The authors would like to acknowledge Jonathan Chou, MD for his assistance on preparation and editing of the final manuscript and Wenzhong Liu for the OCT data. We would also like to acknowledge support from the Macula Society Research Grant (AAF), support from an unrestricted grant to Northwestern University from Research to Prevent Blindness, Inc., New York, NY, USA, and support from NIH-EY019951.

All animals are treated in accordance with the Guide of the Care and Use of Laboratory Animals 2013 Edition, the Association for Research in Vision and Ophthalmology (ARVO) Statement for the Use of Animals in Ophthalmic and Vision Research, and as approved by the Institutional Animal Care and Use Committee for Northwestern University.
Note: The following procedure can be done entirely with one operator; however, it is much more efficiently conducted with two operators with the tasks split accordingly.
1. Prepare Laser and Pre-laser Station
<ol>
	<li>Position the laser and slit lamp where it can be easily accessible. Turn on laser and set to pre-determined parameters (e.g. 75 &#181;m spot size, 100 mW power, 100 msec duration). 
	CAUTION: Ensure operator wears all animal and laser safety personal protective equipment and pertinent laser safety signs are displayed outside of the procedure room.
	<ol>
		<li>Before using any experimental animals, find ideal parameters using a calibration, non-experimental mouse. Laser parameters will depend on the laser used. Ideal parameters are defined by the lowest laser power setting that consistently causes &#8220;bubble&#8221; formation when properly focused. See video for example of lesion with proper &#8220;bubble&#8221; formation.</li>
	</ol>
	</li>
	<li>Prepare the pre-laser station so that anesthesia, animal warmer, tissue wipes, and all eye drops (Tropicamide, Tetracaine, and artificial tears) are easily within reach to operator.
	<ol>
		<li>Ensure that animal warmer is pre-heated to correct temperature (37 &#176;C) before injecting anesthetic into first mouse in order to avoid anesthesia-induced hypothermia.</li>
	</ol>
	</li>
	<li>Place mouse stage on warmer so it can retain heat and remain warm once laser procedure begins.</li>
</ol>
2. Mouse Anesthesia and Pre-laser Preparation
<ol>
	<li>Before injecting anesthesia, inspect the eye macroscopically to ensure it has no deformities or abnormalities that diminish corneal clarity (e.g. cataract).</li>
	<li>Weigh mouse.</li>
	<li>Record weight, sex, and animal ID number.</li>
	<li>Using weight, calculate appropriate amount of anesthetic to be used based on guidelines given by institution (e.g. 100 mg/kg ketamine hydrochloride, 10 mg/kg xylazine OR tribromoethanol 250 mg/kg; for a 20 g mouse inject 0.20 ml&#160;xylazine/ketamine cocktail OR 0.25 ml of tribromoethanol). Have a table of pre-calculated dosages per weight in increments of 1&#160;g in order to reduce mathematical errors.</li>
	<li>Scruff mouse and inject anesthetic intraperitoneally based on calculations in step 2.4.</li>
	<li>Place mouse on animal warmer and wait until mouse is completely anesthetized by checking the pedal reflex via toe pinch (approximately 3-5 min).</li>
	<li>Roll up a tissue wipe and protect the mouse&#8217;s nasal area to prevent aspiration of liquid roll-off. Roll mouse on its side and place a drop (approximately 30 &#181;l) of tetracaine hydrochloride into each eye for topical anesthesia. Wait 2 min for solution to take effect.</li>
	<li>Repeat step 2.7 with one drop of topical Tropicamide for pupillary dilation. Alternatively, use phenylephrine hydrochloride (2.5%) for dilation.</li>
	<li>Wait 2 min for solutions to take effect; keep animal on warmer during this time.</li>
	<li>After appropriate time has elapsed, quickly place the mouse on the mouse stage and place the stage on chin rest of slit lamp.</li>
	<li>Turn on slit lamp to the lowest light brightness and check the degree of pupillary dilation. If pupil is not adequately dilated (approximately 2.5-3 mm), return mouse to animal warmer and wait. Alternatively, administer another drop of Tropicamide. Once eye is sufficiently dilated, proceed to laser procedure.</li>
</ol>
3. Laser Procedure
Note: Ensure other persons in the room wear protective goggles when away from laser-protected slit lamp eye-piece
<ol>
	<li>Adjust the placement of mouse on the mouse-stage, so that it is ideally positioned for visualization of optic nerve (see 3.1.2).
	<ol>
		<li>Orient the mouse on its holder so it lies horizontally, perpendicular to slit lamp beam, with the head at one side and tail at the other. Ideal mouse placement will make optic nerve visualization much easier once cover slip is applied.</li>
		<li>Slightly turn the mouse so it is at a ~170&#176; angle with the head closer to laser operator.</li>
		<li>Ensure that mouse is as close as possible to slit lamp, yet still in a position where it is stable and where the operator&#8217;s hand will have enough space for fine manipulation.</li>
	</ol>
	</li>
	<li>After the mouse is ideally positioned, place one drop of artificial tear solution on a 25 mm x 25 mm glass coverslip.
	<ol>
		<li>Place one drop of artificial tear solution on mouse&#8217;s opposite eye &#8211; this will ensure eye is hydrated and help delay cataract formation.</li>
	</ol>
	</li>
	<li>Hold corner of coverslip between thumb and pointer fingers; position so that the glass is squeezed between tips of both fingers.</li>
	<li>Gently wrap the remaining three fingers around the animal&#8217;s body for support and hand stabilization. Position hand so that the glass coverslip can be easily placed on the mouse&#8217;s eye.
	<ol>
		<li>Be sure that the wrist is stabilized on a firm surface in order to reduce hand tremor.</li>
	</ol>
	</li>
