This work was supported by the National Eye Institute (EY015279 and EY028597) and Delaware INBRE (P20 GM103446).

The following protocol was approved by the University of Delaware Institutional Animal Care and Use Committee under protocol #1039-2021-1. In compliance with the Association for Research in Vision and Ophthalmology (ARVO) for the Use of Animals in Ophthalmic and Vision Research24, all ocular survival surgeries can only be performed in one eye. See Table of Materials for details about reagents and instruments used in this protocol. Alternative approaches for extracting the lens fibers&#160;and suturing&#160;are shown in Figure 2&#160; and&#160;Figure 3.
1. Animals
<ol>
	<li>Use any wild-type or genetically modified laboratory mouse strain that has eyes with intact lenses for this procedure.
	<ol>
		<li>Use C57BL/6 mice as a starting point, as their responses are highly reproducible at both the morphological and molecular levels25,26. 
		NOTE: C57BL/6 or any mouse strain can be used. In the video protocol, the FVB mouse is used for better visual demonstration.</li>
		<li>Perform studies of the response of mutants on other mouse strains with strain-matched controls. 
		NOTE: With practice, this procedure may be performed on mice whose lenses have cataracts as long as the lens capsule is intact and the lens has not undergone significant liquefaction. See the study of mice with lens conditional deletion of the transcription factor Sip127.</li>
	</ol>
	</li>
	<li>Perform this surgery on mice of either sex as young as 8 weeks to as old as 24 months25,26.
	<ol>
		<li>Perform this surgery in even younger mice (after eyelid opening) with practice, although it will be more difficult as these eyes will be significantly smaller.</li>
		<li>Take particular care for surgeries on elderly mice due to their lower tolerance for anesthesia25,26. 
		&#8203;NOTE: Sex differences in surgery responses are more prevalent in elderly mice26.</li>
	</ol>
	</li>
</ol>
2. Preparation of solutions and surgical tools
<ol>
	<li>Obtain anesthetizing and analgesic agents, dilation drops, sterile disposable scalpel, #5 Dumont forceps, 2-110 suture forceps, needles (26 G 1/2 and 27 G 45&#176; bent dispensing tip 1&#34;), syringes, and balanced saline solution (BSS).</li>
	<li>Sterilize surgical instruments in an autoclave.</li>
</ol>
3. Prior to anesthesia
<ol>
	<li>Prior to anesthesia, give the mouse one drop each of 1% atropine and 2.5% phenylephrine to dilate the pupil (see Figure 1A) of the eye undergoing surgery. Apply ophthalmic ointment to the non-operated eye to keep the surface from drying. 
	NOTE: The ARVO statement on the use of animals in eye research24 only allows a single eye to undergo survival surgery, leaving the other eye untouched so that the animal retains vision. Thus, survival surgeries are always unilateral; however, the contralateral, unoperated eye can be operated on just prior to sacrifice to create animal-matched &#39;time zero&#39; controls.</li>
	<li>Make sure the eye drops do not run down the mouse&#39;s face and enter the nose, as this can obstruct its breathing.</li>
	<li>Prepare a syringe with sterile BSS using a sterile 26 G 1/2 needle. Fit the syringe with a sterile 27 G 45&#176; bent blunt dispensing tip of 1 inch (referred to as a dispensing tip below).</li>
</ol>
4. Anesthesia
<ol>
	<li>Following eye drops, anesthetize the mouse with an intraperitoneal injection of xylazine/ketamine/acepromazine (35 mg/kg, 80 mg/kg, and 0.5 mg/kg) solution (Figure 1B). 
	NOTE: Inhaled anesthesia might not be possible unless a specialized nose cone can be utilized to allow for access to the eye needed for surgery.</li>
</ol>
5. Making the incision
<ol>
	<li>Once the mouse is completely under anesthesia, as measured by the lack of toe pinch reflex, and its pupil has dilated completely (2 mm) (Figure 4A), the mouse is ready for surgery.
	<ol>
		<li>Move the anesthetized mouse to the stage of a compound dissecting light microscope. Use a heater pad to ensure the mouse has adequate thermal regulation and use sterile drapes.</li>
