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.

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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. The vendor from which the animals are obtained can affect the resultant CNV size. Poor et al. demonstrated significant differences in final CNV lesion size in C57BL/6 mice from Jackson, Charles Rivers, and Taconic laboratories, with mice from Taconic developing significantly larger CNV than the other two vendors4. Therefore, purchasing from one company and using all mice from that same vendor will help to minimize external differences in CNV lesion size. Age and sex of the mouse has also been shown to be an important factor to consider when designing the experiment. Female mice develop more CNV than male mice of the same age, and older mice of both sexes develop more CNV than younger mice 22,23. Depending on the experimental parameters, operators should keep these factors in mind. For example, if operators are using the laser-CNV procedure to elucidate mechanistic and drug-development purposes, both sexes at different ages should be used to define mechanisms that are specific to the sexes and those that are not.
Proper anesthesia is a critical step for this procedure, especially during the learning process. Most IACUC approved regimens can induce anesthesia for at least 15-30 min, which is more than enough time to deliver 4-5 laser shots to each eye once the procedure has been mastered. However, if too much time elapses, anesthesia can lead to reversible lens opacification, rendering the eye optically impervious for experimental CNV24. The two main protocols used to induce anesthesia are a xylazine/ketamine cocktail or tribromoethanol (TBE) delivered intraperitoneally. Although some reports posit the advantages of TBE to xylazine/ketamine in terms of cataract formation, both eventually result in the development of a cloudy lens if the operator does not work quickly14. One way to extend the time before cataract development is to maintain hydration of the eye, as mentioned in the detailed protocol, through application of artificial tears25. This method, regardless of anesthetic substance used, should help delay cataract formation and give the operator more time to complete the procedure.
Laser settings can be a major factor in the reliable induction of CNV. Calibrating the laser power and duration are an important preliminary step before conducting the procedure on experimental animals. Laser shots that cause hemorrhage or are out of focus without bubble formation should be excluded from calculation, as this disrupts the CNV development process. If multiple vessels are ruptured causing diffuse hemorrhage, the entire eye should be excluded. Ideally, the lowest power and shortest duration of laser that consistently ruptures Bruch&#8217;s membrane causing bubble formation and a lesion with clear demarcation should be used4. Protocols experimenting with different power settings have demonstrated that increased power and duration lead to excessive tissue damage and subsequently lower rates of CNV formation4, 14,17. Furthermore, location of the laser injury can also affect CNV development. Impacts delivered approximately 1 disc diameter (DD) from the optic disk yielded significantly larger area CNV volumes than those delivered &#60;1 DD or 2 or more DDs away23. Depending on the post-laser analysis, lesion location may or may not be crucial. For example, if an OCT image is to be obtained of all lesions, central location around the optic nerve is vital. Contrastingly, if lesions are to be analyzed via flat mount, lesion location is not as crucial. Finally, ensure that last shots are not placed too close to one another or else two lesions may &#8220;grow&#8221; together.
The CNV assay is a robust method to experimentally study neovascular AMD. The angiogenesis resulting from laser induction is similar in location and overall appearance to the angiogenesis observed in human AMD patients. However, it is far from a perfect model. Unlike the human eye, mice do not have a defined macula, and the acute injury-related angiogenesis seen in the laser-induced CNV model fundamentally differs from the genetically influenced, chronic pathology of AMD7,8,14. The mouse retinal environment is healthy and the resultant angiogenesis occurs as a reaction to the trauma caused by the laser impact, rather than genetic/environmental factors and age, as in human AMD. Contrastingly, the human retinal environment in AMD is in a state of chronic inflammation, during which abnormalities in VEGF expression and other cytokines cause CNV3. Clearly, the neovascular development in these two environments is markedly different.
In conclusion, the laser-induced CNV protocol is one that is initially difficult to perform but ultimately rewarding to master. The fine dexterity needed to hold the mouse and coverslip, as well as manipulate the coverslip and mouse head to visualize the optic nerve are exercises that require patience and practice, especially since they need to be performed well using either hands of the operator. However, once the technique is learned it can be an efficient way to implement experimental CNV and can be applied to most aspects of neovascular AMD research.