	<li>Once stable position is obtained, carefully press glass coverslip (with drop of artificial tear still adhered) onto the mouse&#8217;s eye.
	<ol>
		<li>Make sure the coverslip is positioned as perpendicular as possible to the laser beam in order to prevent laser beam scatter or reflection. The coverslip acts as a contact lens to flatten the cornea.</li>
	</ol>
	</li>
	<li>Look through slit lamp and with free hand toggle focus until retina can be visualized. The retina will have a light-yellow/red color depending on the location visualized, distinct, red vessels will be visible.</li>
	<li>Slowly and carefully manipulate mouse head and/or coverslip until visualizing the optic nerve. The optic nerve will be yellow in color with multiple vessels radiating from it.</li>
	<li>Once operator has confirmed visualization of optic nerve, turn on laser focusing beam.</li>
	<li>Once laser beam has been turned on, maneuver laser focusing beam to desired position (approximately 1 disc diameter from the optic nerve).</li>
	<li>Focus laser beam on the RPE of the eye fundus. Proper focus is achieved by having the sharpest and clearest laser beam. If aiming beam looks oval or out of focus, toggle slit lamp focus or re-position glass coverslip.</li>
	<li>Once the aiming beam is focused on RPE, initiate laser administration using the laser&#8217;s foot trigger.
	<ol>
		<li>Be sure to avoid retinal vessels to prevent intraocular hemorrhage.</li>
	</ol>
	</li>
	<li>Watch for the appearance of a bubble immediately after laser administration. The outline of the laser shot should be clear and not hazy in any way.
	<ol>
		<li>If the laser shot does not result in bubble formation, or area of impact looks hazy (cloudy appearance with ill-defined circular border vs. clear, sharply defined border of a successful impact), or if hemorrhage is seen after laser administration, do NOT include these lesions for future analysis.</li>
	</ol>
	</li>
	<li>Repeat steps 3.10-3.12 for all desired CNV positions (usually at 3, 6, 9, and 12 o&#8217;clock positions around optic nerve). If laser inductions are applied at approximately same distance from optic nerve, refocus should not be necessary. However, due to the strong curvature of the mouse eye and small variations that may exist in the retina, refocusing the beam may be necessary between consecutive laser administrations.</li>
	<li>Record in a notebook the location and result of each laser shot administration and result (successful, hazy, hemorrhage, etc.) of each administered shot for the eye. Be sure to place the laser in stand-by mode when not in use.</li>
	<li>Repeat 3.1-3.14 for the mouse&#8217;s other eye, if needed, using the opposite hand for stabilization and a new coverslip.</li>
	<li>After all desired laser shots are administered, turn off laser and slit-lamp.</li>
	<li>Discard coverslip and place mouse on warmer for recovery from anesthesia. Macroscopically inspect eye for any injury and place a drop of artificial tear solution to keep the eye hydrated and potentially prevent future cataract development. Once mouse recovers from anesthesia, return to cage.</li>
</ol>
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td></td>
			<td>AVERTIN</td>
			<td>AVERTIN</td>
			<td>AVERTIN</td>
			<td>XYL/KET</td>
			<td>XYL/KET</td>
		</tr>
		<tr>
			<td>Mouse Weight (g)</td>
			<td>Dose (mg/kg)</td>
			<td>Solution Concentration (mg/ml)</td>
			<td>Anesthetic Dose (ml)</td>
			<td>Dose (mg/kg)</td>
			<td>Anesthetic Dose (ml)</td>
		</tr>
		<tr>
			<td>15</td>
			<td>250</td>
			<td>20</td>
			<td>0.1875</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.15</td>
		</tr>
		<tr>
			<td>16</td>
			<td>250</td>
			<td>20</td>
			<td>0.2</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.16</td>
		</tr>
		<tr>
			<td>17</td>
			<td>250</td>
			<td>20</td>
			<td>0.2125</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.17</td>
		</tr>
		<tr>
			<td>18</td>
			<td>250</td>
			<td>20</td>
			<td>0.225</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.18</td>
		</tr>
		<tr>
			<td>19</td>
			<td>250</td>
			<td>20</td>
			<td>0.2375</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.19</td>
		</tr>
		<tr>
			<td>20</td>
			<td>250</td>
			<td>20</td>
			<td>0.25</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.2</td>
		</tr>
		<tr>
			<td>21</td>
			<td>250</td>
			<td>20</td>
			<td>0.2625</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.21</td>
		</tr>
		<tr>
			<td>22</td>
			<td>250</td>
			<td>20</td>
			<td>0.275</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.22</td>
		</tr>
		<tr>
			<td>23</td>
			<td>250</td>
			<td>20</td>
			<td>0.2875</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.23</td>
		</tr>
		<tr>
			<td>24</td>
			<td>250</td>
			<td>20</td>
			<td>0.3</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.24</td>
		</tr>
		<tr>
			<td>25</td>
			<td>250</td>
			<td>20</td>
			<td>0.3125</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.25</td>
		</tr>
		<tr>
			<td>26</td>
			<td>250</td>
			<td>20</td>
			<td>0.325</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.26</td>
		</tr>
		<tr>
			<td>27</td>
			<td>250</td>
			<td>20</td>
			<td>0.3375</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.27</td>
		</tr>
		<tr>
			<td>28</td>
			<td>250</td>
			<td>20</td>
			<td>0.35</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.28</td>
		</tr>
		<tr>
			<td>29</td>
			<td>250</td>
			<td>20</td>
			<td>0.3625</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.29</td>
		</tr>
		<tr>
			<td>30</td>
			<td>250</td>
			<td>20</td>
			<td>0.375</td>
			<td>100 mg/kg ketamine; 10 mg/kg xylazine</td>
			<td>0.3</td>
		</tr>
	</tbody>
</table>
Table 1: XyIKet Dosage Chart.