		<li>View the eye under 10x-20x magnification using a dissecting microscope (see Table of Materials for specifics).</li>
	</ol>
	</li>
	<li>Make a 1-1.5 mm incision in the center of the cornea using a sterile surgical scalpel. Ensure the incision extends through the cornea and incises the anterior lens capsule (Figure 1C and Figure 4B).</li>
</ol>
6. Separating the lens fibers from the capsular bag
<ol>
	<li>Grasp the syringe with a dispensing tip containing sterile BSS and gently expel 1 drop of BSS above the cornea to wet the surface for further manipulation.</li>
	<li>Separate the lens fiber mass from the lens capsule.
	<ol>
		<li>Insert the dispensing tip through the central corneal incision into the lens capsule incision, then gently move the needle in a circular motion while slowly releasing BSS to separate apical-apical connections between lens epithelial and fiber cells (hydro dissection), ultimately expelling the lens fiber mass (Figure 1D).</li>
		<li>In most cases, the fiber mass will eventually adhere to the dispensing tip during this procedure (Figure 2A). If this happens, gently remove the dispensing tip from the inside eye; and clean with a sterile towel to remove this tissue. Repeat until all lens fiber cells are removed.</li>
		<li>Alternatively, use micro forceps (number 5 Dumont) to extract the lens nucleus (Figure 2B). If using this method, be careful that the original corneal incision exceeds the diameter of the nucleus to ensure easy removal.</li>
	</ol>
	</li>
	<li>Rinse the lens capsule with sterile BSS to remove any remaining fibers.</li>
	<li>Confirm that the lens capsule is left inside the eye (Figure 1E and Figure 4C) by viewing its translucent/glassy appearance through the microscope.</li>
</ol>
7. Inflating the anterior chamber and suturing the corneal incision
<ol>
	<li>Inflate the anterior chamber to its normal depth by injecting BSS through the incision into the anterior chamber.
	<ol>
		<li>Add inhibitors or activators of signaling pathways of interest to the BSS to test their roles in the lens injury response12,15. 
		NOTE: Animals can also be treated systemically with pathway inhibitors, which may be needed depending on the pharmacokinetics of drug turnover as the aqueous humor balance is re-established after surgery15.</li>
	</ol>
	</li>
	<li>Suture the corneal incision with square knots using 10-0 nylon thread&#160;with a tapered, curved 5.3 mm 1/2c needle (Figure 1F and Figure 4D-F).
	<ol>
		<li>Place either one square knot stitch in the central part of the incision or two square knot stitches at both the upper and lower halves of the original incision (see Figure 3). Number of stitches depends on the length of the corneal incision.</li>
		<li>Do not puncture both corneal flaps at one time. Instead, hold one corneal flap using forceps in the non-dominant hand and push the needle/nylon through. Once one side of the cornea has been punctured, push the needle/nylon through the next corneal flap. Close the corneal flaps using square knot stitches.</li>
	</ol>
	</li>
</ol>
8. Post-surgical care
NOTE: Topical antibiotic ointment is applied to the eye, and analgesic agents are administered. Mice do not&#160;usually show signs of distress after initial surgery. If needed, slow-release version of buprenorphine could be used.
<ol>
	<li>Immediately following this surgery, do the following sub-steps.
	<ol>
		<li>Apply one drop of 0.5% erythromycin ophthalmic ointment directly above the suture. Make sure the entire incision is covered in ointment.</li>
		<li>Administer an intraperitoneal injection of buprenorphine (0.1 mg/kg).</li>
	</ol>
	</li>
	<li>Allow the mouse to recover from anesthesia in a cage placed on a heating pad to optimize thermal support.&#160; 
	NOTE: Monitor the animal for signs of trauma, distress, or discomfort. Some signs include infection/scratching at the surgical site, displaced suture, loss of fur, etc.</li>
</ol>
9. Monitoring animal recovery
<ol>
	<li>Monitor the animal&#39;s recovery after surgery until it awakes from anesthesia.</li>
	<li>Check mice every day for the first 3 days following surgery. 