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><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&amp;nbsp;</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>&amp;nbsp;0065-0741-12</td><td>topical anesthesia</td></tr><tr><td>artificial tears</td><td>Alcon</td><td>&amp;nbsp;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 G 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, respectively. After confocal microscopy image acquisition, either area (2-Dimensions) or volume (3-Dimensions) can be calculated and visualized with ImageJ software or Volocity. In addition to quantification, OCT imaging can be used to visualize the CNV lesion in vivo. An example of a cross sectional image of the retina with resultant CNV is shown in Figure 3.
<img alt="Figure 1" src="/files/ftp_upload/53502/53502fig1.jpg" /> 
Figure 1: CNV Perfusion Staining &#38; Area (2D) Calculation Example (25X). (A) FITC dextran perfused CNV lesion. (B) ImageJ area quantification method via thresholding. Mouse Strain: C57BL/6J. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/53502/53502fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53502/53502fig2.jpg" /> 
Figure 2: CNV Immunostaining &#38; Volume (3D) Calculation Example (25X). (A) Isolectin GS-IB4 stained CNV lesion. (B) 3D reconstruction of CNV lesion en face view. (C) 3D reconstruction side view (B and C one tile width = 35 &#181;m). Mouse Strain: C57BL/6J. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/53502/53502fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53502/53502fig3.jpg" /> 
Figure 3: OCT Cross-Sectional CNV Visualization. (A) OCT en face view of CNV lesion. (B) OCT cross-sectional B-scan of retina with CNV circled in yellow. Mouse Strain: C57BL/6J. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/53502/53502fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Mouse Model for Laser-induced Choroidal Neovascularization at JoVE.com

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) ...

a-mouse-model-for-laser-induced-choroidal-neovascularization

Universidad San Sebastian

Research

JoVE Journal

Medicine

17.5K Views.  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.

Video: A Mouse Model for Laser-induced Choroidal Neovascularization

Laser-Induced Chronic Ocular Hypertension Model on SD Rats

Glaucoma is one of the major causes of blindness in the world. Elevated intraocular pressure is a major risk factor. Laser photocoagulation induced ocular hypertension is one of the well established animal models. This video demonstrates how to induce ocular hypertension by Argon laser photocoagulation in rat.

Glaucoma is one of the major causes of blindness in the world. Elevated intraocular pressure is a major risk factor. Laser photocoagulation induced ocular hypertension is one of the well established animal models. This video demonstrates how to induce ocular hypertension by Argon laser photocoagulation in rat.

Glaucoma is one of the major causes of blindness in the world. Elevated intraocular pressure is a major risk factor. Laser photocoagulation induced ocular ...

Biology

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 ...

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 ...

Implantation and Evaluation of Melanoma in the Murine Choroid via Optical Coherence Tomography

Establishing experimental choroidal melanoma models is challenging in terms of the ability to induce tumors at the correct localization. In addition, difficulties in observing posterior choroidal melanoma in vivo limit tumor location and growth evaluation in real-time. The approach described here optimizes techniques for establishing choroidal melanoma in mice via a multi-step sub-choroidal B16LS9 cell injection procedure. To enable precision in injecting into the small dimensions of the mouse uvea, the complete procedure is performed under a microscope. First, a conjunctival peritomy is formed in the dorsal-temporal area of the eye. Then, a tract into the sub-choroidal space is created by inserting a needle through the exposed sclera. This is followed by the insertion of a blunt needle into the tract and the injection of melanoma cells into the choroid. Immediately after injection, noninvasive optical coherence tomography (OCT) imaging is utilized to determine tumor location and progress. Retinal detachment is evaluated as a predictor of tumor site and size. The presented method enables the reproducible induction of choroid-localized melanoma in mice and the live imaging of tumor growth evaluation. As such, it provides a valuable tool for studying intraocular tumors.

Implantation and Evaluation of Melanoma in the Murine Choroid via Optical Coherence Tomography

The present protocol describes the implantation and evaluation of melanoma in the murine choroid utilizing optical coherence tomography.

Establishing experimental choroidal melanoma models is challenging in terms of the ability to induce tumors at the correct localization. In addition, ...

Cancer Research

In Vivo Multimodal Imaging and Analysis of Mouse Laser-Induced Choroidal Neovascularization Model