<ol>
	<li>Bressler, N. M., Bressler, S. B., Fine, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Age+related+macular+degeneration.">Age related macular degeneration.</a> Surv Ophthalmol. 32, 375-413 (1988).</li><li>Congdon, N., 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=Causes+and+Prevalence+of+Visual+Impairment+Among+Adults+in+the+United+States.">Causes and Prevalence of Visual Impairment Among Adults in the United States.</a> Arch Ophthalmol. 122, 477-485 (2004).</li><li>Jager, R. D., Mieler, W. F., Miller, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Age-Related+Macular+Degeneration.">Age-Related Macular Degeneration.</a> NEJM. 358, 2606-2617 (2008).</li><li>Poor, S. H., 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=Reliability+of+the+Mouse+Model+of+Choroidal+Neovascularization+Induced+by+Laser+Photocoagulation.">Reliability of the Mouse Model of Choroidal Neovascularization Induced by Laser Photocoagulation.</a> IOVS. 55, 6525-6534 (2014).</li><li>Ryan, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+development+of+an+experimental+model+of+subretinal+neovascularization+in+disciform+macular+degeneration.">The development of an experimental model of subretinal neovascularization in disciform macular degeneration.</a> Trans Am Ophthalmol Soc. 77, 707-745 (1979).</li><li>Miller, H., Miller, B., Ishibashi, T., Ryan, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pathogenesis+of+laser+induced+choroidal+subretinal+neovascularization.">Pathogenesis of laser induced choroidal subretinal neovascularization.</a> IOVS. 31, 899-908 (1990).</li><li>Pennesi, M. E., Neuringer, M., Courtney, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+age+related+macular+degeneration.">Animal models of age related macular degeneration.</a> Molr Aspects Mede. 33, 487-509 (2012).</li><li>Grossniklaus, H. E., Kang, S. J., Berglin, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+Models+of+Choroidal+and+Retinal+Neovascularization.">Animal Models of Choroidal and Retinal Neovascularization.</a> Prog Retin Eye Res. 29, 500-519 (2010).</li><li>Zeiss, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=REVIEW+PAPER:+Animals+as+Models+of+Age+Related+Macular+Degeneration+An+Imperfect+Measure+of+the+Truth.">REVIEW PAPER: Animals as Models of Age Related Macular Degeneration An Imperfect Measure of the Truth.</a> Vet Pathol Online. 47, 396-413 (2010).</li><li>Tobe, T., 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=Targeted+Disruption+of+the+FGF2+Gene+Does+Not+Prevent+Choroidal+Neovascularization+in+a+Murine+Model.">Targeted Disruption of the FGF2 Gene Does Not Prevent Choroidal Neovascularization in a Murine Model.</a> Am J Pathol. 153, 1641-1646 (1998).</li><li>He, L., Marneros, A. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Macrophages+Are+Essential+for+the+Early+Wound+Healing+Response+and+the+Formation+of+a+Fibrovascular+Scar.">Macrophages Are Essential for the Early Wound Healing Response and the Formation of a Fibrovascular Scar.</a> Am J Pathol. 182, 2407-2417 (2013).</li><li>Hoerster, 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=In+vivo+and+ex+vivo+characterization+of+laser+induced+choroidal+neovascularization+variability+in+mice.">In vivo and ex vivo characterization of laser induced choroidal neovascularization variability in mice.</a> Graefes Arch Clin Exp Ophthalmol. 250, 1579-1586 (2012).</li><li>Jawad, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Role+of+Macrophage+Class+A+Scavenger+Receptors+in+a+Laser+Induced+Murine+Choroidal+Neovascularization+Model.">The Role of Macrophage Class A Scavenger Receptors in a Laser Induced Murine Choroidal Neovascularization Model.</a> IOVS. 54, 5959-5970 (2013).