	NOTE: It is usually not needed to remove stitches because mice are either sacrificed prior to full healing (2 weeks PCS) or stitches spontaneously fall out.</li>
	<li>Administer buprenorphine orally, as needed for pain management after 4-8 h post-surgery, at a dosage of 0.5 mg/kg (in Jello) or a dosage of 0.1 mg/kg for those not eating. Alternatively, administer buprenorphine extended-release just prior to surgery. 
	NOTE: When possible, preemptive analgesics should be utilized, yet this may require adjustments to the analgesic protocol that require review and approval from the institutional animal care and use committee.</li>
</ol>
10. Euthanasia
<ol>
	<li>Euthanize the mouse at a desired time post-surgery
	<ol>
		<li>Removed the operated eye using the unoperated eye as control tissue; or re-anesthetize the animal, perform lens fiber cell removal surgery on the previously unoperated contralateral eye, then immediately sacrifice the animal (without allowing it to awake from anesthesia) to provide a time zero operated control.</li>
		<li>Ensure quick dissection and tissue harvest. 
		NOTE: LECs undergo transcriptomic changes within minutes of surgery so rapid tissue harvest and fixation/freezing are necessary when studying early LEC responses to surgery.</li>
	</ol>
	</li>
</ol>
11. Performing further analysis on harvested lens tissues
<ol>
	<li>Use the obtained tissue for immunostaining, RNA sequencing, cell culture, or&#160;western analysis12,15,23,25,26,28. Flow cytometry is theoretically possible as well12, although the small number of LECs in the mouse lens (~40,000)29 and the presence of capsule associated cell debris following surgery make this challenging.</li>
	<li>Immunostaining
	<ol>
		<li>Euthanize the mouse in a chamber via CO2 inhalation (4 L/min) from a gas cylinder. Increase the flow rate to 8-10 L/min once the mouse is unconscious and continue this exposure until complete cessation of breathing plus 1 min. 
		NOTE: The mouse can be euthanized using an appropriate method per AVMA Guidelines for the Euthanasia of Animals or per Institutional Animal Care and Use Committee approved guidelines or policies.</li>
		<li>Ensure proper sacrifice by employing a secondary method of euthanasia, e.g., cervical dislocation. 
		NOTE: The Duncan lab has investigated time points ranging from minutes to months PCS12,15,23.</li>
		<li>After the mouse has been humanely sacrificed, isolate the operated eye for analysis.
		<ol>
			<li>Hold micro forceps in the non-dominant hand to grasp the operated eye while holding micro scissors in the dominant hand.</li>
			<li>Gently clip the optic nerve and muscles anchoring the eye. 
			NOTE: Be particularly careful handling eyes that were recently operated on versus those that have had longer to heal PCS. Recently operated eyes are more structurally fragile.</li>
		</ol>
		</li>
		<li>Embed the dissected eye in OCT media for later cryo-sectioning, immunostaining, and semi-quantitative confocal microscopy28,30.</li>
	</ol>
	</li>
	<li>Isolation of the capsular bag with associated cells for RNA sequencing, cell culture, or western blot analysis
	<ol>
		<li>Euthanize the mouse in a chamber via CO2 inhalation (4 L/min) from a gas cylinder. Increase the flow rate to 8-10 L/min once the mouse is unconscious and continue this exposure until complete cessation of breathing plus 1 min.</li>
		<li>Ensure proper sacrifice by employing a secondary method of euthanasia, e.g., cervical dislocation.</li>
		<li>If an early timepoint PCS (within 1-2 days) is required, either remove stitches prior to tissue extraction or make another incision in the cornea through which to extract the lens capsule. 
		NOTE: At later time points PCS, the stitches will likely have fallen out, so a new incision will be needed.</li>
	</ol>
	</li>
	<li>Locate the translucent lens capsule.
	<ol>
		<li>Gently insert the micro forceps through the corneal opening, grasp the capsule, and remove it from the anterior chamber.</li>
		<li>Place the capsule in a 1.5 mL microcentrifuge tube or culture dish for further analysis.</li>
	</ol>
	</li>
</ol>