Laser-induced choroidal neovascularization (CNV) is a well-established model to mimic the wet form of age-related macular degeneration (AMD). In this protocol, we aim to guide the reader not simply through the technical considerations of generating laser-induced lesions to trigger neovascular processes, but rather focus on the powerful information that can be obtained from multimodal longitudinal in vivo imaging throughout the follow-up period.
The laser-induced mouse CNV model was generated by a diode laser administration. Multimodal in vivo imaging techniques were used to monitor CNV induction, progression and regression. First, spectral domain optical coherence tomography (SD-OCT) was performed immediately after the lasering to verify a break of Bruch&#39;s membrane. Subsequent in vivo imaging using fluorescein angiography (FA) confirmed successful damage of Bruch&#39;s membrane from serial images acquired at the choroidal level. Longitudinal follow-up of CNV proliferation and regression on days 5, 10, and 14 after the lasering was performed using both SD-OCT and FA. Simple and reliable grading of leaky CNV leasions from FA images is&#160;presented. Automated segmentation for measurement of total retinal thickness, combined with manual caliber application for measurement of retinal thickness at CNV sites, allow unbiased evaluation of the presence of edema. Finally, histological verification of CNV is performed using isolectin GS-IB4 staining on choroidal flatmounts. The staining is thresholded, and the isolectin-positive area is calculated with ImageJ.
This protocol is especially useful in therapeutics studies requiring high-throughput-like screening of CNV pathology as it allows fast, multimodal, and reliable classification of CNV pathology and retinal edema. In addition, high resolution SD-OCT enables the recording of other pathological hallmarks, such as the accumulation of subretinal or intraretinal fluid. However, this method does not provide a possibility to automate CNV volume analysis from SD-OCT images, which has to be performed manually.

In Vivo Multimodal Imaging and Analysis of Mouse Laser-Induced Choroidal Neovascularization Model

Here, we present the usefulness of longitudinal in vivo imaging in the follow-up of morphological changes of laser-induced choroidal neovascularization in mice.

Laser-induced choroidal neovascularization (CNV) is a well-established model to mimic the wet form of age-related macular degeneration (AMD). In this ...

Neuroscience

Optimization of the Retinal Vein Occlusion Mouse Model to Limit Variability

Mouse models of retinal vein occlusion (RVO) are often used in ophthalmology to study hypoxic-ischemic injury in the neural retina. In this report, a detailed method pointing out critical steps is provided with recommendations for optimization to achieve consistently successful occlusion rates across different genetically modified mouse strains. The RVO mouse model consists primarily of the intravenous administration of a photosensitizer dye followed by laser photocoagulation using a retinal imaging microscope attached to an ophthalmic guided laser. Three variables were identified as determinants of occlusion consistency. By adjusting the wait time after rose bengal administration and balancing the baseline and experimental laser output, the variability across experiments can be limited and a higher success rate of occlusions achieved. This method can be used to study retinal diseases that are characterized by retinal edema and hypoxic-ischemic injury. Additionally, as this model induces vascular injury, it can also be applied to study the neurovasculature, neuronal death, and inflammation.

Here, we describe an optimized protocol for retinal vein occlusion using rose bengal and a laser-guided retinal imaging microscope system with recommendations to maximize its reproducibility in genetically modified strains.

Mouse models of retinal vein occlusion (RVO) are often used in ophthalmology to study hypoxic-ischemic injury in the neural retina. In this report, a ...

Application of Optical Coherence Tomography to a Mouse Model of Retinopathy

Optical coherence tomography (OCT) offers a noninvasive method for the diagnosis of retinopathy. The OCT machine can capture retinal crosssectional images from which the retinal thickness can be calculated. Although OCT is widely used in clinical practice, its application in basic research is not as prevalent, especially in small animals such as mice. Because of the small size of their eyeballs, it is challenging to conduct fundus imaging examinations in mice. Therefore, a specialized retinal imaging system is required to accommodate OCT imaging on small animals. This article demonstrates a small-animal-specific system for OCT examination procedures and a detailed method for image analysis. The results of retinal OCT examination of very-low-density lipoprotein receptor (Vldlr) knockout mice and C57BL/6J mice are presented. The OCT images of C57BL/6J mice showed retinal layers, while those of Vldlr knockout mice showed subretinal neovascularization and retinal thinning. In summary, OCT examination could facilitate the noninvasive detection and measurement of retinopathy in mouse models.

Here, we describe an in vivo imaging technique using optical coherence tomography to facilitate the diagnosis and quantitative measurement of retinopathy in mice.

Optical coherence tomography (OCT) offers a noninvasive method for the diagnosis of retinopathy. The OCT machine can capture retinal crosssectional images ...

Image-Guided Optical Coherence Tomography to Assess Structural Changes in Rodent Retinas

Ocular diseases, such as age-related macular degeneration, glaucoma, retinitis pigmentosa, and uveitis, are always accompanied by retinal structural changes. These diseases affecting the fundus always exhibit typical abnormalities in certain cell types in the retina, including photoreceptor cells, retinal ganglion cells, cells in the retinal blood vessels, and cells in the choroidal vascular cells. Noninvasive, highly efficient, and adaptable imaging techniques are required for both clinical practice and basic research. Image-guided optical coherence tomography (OCT) satisfies these requirements because it combines fundus photography and high-resolution OCT, providing an accurate diagnosis of tiny lesions as well as important changes in the retinal architecture. This study details the procedures of data collection and data analysis for image-guided OCT and demonstrates its application in rodent models of choroidal neovascularization (CNV), optic nerve crush (ONC), light-induced retinal degeneration, and experimental autoimmune uveitis (EAU). This technique helps researchers in the eye field to identify rodent retinal structural changes conveniently, reliably, and tractably.