</li><li>Lambert, V., 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=Laser+induced+choroidal+neovascularization+model+to+study+age+related+macular+degeneration+in+mice.">Laser induced choroidal neovascularization model to study age related macular degeneration in mice.</a> Nat Protocols. 8, 2197-2211 (2013).</li><li>Sakurai, E., Anand, A., Ambati, B. K., van Rooijen, N., Ambati, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Macrophage+Depletion+Inhibits+Experimental+Choroidal+Neovascularization.">Macrophage Depletion Inhibits Experimental Choroidal Neovascularization.</a> IOVS. 44, 3578-3585 (2003).</li><li>Espinosa Heidmann, D. G., 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=Macrophage+Depletion+Diminishes+Lesion+Size+and+Severity+in+Experimental+Choroidal+Neovascularization.">Macrophage Depletion Diminishes Lesion Size and Severity in Experimental Choroidal Neovascularization.</a> IOVS. 44, 3586-3592 (2003).</li><li>Giani, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+Vivo+Evaluation+of+Laser+Induced+Choroidal+Neovascularization+Using+Spectral+Domain+Optical+Coherence+Tomography.">In Vivo Evaluation of Laser Induced Choroidal Neovascularization Using Spectral Domain Optical Coherence Tomography.</a> IOVS. 52, 3880-3887 (2011).</li><li>Kwak, N., Okamoto, N., Wood, J. M., Campochiaro, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=VEGF+Is+Major+Stimulator+in+Model+of+Choroidal+Neovascularization.">VEGF Is Major Stimulator in Model of Choroidal Neovascularization.</a> IOVS. 41, 3158-3164 (2000).</li><li>Reich, S. J., 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=Small+interfering+RNA+(siRNA)+targeting+VEGF+effectively+inhibits+ocular+neovascularization+in+a+mouse+model.">Small interfering RNA (siRNA) targeting VEGF effectively inhibits ocular neovascularization in a mouse model.</a> Molr Vis. 9, 210-216 (2003).</li><li>Baffi, J., Byrnes, G., Chan, C. C., Csaky, K. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Choroidal+Neovascularization+in+the+Rat+Induced+by+Adenovirus+Mediated+Expression+of+Vascular+Endothelial+Growth+Factor.">Choroidal Neovascularization in the Rat Induced by Adenovirus Mediated Expression of Vascular Endothelial Growth Factor.</a> IOVS. 41, 3582-3589 (2000).</li><li>Claybon, A., Bishop, A. J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissection+of+a+Mouse+Eye+for+a+Whole+Mount+of+the+Retinal+Pigment+Epithelium.">Dissection of a Mouse Eye for a Whole Mount of the Retinal Pigment Epithelium.</a> JoVE. , 2563 (2011).</li><li>Espinosa-Heidmann, D. G., 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=Age+as+an+Independent+Risk+Factor+for+Severity+of+Experimental+Choroidal+Neovascularization.">Age as an Independent Risk Factor for Severity of Experimental Choroidal Neovascularization.</a> IOVS. 43, 1567-1573 (2002).</li><li>Zhu, Y., 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=Improvement+and+Optimization+of+Standards+for+a+Preclinical+Animal+Test+Model+of+Laser+Induced+Choroidal+Neovascularization.">Improvement and Optimization of Standards for a Preclinical Animal Test Model of Laser Induced Choroidal Neovascularization.</a> PLoS ONE. 9, e94743 (2014).</li><li>Weinstock, M., Stewart, H. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Occurrence+in+rodents+of+reversible+drug+induced+opacities+of+the+lens.">Occurrence in rodents of reversible drug induced opacities of the lens.</a> British J Ophthalmol. 45, 408-414 (1961).</li><li>Ridder III, W. H., Nusinowitz, S., Heckenlively, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Causes+of+Cataract+Development+in+Anesthetized+Mice.">Causes of Cataract Development in Anesthetized Mice.</a> Exp Eye Res. 75, 365-370 (2002).</li></ol>

