of the morphogenesis of tissues and organs at the cellular level. Here, we demonstrate a protocol to generate chimeric retinas and brains in zebrafish embryos and to perform live imaging of the donor cells. The protocol covers the preparation of transplantation needles, the transplantation of GFP-expressing donor blastomeres to GFP-negative hosts, and the examination of donor cell behavior under live confocal microscopy. With slight modifications, this protocol can also be used to study the embryonic development of other tissues and organs in zebrafish. The advantages of using GFP to label donor cells are also discussed. 

We demonstrate a protocol to generate chimeric zebrafish embryos for live imaging cellular behavior during embryogenesis.

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their ...

Biology

Multicolor Time-lapse Imaging of Transgenic Zebrafish: Visualizing Retinal Stem Cells Activated by Targeted Neuronal Cell Ablation

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound transgenic fish which express different fluorescent reporters in neighboring cell types provide a means of following cellular interactions 2 and/or tissue-level responses to experimental manipulations over time. In this video, we demonstrate methods that can be used for imaging multiple transgenically labeled cell types serially in individual fish over time courses that can span from minutes to several days. The techniques described are applicable to any study seeking to correlate the "behavior" of neighboring cells types over time, including: 1) serial 'catch and release' methods for imaging a large number of fish over successive days, 2) simplified approaches for separating fluorophores with overlapping excitation/emission profiles (e.g., GFP and YFP), 3) use of hypopigmented mutant lines to extend the time window available for high-resolution imaging into late larval stages of development, 4) use of membrane targeted fluorescent reporters to reveal fine morphological detail of individual cells as well as cellular details in larger populations of cells, and 5) a previously described method for chemically-induced ablation of transgenically targeted cell types; i.e., nitroreductase (NTR) mediated conversion of prodrug substrates, such as metronidazole (MTZ), to cytotoxic derivatives 3,5.
As an example of these approaches, we will visualize the ablation and regeneration of a subtype of retinal bipolar neuron within individual fish over several days. Simultaneously we will monitor several other retinal cell types, including neighboring non-targeted bipolar cells and potential degeneration-stimulated retinal stem cells (i.e., M&#971;ller glia). This strategy is being applied in our lab to characterize cell- and tissue-level (e.g., stem cell niche) responses to the selective loss and regeneration of targeted neuronal cell types.

In this video, techniques for multicolor confocal time-lapse imaging and targeted cell ablation are provided. Time-lapse imaging is used to monitor the behavior of multiple cell types of interest in vivo. Targeted cell ablation facilitates the study neural circuit function and cell-specific neuronal regeneration paradigms.

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound ...

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 ...

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding diseases, and future therapies hinge on fundamental understandings about how different retinal cells function normally. Gaining such information with biochemical methods has proven difficult because contributions of particular cell types are diminished in the retinal cell milieu. Live retinal imaging can provide a view of numerous biological processes on a subcellular level, thanks to a growing number of genetically encoded fluorescent biosensors. However, this technique has thus far been limited to tadpoles and zebrafish larvae, the outermost retinal layers of isolated retinas, or lower resolution imaging of retinas in live animals. Here we present a method for generating live ex vivo retinal slices from adult zebrafish for live imaging via confocal microscopy. This preparation yields transverse slices with all retinal layers and most cell types visible for performing confocal imaging experiments using perfusion. Transgenic zebrafish expressing fluorescent proteins or biosensors in specific retinal cell types or organelles are used to extract single-cell information from an intact retina. Additionally, retinal slices can be loaded with fluorescent indicator dyes, adding to the method&#39;s versatility. This protocol was developed for imaging Ca2+ within zebrafish cone photoreceptors, but with proper markers it could be adapted to measure Ca2+ or metabolites in M&#252;ller cells, bipolar and horizontal cells, microglia, amacrine cells, or retinal ganglion cells. The retinal pigment epithelium is removed from slices so this method is not suitable for studying that cell type. With practice, it is possible to generate serial slices from one animal for multiple experiments. This adaptable technique provides a powerful tool for answering many questions about retinal cell biology, Ca2+, and energy homeostasis.

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

Imaging retinal tissue can provide single-cell information that cannot be gathered from traditional biochemical methods. This protocol describes preparation of retinal slices from zebrafish for confocal imaging. Fluorescent genetically encoded sensors or indicator dyes allow visualization of numerous biological processes in distinct retinal cell types.

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding ...

Neuroscience

4-Dimensional Imaging of Zebrafish Optic Cup Morphogenesis

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of visual impairment, yet mechanisms of eye morphogenesis are still poorly understood. The basic organization of the embryonic eye is conserved throughout vertebrates, thus live imaging of zebrafish embryos has become a powerful approach to directly observe eye development at real time under normal and pathological conditions. Dynamic cell processes including movements, morphologies, interactions, division, and death can be visualized in the embryo. We have developed methods for uniform labeling of subcellular structures and timelapse confocal microscopy of early eye development in zebrafish. This protocol outlines the method of generating capped mRNA for injection into the 1-cell zebrafish embryo, mounting embryos at optic vesicle stage (~12 hours post fertilization, hpf), and performing multi-dimensional timelapse imaging of optic cup morphogenesis on a laser scanning confocal microscope, such that multiple datasets are acquired sequentially in the same imaging session. Such an approach yields data that can be used for a variety of purposes, including cell tracking, volume measurements, three-dimensional (3D) rendering, and visualization. Our approaches allow us to pinpoint the cellular and molecular mechanisms driving optic cup development, in both wild type and genetic mutant conditions. These methods can be employed directly by other groups or adapted to visualize many additional aspects of zebrafish eye development.

This protocol describes an approach for in toto labeling and multidimensional imaging of zebrafish early eye development. We describe labeling, embedding, and four dimensional (4D) imaging using laser scanning confocal microscopy, and considerations for optimizing acquisition of datasets for dissecting mechanisms of optic cup morphogenesis.

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of ...

ROS Live Cell Imaging During Neuronal Development

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the different ROS, hydrogen peroxide (H2O2) is best characterized with respect to roles in cellular signaling. H2O2 has been implicated during the development in several species. For example, a transient increase in H2O2 has been detected in zebrafish embryos during the first days following fertilization. Furthermore, depleting an important cellular H2O2 source, NADPH oxidase (NOX), impairs nervous system development such as the differentiation, axonal growth, and guidance of retinal ganglion cells (RGCs) both in&#160;vivo and in vitro. Here, we describe a method for imaging intracellular H2O2 levels in cultured zebrafish neurons and whole larvae during development using the genetically encoded H2O2-specific biosensor, roGFP2-Orp1. This probe can be transiently or stably expressed in zebrafish larvae. Furthermore, the ratiometric readout diminishes the probability of detecting artifacts due to differential gene expression or volume effects. First, we demonstrate how to isolate and culture RGCs derived from zebrafish embryos that transiently express roGFP2-Orp1. Then, we use whole larvae to monitor H2O2 levels at the tissue level. The sensor has been validated by the addition of H2O2. Additionally, this methodology could be used to measure H2O2 levels in specific cell types and tissues by generating transgenic animals with tissue-specific biosensor expression. As zebrafish facilitate genetic and developmental manipulations, the approach demonstrated here could serve as a pipeline to test the role of H2O2 during neuronal and general embryonic development in vertebrates.

This protocol describes the use of a genetically encoded hydrogen peroxide (H2O2)-biosensor in cultured zebrafish neurons and larvae for assessing the physiological signaling roles of H2O2 during nervous system development. It can be applied to different cell types and modified with experimental treatments to study reactive oxygen species (ROS) in general development.

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the ...

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

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