The protocol presented here details the procedures of data collection and data analysis for image-guided optical coherence tomography (OCT) and demonstrates its application in multiple rodent models of ocular diseases.

Ocular diseases, such as age-related macular degeneration, glaucoma, retinitis pigmentosa, and uveitis, are always accompanied by retinal structural ...

Characterizing Different Lesion Types of Laser-Induced CNV in Mouse

Characterization of Vascular Morphology of Neovascular Age-Related Macular Degeneration by Indocyanine Green Angiography

Age-related macular degeneration (AMD) is a leading cause of blindness among older individuals, and its prevalence is rapidly increasing due to the aging population. Choroidal neovascularization (CNV) or wet AMD, which accounts for 10%-20% of all AMD cases, is responsible for an alarming 80%-90% of AMD-related blindness. Current anti-VEGF therapies show suboptimal responses in approximately 50% of patients. Resistance to anti-VEGF treatment in CNV patients is often associated with arteriolar CNV, while responders tend to have capillary CNV. While fluorescein angiography (FA) is commonly used to assess leakage patterns in wet AMD patients and animal models, it does not provide information about CNV vascular morphology (arteriolar CNV vs. capillary CNV). This protocol introduces the use of indocyanine green angiography (ICGA) to characterize lesion types in laser-induced CNV mouse models. This method is crucial for investigating the mechanisms and treatment strategies for anti-VEGF resistance in wet AMD. It is recommended to incorporate ICGA alongside FA for comprehensive assessment of both leakage and vascular features of CNV in mechanistic and therapeutic studies.

Author Spotlight: Overcoming Anti-VEGF Resistance Through Advanced Vascular Morphology Assessment in Choroidal Neovascularization

Currently, fluorescein angiography (FA) is the preferred method for identifying leakage patterns in animal models of choroidal neovascularization (CNV). However, FA does not provide information about vascular morphology. This protocol outlines the use of indocyanine green angiography (ICGA) to characterize different lesion types of laser-induced CNV in mouse models.

Age-related macular degeneration (AMD) is a leading cause of blindness among older individuals, and its prevalence is rapidly increasing due to the aging ...

Monitoring Dynamic Growth of Retinal Vessels in Oxygen-Induced Retinopathy Mouse Model

Oxygen-induced retinopathy (OIR) is widely used to study abnormal vessel growth in ischemic retinal diseases, including retinopathy of prematurity (ROP), proliferative diabetic retinopathy (PDR), and retinal vein occlusion (RVO). Most OIR studies observe retinal neovascularization at specific time points; however, the dynamic vessel growth in live mice along a time course, which is essential for understanding the OIR-related vessel diseases, has been understudied. Here, we describe a step-by-step protocol for the induction of the OIR mouse model, highlighting the potential pitfalls, and providing an improved method to quickly quantify areas of vaso-obliteration (VO) and neovascularization (NV) using immunofluorescence staining. More importantly, we monitored vessel regrowth in live mice from P15 to P25 by performing fluorescein fundus angiography (FFA) in the OIR mouse model. The application of FFA to the OIR mouse model allows us to observe the remodeling process during vessel regrowth.

This protocol describes a detailed method for the preparation and immunofluorescence staining of mice retinal flat mounts and analysis. The use of fluorescein fundus angiography (FFA) for mice pups and image processing are described in detail as well.

Oxygen-induced retinopathy (OIR) is widely used to study abnormal vessel growth in ischemic retinal diseases, including retinopathy of prematurity (ROP), ...

A Mouse Model for Laser-induced Choroidal Neovascularization

Summary

Explore More Videos

Chapters in this video

Laser-Induced Chronic Ocular Hypertension Model on SD Rats

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

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

Implantation and Evaluation of Melanoma in the Murine Choroid via Optical Coherence Tomography

In Vivo Multimodal Imaging and Analysis of Mouse Laser-Induced Choroidal Neovascularization Model

Optimization of the Retinal Vein Occlusion Mouse Model to Limit Variability

Application of Optical Coherence Tomography to a Mouse Model of Retinopathy

Image-Guided Optical Coherence Tomography to Assess Structural Changes in Rodent Retinas

Characterization of Vascular Morphology of Neovascular Age-Related Macular Degeneration by Indocyanine Green Angiography

Monitoring Dynamic Growth of Retinal Vessels in Oxygen-Induced Retinopathy Mouse Model