Replicating Cataract Surgical Techniques in a Mouse Model Through LEC Extraction

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The Duncan lab has received financial support from Pliantx to test anti-PCO therapies using this surgery model.

This surgical technique requires advanced mouse handling and micro-surgical skills that take practice to develop. The most difficult to master is the placement of fine sutures in the cornea to close the wound. Suppose the experimenter is a total novice to suturing. In that case, it is recommended to first practice using string and cloth, then move to practice using large-diameter sutures on small fruits to master the needed hand movements to make surgical square knots. Then, the experimenter must practice making incision...

The lens is a highly organized, transparent tissue that refracts light to produce a clearly focused image onto the retina1,2,3. This specialized organ is surrounded by an uninterrupted basement membrane (the capsule), which isolates the lens from other parts of the eye. The inner anterior surface of the capsule anchors a monolayer of lens epithelial cells (LECs), which then differentiate at the lens equator into lens fiber cells, which comprise the vast majority of the lens3. A cataract occurs when the lens loses its transparency due to factors like aging, genetic mutation, UV radiation, oxidative stress, and ocular trauma4. Cataract is a leading cause of blindness worldwide, particularly in countries with poor medical care5. However, this condition is now readily treated by surgical removal of opaque lens cells by phacoemulsification in which the central lens capsule and attached lens epithelium are removed, followed by the use of a vibrating probe to break the cellular mass of the lens into smaller fragments that can be suctioned out; leaving behind a capsular bag with some attached equatorial LECs1. Vision is then most commonly restored by post-surgical implantation of an artificial intraocular lens (IOL).
While cataract surgery (CS) is a highly effective and minimally invasive treatment for cataracts, a patient&#39;s post-surgical recovery can be acutely hampered by the development of ocular inflammation5,6,8. This inflammation can cause post-surgical pain, retinal edema leading to retinal detachment, as well as exacerbation of other inflammatory and fibrotic conditions like uveitis and glaucoma4,7,8,9. Ocular inflammation post-CS (PCS) is generally treated with anti-inflammatory eye drops which are plagued by poor patient compliance or dropless cataract surgery that can lead to an increased prevalence of floaters8,10,11. In the long term, the outcomes of CS can be compromised by the development of posterior capsular opacification (PCO)12. PCO occurs when residual LECs left behind after surgery undergo a wound healing response, proliferate, and migrate onto the posterior lens capsule at&#160;months to years PCS contributing to secondary visual obstruction13,14.
Understanding the molecular mechanisms by which CS results in such acute and chronic responses represents a major challenge in the field, as much of the literature investigating PCO relies on the induction of LEC conversion to myofibroblasts in culture via treatment with active transforming growth factor beta (TGF&#946;)12,15. Human capsular bag models created from cadaver lenses cultured in vitro better reflect the biology of the lens PCS as LECs are cultured on their native basement membrane but are difficult to manipulate mechanistically, do not recapitulate the intraocular environment, and are inherently variable due to lens to lens (or donor to donor) variability16,17. Rabbit1,18 and non-human primate19,20&#160;in vivo models of cataract surgery overcome some of these issues but still are not easy to manipulate for mechanistic studies. Notably, a prior study of mammalian lens regeneration found significant lens regeneration within four weeks after lens fiber cells were removed from mice, leaving the lens epithelial cells and capsule behind, while no lens regrowth occurred when the entire lens was removed21,22. We subsequently streamlined this procedure and optimized it for the study of the acute response of LECs to lens fiber cell removal and their subsequent conversion to a mixed population of cells with myofibroblast and fiber cell properties.
Using this mouse model of cataract surgery, we have shown that lens epithelial cells drastically remodel their transcriptome by 6 hours PCS to produce numerous proinflammatory cytokines23, while they begin expressing fibrotic markers as early as 24 h PCS12,23, prior to the onset of canonical TGF&#946; signaling. Mechanistic experiments in knockout mice using this model revealed that cellular fibronectin expression by LECs is required for a sustained fibrotic response PCS, likely due to both its role in the assembly of the fibrotic ECM and cell signaling12. Other studies showed that &#945;V&#946;8-integrin is required for the transition of LECs to myofibroblasts due to its function in TGF&#946; activation, and the potential of this approach to identify anti-PCO therapeutic leads was confirmed as a &#945;V&#946;8-integrin antagonist also blocked LEC EMT15.
Here, we provide a detailed protocol describing how to remove the lens fiber cells from living mice (Figure 1and&#160;Figure 4) while leaving behind the lens capsule and LECs to model the LEC response to cataract surgery.&#160;