The authors declare they have no competing financial interests.

There are multiple factors that can affect laser delivery and resultant CNV lesion development after successful laser-induction. These factors should be controlled for and standardized in order to have the most reliable results. The most pertinent of these factors are mouse selection (genotype, age, and sex), anesthetic selection, and laser settings.
The specific mouse model used can have a significant effect on the course of CNV development. The most widely used genotype is the C57BL/6 mouse....

Age-related macular degeneration (AMD) is one of the leading causes of blindness in individuals over the age of 501-3. AMD can be classified into two forms: atrophic (&#8220;dry&#8221;) AMD and neovascular (&#8220;wet&#8221;) AMD. The former is characterized by geographic atrophy of the retinal pigment epithelium (RPE), choriocapillaris, and photoreceptors, while the latter is characterized by the invasion of abnormal vessels from the choroid into the outer retinal layers causing leakage, hemorrhage, and fibrosis, and ultimately leading to blindness1,2. Of the two forms, neovascular AMD accounts for the majority of vision loss1. Fortunately, this form has numerous effective pharmacological management options, whereas its atrophic counterpart currently has no proven medical treatments3. Moreover, because the neovascular form has been easily re-capitulated in an animal model, it has been more widely accessible to basic AMD research exploring the underlying pathological mechanisms in order to develop novel therapies4.
The first animal model of experimental choroidal neovascularization (CNV) was developed by Ryan et al. in non-human primates5. This model induced rupture of Bruch&#8217;s membrane via laser photocoagulation, which caused a local inflammatory response resulting in angiogenesis similar to that seen in neovascular AMD. The histopathological progression of angiogenesis post-laser induction was found to mimic neovascular AMD, which confirmed the model&#8217;s validity6. Non-human primates offer the most similar anatomy to humans, but unfortunately, are expensive to maintain, cannot be easily genetically manipulated, and have a slow time course of disease progression7. Contrastingly, rodent models are much more cost-effective to maintain, can be genetically manipulated with relative ease, and have a much faster course of disease progression (experiments can be conducted on a time scale of weeks versus months). These experiments should only be conducted in pigmented rodents as it is very difficult to visualize in albino animals.
The mouse laser-induced CNV model, first developed by the Campochiaro group in the late 90&#8217;s10, has grown to be the dominant animal model in the majority of recent studies11-16. Due to the complex and still unclear pathogenesis of CNV, the laser model has been applied in all aspects of wet AMD research ranging from studying the molecular mechanisms driving angiogenesis to evaluating new treatment modalities for future human use. For example, Sakurai et al. and Espinosa-Heidmann et al. used the laser model to investigate the effect of macrophages on the development of CNV using transgenic mice and pharmacological depletion treatments15, 16. Giani et al. and Hoerster et al. used optical-coherence tomography (OCT) to image the laser-induced CNV in an effort to characterize the progression of CNV and compare the histopathologic findings to the findings seen on OCT imaging12,17. Finally, studies involving intravitreal injection of anti-angiogenic agents have been used as pre-requisites for human trials and were vital in developing the first generation of anti-VEGF agents used in management of neovascular AMD today10,18,19.
Alternative models for experimental CNV utilize surgical methods to induce CNV. This procedure involves injecting pro-angiogenic substances (e.g. recombinant viral vectors overexpressing VEGF, subretinal injection of RPE cells and/or polystyrene beads) to mimic the increased VEGF expression seen in neovascular AMD, with the goal of causing angiogenesis8,20. However, this method yields a drastically lower incidence of neovascularization; these studies showed that CNV in C57/BL6 mice occurs in 31% of injections versus the ~70% success rate seen in the laser photocoagulation method in the same strain of mice8,14. For these reasons, and given the advantages of using rodents versus non-human primates, the mouse model of laser-induced CNV has become the standard animal model of CNV for most neovascular AMD study experiments8.
The mouse eye is a miniscule, delicate tissue to work with. Maneuvering of the eye to visualize the retina is difficult and requires much practice until mastery is achieved. This task is complicated by the fact that it must be learned with the dominant and non-dominant hand. Furthermore, after the fine movements required to visualize the retina have been learned, the coordination between both hands and the foot pedal operating the laser are important. In this paper, we sought to distill the challenges of learning all of the physical manipulations involved in the laser-induced CNV procedure into a guide that would help operators achieve rapid success with this model.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>532 nm (green) argon ophthalmic laser</td>
			<td>IRIDEX</td>
			<td>GLx</td>
			<td>any ophthalmic 532 nm (green) argon laser can be used</td>
		</tr>
		<tr>
			<td>slit lamp</td>
			<td>Carl Zeiss</td>
			<td>30SL-M</td>
			<td>any slit lamp can be used as long as it is compatible with the laser</td>
		</tr>
		<tr>
			<td>tribromoethanol</td>
			<td>Sigma</td>
			<td>T48402-25G</td>
			<td>used to make anesthetic</td>
		</tr>
		<tr>
			<td>tert-amyl alcohol</td>
			<td>Sigma</td>
			<td>152463-1L</td>
			<td>used to make anesthetic</td>
		</tr>
		<tr>
			<td>amber glass vials + septa</td>
			<td>Wheaton</td>
			<td>WH-223696</td>
			<td>tribromoethanol storage</td>
		</tr>
		<tr>
			<td>tissue wipes</td>
			<td>VWR</td>
			<td>82003-820</td>
			<td>miscellaneous&#160;</td>
		</tr>
		<tr>
			<td>1% tropicamide</td>
			<td>Falcon Pharmaceuticals</td>
			<td>RXD2974251</td>
			<td>pupillary dilation</td>
		</tr>
		<tr>
			<td>0.5% tetracaine hydrochloride</td>
			<td>Alcon</td>
			<td>&#160;0065-0741-12</td>
			<td>topical anesthesia</td>
		</tr>
		<tr>
			<td>artificial tears</td>
			<td>Alcon</td>
			<td>&#160;58768-788-25</td>
			<td>hydration</td>
		</tr>
		<tr>
			<td>heat therapy pump (for animal warming)</td>
			<td>Kent Scientific</td>
			<td>HTP-1500</td>
			<td>used to maintain animal body temp</td>
		</tr>
		<tr>
			<td>warming pad</td>
			<td>Kent Scientific</td>
			<td>TPZ-0510EA</td>
			<td>maintains animal body temperature</td>
		</tr>
		<tr>
			<td>30 gauge insulin needles</td>
			<td>BD</td>
			<td>328418</td>
			<td>IP anesthesia injection</td>
		</tr>
		<tr>
			<td>scale</td>
			<td>American Weigh Scale</td>
			<td>AWS-1KG-BLK</td>
			<td>mouse weighing</td>
		</tr>
		<tr>
			<td>cover slip (25 mm x 25 mm)</td>
			<td>VWR</td>
			<td>48366089</td>
			<td>flatten cornea to visualize mouse retina</td>
		</tr>
		<tr>
			<td>xylazine</td>
			<td>obtained from institution</td>
			<td>obtained from institution</td>
			<td>anesthesia</td>
		</tr>
		<tr>
			<td>ketamine</td>
			<td>obtained from institution</td>
			<td>obtained from institution</td>
			<td>anesthesia</td>
		</tr>
		<tr>
			<td>Volocity</td>
			<td>PerkinElmer</td>
			<td></td>
			<td>used for volumetric re-construction</td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>National Institutes of Health</td>
			<td></td>
			<td>used for image analysis</td>
		</tr>
	</tbody>
</table>

a mouse model for laser-induced choroidal neovascularization

The mouse laser-induced choroidal neovascularization (CNV) model has been a crucial mainstay model for neovascular age-related macular degeneration (AMD) research. By administering targeted laser injury to the RPE and Bruch&#8217;s membrane, the procedure induces angiogenesis, modeling the hallmark pathology observed in neovascular AMD. 
First developed in non-human primates, the laser-induced CNV model has come to be implemented into many other species, the most recent of which being the mouse. Mouse experiments are advantageously more cost-effective, experiments can be executed on a much faster timeline, and they allow the use of various transgenic models. The miniature size of the mouse eye, however, poses a particular challenge when performing the procedure. Manipulation of the eye to visualize the retina requires practice of fine dexterity skills as well as simultaneous hand-eye-foot coordination to operate the laser. However, once mastered, the model can be applied to study many aspects of neovascular AMD such as molecular mechanisms, the effect of genetic manipulations, and drug treatment effects. 
The laser-induced CNV model, though useful, is not a perfect model of the disease. The wild-type mouse eye is otherwise healthy, and the chorio-retinal environment does not mimic the pathologic changes in human AMD. Furthermore, injury-induced angiogenesis does not reflect the same pathways as angiogenesis occurring in an age-related and chronic disease state as in AMD.
Despite its shortcomings, the laser-induced CNV model is one of the best methods currently available to study the debilitating pathology of neovascular AMD. Its implementation has led to a deeper understanding of the pathogenesis of AMD, as well as contributing to the development of many of the AMD therapies currently available.

Quantification of CNV lesions can be performed through analysis of flat-mounted choroids using immunofluorescence staining to label the CNV vessels. The two most frequently employed methods of tissue preparation are FITC-dextran labeling, done via perfusion immediately before animal sacrifice, or post-mortem immuno-staining with an endothelial cell marker. Both of these methods have been described previously in detail13,14,21; Figures 1 and 2 show examples of each, respectivel...

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A Mouse Model for Laser-induced Choroidal Neovascularization

Here, we present the mouse laser-induced choroidal neovascularization (CNV) protocol, an experimental model that re-creates the vascular hallmarks of neovascular age-related macular degeneration (AMD). Once mastered, it can reliably and effectively induce CNV as a model system to test various experimental measures.

The mouse laser-induced choroidal neovascularization (CNV) model has been a crucial mainstay model for neovascular age-related macular degeneration (AMD) ...

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17.5K vues.  Northwestern University Feinberg School of Medicine. Here, we present the mouse laser-induced choroidal neovascularization (CNV) protocol, an experimental model that re-creates the vascular hallmarks of neovascular age-related macular degeneration (AMD). Once mastered, it can reliably and effectively induce CNV as a model system to test various experimental measures.

Vidéo: A Mouse Model for Laser-induced Choroidal Neovascularization

A Laser-induced Mouse Model of Chronic Ocular Hypertension to Characterize Visual Defects

Glaucoma, frequently associated with elevated intraocular pressure (IOP), is one of the leading causes of blindness. We sought to establish a mouse model of ocular hypertension to mimic human high-tension glaucoma. Here laser illumination is applied to the corneal limbus to photocoagulate the aqueous outflow, inducing angle closure. The changes of IOP are monitored using a rebound tonometer before and after the laser treatment. An optomotor behavioral test is used to measure corresponding changes in visual capacity. The representative result from one mouse which developed sustained IOP elevation after laser illumination is shown. A decreased visual acuity and contrast sensitivity is observed in this ocular hypertensive mouse. Together, our study introduces a valuable model system to investigate neuronal degeneration and the underlying molecular mechanisms in glaucomatous mice.

Chronic ocular hypertension is induced using laser photocoagulation of the trabecular meshwork in mouse eyes. The intraocular pressure (IOP) is elevated for several months after laser treatment. The decrease of visual acuity and contrast sensitivity of experimental animals are monitored using the optomotor test.

Glaucoma, frequently associated with elevated intraocular pressure (IOP), is one of the leading causes of blindness. We sought to establish a mouse model ...

Detecting Abnormalities in Choroidal Vasculature in a Mouse Model of Age-related Macular Degeneration by Time-course Indocyanine Green Angiography

Indocyanine Green&#160;Angiography (or ICGA) is a technique performed by ophthalmologists to diagnose abnormalities of the choroidal and retinal vasculature of various eye diseases such as age-related macular degeneration (AMD). ICGA is especially useful to image the posterior choroidal vasculature of the eye due to its capability of penetrating through the pigmented layer with its infrared spectrum. ICGA time course can be divided into early, middle, and late phases. The three phases provide valuable information on the pathology of eye problems. Although time-course ICGA by intravenous (IV) injection is widely used in the clinic for the diagnosis and management of choroid problems, ICGA by intraperitoneal injection (IP) is commonly used in animal research. Here we demonstrated the technique to obtain high-resolution ICGA time-course images in mice by tail-vein injection and confocal scanning laser ophthalmoscopy. We used this technique to image the choroidal lesions in a mouse model of age-related macular degeneration. Although it is much easier to introduce ICG to the mouse vasculature by IP, our data indicate that it is difficult to obtain reproducible ICGA time course images by IP-ICGA. In contrast, ICGA via tail vein injection provides high quality ICGA time-course images comparable to human studies. In addition, we showed that ICGA performed on albino mice gives clearer pictures of choroidal vessels than that performed on pigmented mice. We suggest that time-course IV-ICGA should become a standard practice in AMD research based on animal models.