<table><tbody><tr><td>0.5% Erythryomycin opthalmic ointment USP</td><td>Baush Lomb</td><td>24208-910-55</td><td></td></tr><tr><td>1% Atropine sulfate ophthalmic solution</td><td>Amneal</td><td>60219-1748-2</td><td></td></tr><tr><td>1% Tropicamide opthalmic solution USP</td><td>Akorn</td><td>17478-102-12</td><td></td></tr><tr><td>10-0 Nylon suture</td><td>Ethicon</td><td>7707G</td><td></td></tr><tr><td>2.5% Phenylephrine hydrochloride ophtahlmic solution USP</td><td>Akorn</td><td>17478-200-12</td><td></td></tr><tr><td>26 G 1/2 Needles - straight</td><td>BD PrecisionGlide</td><td>5111</td><td></td></tr><tr><td>27 G 45&amp;deg; bent dispensing tip 1&quot;</td><td>Harfington</td><td></td><td></td></tr><tr><td>Balanced saline solution</td><td>Phoenix</td><td>57319-555-06</td><td></td></tr><tr><td>Bupranorphine (0.1 mg/kg)</td><td>APP Pharmaceticals</td><td>401730D</td><td></td></tr><tr><td>Chlorhexidine solution</td><td></td><td></td><td></td></tr><tr><td>Needle holder 2-110</td><td>Duckworth &amp;amp; Kent</td><td>2-110-3E</td><td></td></tr><tr><td>Noyes scissors, straight</td><td>Fine Science Tools</td><td>12060-02</td><td></td></tr><tr><td>SMZ800 Nikon model microscope</td><td>Nikon</td><td></td><td></td></tr><tr><td>Sterile disposable scalpel No.11</td><td>Feather</td><td>2975#11</td><td></td></tr><tr><td>Tweezers #5 Dumont</td><td>Electron Microscopy Sciences</td><td>72700-D</td><td></td></tr><tr><td>Xylazine/Ketamine/Acepromazine (35 mg/kg, 80 mg/kg, 0.5 mg/kg) solution</td><td>APP Pharmaceticals</td><td>401730D</td><td></td></tr></tbody></table>