Indocyanine Green Angiography (or ICGA) performed by tail vein injection provides high quality ICGA time course images to characterize abnormalities in mouse choroid.

Indocyanine Green&#160;Angiography (or ICGA) is a technique performed by ophthalmologists to diagnose abnormalities of the choroidal and retinal ...

An Alkali-burn Injury Model of Corneal Neovascularization in the Mouse

Under normal conditions, the cornea is avascular, and this transparency is essential for maintaining good visual acuity. Neovascularization (NV) of the cornea, which can be caused by trauma, keratoplasty or infectious disease, breaks down the so called &lsquo;angiogenic privilege' of the cornea and forms the basis of multiple visual pathologies that may even lead to blindness. Although there are several treatment options available, the fundamental medical need presented by corneal neovascular pathologies remains unmet. In order to develop safe, effective, and targeted therapies, a reliable model of corneal NV and pharmacological intervention is required. Here, we describe an alkali-burn injury corneal neovascularization model in the mouse. This protocol provides a method for the application of a controlled alkali-burn injury to the cornea, administration of a pharmacological compound of interest, and visualization of the result. This method could prove instrumental for studying the mechanisms and opportunities for intervention in corneal NV and other neovascular disorders.

Neovascularization (NV) of the cornea can complicate multiple visual pathologies. Utilizing a controlled, alkali-burn injury model, a quantifiable level of corneal NV can be produced for mechanistic study of corneal NV and evaluation of potential therapies for neovascular disorders.

Under normal conditions, the cornea is avascular, and this transparency is essential for maintaining good visual acuity. Neovascularization (NV) of the ...

A Mouse Fetal Skin Model of Scarless Wound Repair

Early in utero, but not in postnatal life, cutaneous wounds undergo regeneration and heal without formation of a scar. Scarless fetal wound healing occurs across species but is age dependent. The transition from a scarless to scarring phenotype occurs in the third trimester of pregnancy in humans and around embryonic day 18 (E18) in mice. However, this varies with the size of the wound with larger defects generating a scar at an earlier gestational age. The emergence of lineage tracing and other genetic tools in the mouse has opened promising new avenues for investigation of fetal scarless wound healing. However, given the inherently high rates of morbidity and premature uterine contraction associated with fetal surgery, investigations of fetal scarless wound healing in vivo require a precise and reproducible surgical model. Here we detail a reliable model of fetal scarless wound healing in the dorsum of E16.5 (scarless) and E18.5 (scarring) mouse embryos.

During mammalian development, early gestational skin wounds heal without a scar. Here we detail a reliable and reproducible model of fetal scarless wound healing in the cutaneous dorsum of E16.5 (scarless) and E18.5 (scarring) mouse embryos.

Early in utero, but not in postnatal life, cutaneous wounds undergo regeneration and heal without formation of a scar. Scarless fetal wound healing occurs ...

A Neonatal Mouse Spinal Cord Compression Injury Model

Spinal cord injury (SCI) typically causes devastating neurological deficits, particularly through damage to fibers descending from the brain to the spinal cord. A major current area of research is focused on the mechanisms of adaptive plasticity that underlie spontaneous or induced functional recovery following SCI. Spontaneous functional recovery is reported to be greater early in life, raising interesting questions about how adaptive plasticity changes as the spinal cord develops. To facilitate investigation of this dynamic, we have developed a SCI model in the neonatal mouse. The model has relevance for pediatric SCI, which is too little studied. Because neural plasticity in the adult involves some of the same mechanisms as neural plasticity in early life1, this model may potentially have some relevance also for adult SCI. Here we describe the entire procedure for generating a reproducible spinal cord compression (SCC) injury in the neonatal mouse as early as postnatal (P) day 1. SCC is achieved by performing a laminectomy at a given spinal level (here described at thoracic levels 9-11) and then using a modified Yasargil aneurysm mini-clip to rapidly compress and decompress the spinal cord. As previously described, the injured neonatal mice can be tested for behavioral deficits or sacrificed for ex vivo physiological analysis of synaptic connectivity using electrophysiological and high-throughput optical recording techniques1. Earlier and ongoing studies using behavioral and physiological assessment have demonstrated a dramatic, acute impairment of hindlimb motility followed by a complete functional recovery within 2 weeks, and the first evidence of changes in functional circuitry at the level of identified descending synaptic connections1.

This article describes a method for generating a reproducible spinal cord compression injury (SCI) in the neonatal mouse. The model provides an advantageous platform for studying mechanisms of adaptive plasticity that underlie spontaneous functional recovery.

Spinal cord injury (SCI) typically causes devastating neurological deficits, particularly through damage to fibers descending from the brain to the spinal ...

Puncture-Induced Iris Neovascularization as a Mouse Model of Rubeosis Iridis

We describe a model of puncture-induced iris neovascularization as a general model for noninvasive evaluation of angiogenesis. The model is also relevant for targeting neovascular glaucoma, a sight-threatening complication of diabetic retinopathy. This method is based on the induction of iris vascular response by a series of self-sealing uveal punctures on BALB/c mice and takes advantage of the postpartum maturation of mouse ocular vasculature. Mouse pups undergo uveal punctures from postnatal day 12.5, when the pups naturally open their eyes, until postnatal day 24.5. Due to the transparency of the cornea, iris vasculature can be analyzed easily through time by noninvasive in vivo methods. Furthermore, the semitransparent iris of BALB/c mice can be flatmounted for detailed immunohistologic analysis with minimal non-specific background staining. In this model, angiogenesis is mainly driven by the inflammatory and plasminogen activating systems. The puncture-induced model is the first to induce iris neovascularization in small rodents, and has the advantage of allowing direct noninvasive in vivo analysis of the angiogenic process. Moreover, the model can be combined with angiogenic modulating substances, which highlights its potential in the study of angiogenesis with an in vivo perspective.

Iris neovascularization, a common complication of ischemic retinal disease, may lead to sight-threatening neovascular glaucoma. Here, we describe a murine protocol for inducing experimental iris neovascularization that may be used for noninvasive evaluation of angiogenesis-modulating substances.

We describe a model of puncture-induced iris neovascularization as a general model for noninvasive evaluation of angiogenesis. The model is also relevant ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...

A Novel Approach to Monitoring Graft Neovascularization in the Human Gingiva

Laser speckle contrast imaging (LSCI) is a novel method for measuring superficial blood perfusion over large areas. Since it is non-invasive and avoids direct contact with the measured area, it is suitable for monitoring blood flow changes during wound healing in human patients. Vestibuloplasty is periodontal surgery to the oral vestibule, aiming to restore vestibular depth with simultaneous enlargement of the keratinized gingiva. In this special clinical case, a split thickness flap was elevated at the first upper premolar and a xenogenic collagen matrix was adapted to the resulting recipient bed. LSCI was used to monitor the re- and neovascularization of the graft and the surrounding mucosa for one year. A protocol is introduced for the correct adjustment of microcirculation measurement in the oral mucosa, highlighting difficulties and possible failures.
The clinical case study presented demonstrated that &#8212; following the appropriate protocol &#8212; LSCI is a suitable and reliable method for following up microcirculation in a healing wound in the human oral mucosa and gives useful information on graft integration.

This study introduces a protocol for measuring microcirculation in human oral mucosa by laser speckle contrast imaging. The monitoring of wound healing after vestibuloplasty combined with a xenogenic collagen graft is presented on a clinical case.

Laser speckle contrast imaging (LSCI) is a novel method for measuring superficial blood perfusion over large areas. Since it is non-invasive and avoids ...

A Mouse Model for Chronic Pancreatitis via Bile Duct TNBS Infusion

Chronic pancreatitis (CP) is a complex disease involving pancreatic inflammation and fibrosis, glandular atrophy, abdominal pain and other symptoms. Several rodent models have been developed to study CP, of which the bile duct 2,4,6 -trinitrobenzene sulfonic acid (TNBS) infusion model replicates the features of neuropathic pain seen in CP. However, bile duct drug infusion in mice is technically challenging. This protocol demonstrates the procedure of bile duct TNBS infusion for generation of a CP mouse model. TNBS was infused into the pancreas through the ampulla of Vater in the duodenum. This protocol optimized drug volume, surgical techniques, and drug handling during the procedure. TNBS-treated mice showed features of CP as reflected by bodyweight and pancreas weight reductions, changes in pain-associated behaviors, and abnormal pancreatic morphology. With these improvements, mortality associated with TNBS injection was minimal. This procedure is not only critical in generating pancreatic disease models but is also useful in local pancreatic drug delivery.

Chronic pancreatitis (CP) is a disease characterized by inflammation and fibrosis of the pancreas, often associated with intractable abdominal pain. This article focuses on refining the technique to generate a mouse model of CP via bile duct infusion with 2,4,6 -trinitrobenzene sulfonic acid (TNBS).

Chronic pancreatitis (CP) is a complex disease involving pancreatic inflammation and fibrosis, glandular atrophy, abdominal pain and other symptoms. ...

Controlled Semi-Automated Laser-Induced Injuries for Studying Spinal Cord Regeneration in Zebrafish Larvae

Zebrafish larvae possess a fully functional central nervous system (CNS) with a high regenerative capacity only a few days after fertilization. This makes this animal model very useful for studying spinal cord injury and regeneration. The standard protocol for inducing such lesions is to transect the dorsal part of the trunk manually. However, this technique requires extensive training and damages additional tissues. A protocol was developed for laser-induced lesions to circumvent these limitations, allowing for high reproducibility and completeness of spinal cord transection over many animals and between different sessions, even for an untrained operator. Furthermore, tissue damage is mainly limited to the spinal cord itself, reducing confounding effects from injuring different tissues, e.g., skin, muscle, and CNS. Moreover, hemi-lesions of the spinal cord are possible. Improved preservation of tissue integrity after laser injury facilitates further dissections needed for additional analyses, such as electrophysiology. Hence, this method offers precise control of the injury extent that is unachievable manually. This allows for new experimental paradigms in this powerful model in the future.

The present protocol describes a method to induce tissue-specific and highly reproducible injuries in zebrafish larvae using a laser lesion system combined with an automated microfluidic platform for larvae handling.

Zebrafish larvae possess a fully functional central nervous system (CNS) with a high regenerative capacity only a few days after fertilization. This makes ...