modeling cataract surgery in mice

Cataract surgery (CS) is an effective treatment for cataracts, a major cause of visual disability worldwide. However, CS leads to ocular inflammation, and in the long term, it can result in posterior capsular opacification (PCO) and/or lens dislocation driven by the post-surgical overgrowth of lens epithelial cells (LECs) and their conversion to myofibroblasts and/or aberrant fiber cells. However, the molecular mechanisms by which CS results in inflammation and PCO are still obscure because most in vitro models do not recapitulate the wound healing response of LECs seen in vivo, while traditional animal models of cataract surgery, such as rabbits, do not allow the genetic manipulation of gene expression to test mechanisms. Recently, our laboratory and others have successfully used genetically modified mice to study the molecular mechanisms that drive the induction of proinflammatory signaling and LEC epithelial to mesenchymal transition, leading to new insight into PCO pathogenesis. Here, we report the established protocol for modeling cataract surgery in mice, which allows for robust transcriptional profiling of the response of LECs to lens fiber cell removal via RNAseq, the evaluation of protein expression by semi-quantitative immunofluorescence, and the use of modern mouse genetics tools to test the function of genes that are hypothesized to participate in the pathogenesis of acute sequelae like inflammation as well as the later conversion of LECs to myofibroblasts and/or aberrant lens fiber cells.

Based on the results from this surgical method, the Duncan Lab has used this mouse model of cataract surgery to construct an injury response time course of lens epithelial cells post-surgery (Figure 5). By 6 h after surgery, 5% of the lens epithelial transcriptome is differentially expressed (Figure 6A), including the upregulation of numerous immediate early response genes and proinflammatory cytokines23 (Figure 6B</s...

Watch this Scientific Journal Video about Unraveling Molecular Mechanisms in PCO and Fibrosis Following Cataract Surgery at JoVE.com

Author Spotlight: Unraveling the Molecular Mechanisms in PCO and Fibrosis Following Cataract Surgery

This method models cataract surgery in vivo by removing lens fiber cells from adult mice and leaving behind the capsular bag with attached lens epithelial cells (LECs). The injury response is then assessed at various times post-surgery using molecular and morphological criteria.

Modeling Cataract Surgery in Mice

Cataract surgery (CS) is an effective treatment for cataracts, a major cause of visual disability worldwide. However, CS leads to ocular inflammation, and ...

Research

JoVE Journal

Biology

914 Views.  University of Delaware. This method models cataract surgery in vivo by removing lens fiber cells from adult mice and leaving behind the capsular bag with attached lens epithelial cells (LECs). The injury response is then assessed at various times post-surgery using molecular and morphological criteria. 

Video: Author Spotlight: Unraveling the Molecular Mechanisms in PCO and Fibrosis Following Cataract Surgery

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 ...

Medicine

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.

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) ...

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

Ultrahigh Resolution Mouse Optical Coherence Tomography to Aid Intraocular Injection in Retinal Gene Therapy Research

HR-SD-OCT is utilized to monitor the progression of photoreceptor degeneration in live mouse models, assess the delivery of therapeutic agents into the subretinal space, and to evaluate toxicity and efficacy in vivo. HR-SD-OCT uses near infrared light (800-880 nm) and has optics specifically designed for the unique optics of the mouse eye with sub-2-micron axial resolution. Transgenic mouse models of outer retinal (photoreceptor) degeneration and controls were imaged to assess the disease progression. Pulled glass microneedles were used to deliver sub retinal injections of adeno-associated virus (AAV) or nanoparticles (NP) via a trans-scleral and trans-choroidal approach. Careful positioning of the needle into the subretinal space was required prior to a calibrated pressure injection, which delivers fluid into the sub retinal space. Real time subretinal surgery was conducted on our retinal imaging system (RIS). HR-SD-OCT demonstrated progressive uniform retinal degeneration due to expression of a toxic mutant human mutant rhodopsin (P347S) (RHOP347S) transgene in mice. HR-SD-OCT allows rigorous quantification of all the retinal layers. Outer nuclear layer (ONL) thickness and photoreceptor outer segment length (OSL) measurements correlate with photoreceptor vitality, degeneration, or rescue. The RIS delivery system allows real-time visualization of subretinal injections in neonatal (~P10-14) or adult mice, and HR-SD-OCT immediately determines success of delivery and maps areal extent. HR-SD-OCT is a powerful tool that can evaluate the success of subretinal surgery in mice, in addition to measuring vitality of photoreceptors in vivo. HR-SD-OCT can also be used to identify uniform animal cohorts to evaluate the extent of retinal degeneration, toxicity, and therapeutic rescue in preclinical gene therapy research studies.

Here we demonstrate a novel approach to using high resolution spectral-domain optical coherence tomography (HR-SD-OCT) to assist delivery of gene therapy agents into the subretinal space, assess its areal coverage, and characterize photoreceptor vitality.

HR-SD-OCT is utilized to monitor the progression of photoreceptor degeneration in live mouse models, assess the delivery of therapeutic agents into the ...

In Vivo Imaging of Cx3cr1gfp/gfp Reporter Mice with Spectral-domain Optical Coherence Tomography and Scanning Laser Ophthalmoscopy

Spectral domain optical coherence tomography (SD-OCT) and scanning laser ophthalmoscopy (SLO) are extensively used in experimental ophthalmology. In the present protocol, mice expressing green fluorescent protein (gfp) under the promoter of Cx3cr1 (BALB/c-Cx3cr1gfp/gfp) were used to image microglia cells in vivo in the retina. Microglia are resident macrophages of the retina and have been implicated in several retinal diseases1,2,3,4,5,6. This protocol provides a detailed approach for generation of retinal B-scans, with SD-OCT, and imaging of microglia cell distribution in Cx3cr1gfp/gfp mice with SLO in vivo, using an ophthalmic imaging platform system. The protocol can be used in several reporter mouse lines. However, there are some limitations to the protocol presented here. First, both SLO and SD-OCT, when used in the high-resolution mode, collect data with high axial resolution but the lateral resolution is lower (3.5 &#181;m and 6 &#181;m, respectively). Moreover, the focus and saturation level in SLO is highly dependent on parameter selection and correct alignment of the eye. Additionally, using devices designed for human patients in mice is challenging due to the higher total optical power of the mouse eye compared to the human eye; this can lead to lateral magnification inaccuracies7, which are also dependent on the magnification by the mouse lens among others. However, despite that the axial scan position is dependent upon lateral magnification, the axial SD-OCT measurements are accurate8.

In Vivo Imaging of Cx3cr1gfp/gfp Reporter Mice with Spectral-domain Optical Coherence Tomography and Scanning Laser Ophthalmoscopy

This protocol describes how high-resolution imaging techniques such as spectral domain optical coherence tomography and scanning laser ophthalmoscopy can be utilized in small rodents, using an ophthalmic imaging platform system, to obtain information on retinal thickness and microglial cell distribution, respectively.

Spectral domain optical coherence tomography (SD-OCT) and scanning laser ophthalmoscopy (SLO) are extensively used in experimental ophthalmology. In the ...

A Model of Glaucoma Induced by Circumlimbal Suture in Rats and Mice

The circumlimbal suture is a technique for inducing experimental glaucoma in rodents by chronically elevating intraocular pressure (IOP), a well-known risk factor for glaucoma. This protocol demonstrates a step-by-step guide on this technique in Long Evans rats and C57BL/6 mice. Under general anesthesia, a "purse-string" suture is applied on the conjunctiva, around the equator and behind the limbus of the eye. The fellow eye serves as an untreated control. Over the duration of our study, which was a period of 8 weeks for rats and 12 weeks for mice, IOP remained elevated, as measured regularly by rebound tonometry in conscious animals without topical anesthesia. In both species, the sutured eyes showed electroretinogram features consistent with preferential inner retinal dysfunction. Optical coherence tomography showed selective thinning of the retinal nerve fiber layer. Histology of the rat retina in cross-section found reduced cell density in the ganglion cell layer, but no change in other cellular layers. Staining of flat-mounted mouse retinae with a ganglion cell specific marker (RBPMS) confirmed ganglion cell loss. The circumlimbal suture is a simple, minimally invasive and cost-effective way to induce ocular hypertension that leads to ganglion cell injury in both rats and mice.

Chronic ocular hypertension is induced by applying a circumlimbal suture in rats and mice, leading to functional and structural deterioration of the retinal ganglion cells consistent with glaucoma.

The circumlimbal suture is a technique for inducing experimental glaucoma in rodents by chronically elevating intraocular pressure (IOP), a well-known ...

Modeling Stroke in Mice: Focal Cortical Lesions by Photothrombosis

Stroke is a leading cause of death and acquired adult disability in developed countries. Despite extensive investigation for novel therapeutic strategies, there remain limited therapeutic options for stroke patients. Therefore, more research is needed for pathophysiological pathways such as post-stroke inflammation, angiogenesis, neuronal plasticity, and regeneration. Given the inability of in vitro models to reproduce the complexity of the brain, experimental stroke models are essential for the analysis and subsequent evaluation of novel drug targets for these mechanisms. In addition, detailed standardized models for all procedures are urgently needed to overcome the so-called replication crisis. As an effort within the ImmunoStroke research consortium, a standardized photothrombotic mouse model using an intraperitoneal injection of Rose Bengal and the illumination of the intact skull with a 561 nm laser is described. This model allows the performance of stroke in mice with allocation to any cortical region of the brain without invasive surgery; thus, enabling the study of stroke in various areas of the brain. In this video, the surgical methods of stroke induction in the photothrombotic model along with&#160;histological analysis are demonstrated.

Described here is the photothrombotic stroke model, where a stroke is produced through the intact skull by inducing permanent microvascular occlusion using laser illumination after administration of a photosensitive dye.

Stroke is a leading cause of death and acquired adult disability in developed countries. Despite extensive investigation for novel therapeutic strategies, ...

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 ...

Installation of Optical Windows in the Brain of Transgenic Mice to Image Neural Activities

Source: Lin, X., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/56297">Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice.</a> J. Vis. Exp. (2019).
This video demonstrates the brain&#39;s neural activity recording through an optical window in transgenic mice. The mice express a fluorescent protein that enables the detection of neural activities. A small portion of the skull is carefully removed to install an optical window, and the skull is then covered with black cement to block light before imaging.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

JoVE Encyclopedia of Experiments

Neurophysiology

Imaging and Optical Techniques

Implantation of Cranial Imaging Window in Mouse Model: A Surgical Procedure to Implant Glass-based Coverslip for Stable Optical Access to Regions of Murine Brain

Source: Alieva, M. et al. <a href="https://www-jove-com.bdigitaluss.remotexs.co/v/59278/longitudinal-intravital-imaging-brain-tumor-cell-behavior-response-to">Longitudinal Intravital Imaging of Brain Tumor Cell Behavior in Response to an Invasive Surgical Biopsy.</a>&#160;J. Vis. Exp.&#160; (2019)
This video presents a surgical procedure to implant cranial window in a mouse model for stable optical access to different regions of the brain.

Source: Alieva, M. et al. Longitudinal Intravital Imaging of Brain Tumor Cell Behavior in Response to an Invasive Surgical Biopsy.&#160;J. Vis. Exp.&#160; ...

Modeling Cataract Surgery in Mice

Summary

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A Mouse Model for Laser-induced Choroidal Neovascularization

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

Ultrahigh Resolution Mouse Optical Coherence Tomography to Aid Intraocular Injection in Retinal Gene Therapy Research

In Vivo Imaging of Cx3cr1^gfp/gfp Reporter Mice with Spectral-domain Optical Coherence Tomography and Scanning Laser Ophthalmoscopy

A Model of Glaucoma Induced by Circumlimbal Suture in Rats and Mice

Modeling Stroke in Mice: Focal Cortical Lesions by Photothrombosis

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

Installation of Optical Windows in the Brain of Transgenic Mice to Image Neural Activities

Implantation of Cranial Imaging Window in Mouse Model: A Surgical Procedure to Implant Glass-based Coverslip for Stable Optical Access to Regions of Murine Brain