This work was supported by research grants from the National Institutes of Health (EY022383 and EY022683; EJD) and Core grant (P30EY001765), Imaging and Microscopy Core Module.

Ethics Statement: All procedures were performed in accordance with the guidelines set forth by the Johns Hopkins University Institutional Animal Care and Use Committee.
Note: Mice used during filming are C57BL/6 mice from Jackson, although other rodent strains or species may also be used. When using other strains or species, be aware that anesthesia dosages and injury timeline may vary. It is important to adapt I/R conditions to accommodate strain, species, and experimental variations.
1. Prepare the Anesthesia Cocktail
<ol>
	<li>Combine 1.25 ml Ketamine, 0.625 ml Xylazine, 0.375 ml Acepromazine, and 22.75 ml phosphate buffered saline in a 50 ml centrifuge tube. 
	NOTE: For the remainder of the manuscript, this solution will be referred to as the cocktail.</li>
	<li>Filter the cocktail into a new sterile 50 ml centrifuge tube using a 60 ml syringe and a 0.20 &#181;m filter. Label and date this new tube.</li>
	<li>Fully wrap the cocktail tube in aluminum foil to prevent light-induced degradation of the anesthetic.&#160; 
	NOTE: The cocktail may be stored at room temperature and reused until the expiration date of its earliest-expiring ingredient.</li>
</ol>
2. Prepare the Anesthesia Booster
<ol>
	<li>In a 50 ml centrifuge tube, combine 4 ml Ketamine and 16 ml phosphate buffered saline. 
	NOTE: This solution will henceforth be referred to as the booster.</li>
	<li>Filter the booster into a new sterile 50 ml centrifuge tube using a 60 ml syringe and a 0.20 &#181;m filter. Label and date this new tube.&#160; 
	NOTE: The booster may be stored at room temperature and reused until the expiration date of its earliest-expiring ingredient.</li>
</ol>
3. Prepare the Surgical Suite
<ol>
	<li>Set the room temperature between 18 &#176;C and 21 &#176;C.</li>
	<li>Turn on the surgery table, and adjust its surface heat to the highest temperature.</li>
	<li>Cover all work surfaces with surgical underpads.</li>
	<li>Arrange an empty cage or other container on the surgical table to warm.</li>
	<li>Arrange a scale and an ear tagger on the worktop.</li>
</ol>
4. Prepare the Balanced Salt Solution with Heparin Sodium
<ol>
	<li>Add 0.5 ml of heparin sodium to a 500 ml IV bottle of balanced salt solution (HBSS).</li>
	<li>Insert the sharp end of the primary set prepierced Y-site tubing into the bottle of 0.1% heparin sodium.</li>
</ol>
5. Set up the IV Pole
<ol>
	<li>Hang the bottle of 0.1% heparin sodium from the IV pole extension, and snap open the air filter cap on the primary set prepierced Y-site tubing.</li>
	<li>Elevate the 0.1% heparin sodium bottle to 163 cm (120 mm Hg). Measure the height from the tabletop to the peak of the sodium heparin drip.</li>
	<li>Remove any air bubbles in the IV tubing by manually flicking the primary set prepierced Y-site tubing.</li>
</ol>
6. Set up the Sodium Heparin Drip
<ol>
	<li>Connect the primary set prepierced Y-site tubing to the five-valve manifold.</li>
	<li>Connect 30-gauge &#189; inch needles to the five-port manifold using Luer male to Luer male tube fittings.</li>
	<li>Insert each of the 30-gauge &#189; inch needles into its own 10-inch segment of 30-gauge tubing.</li>
	<li>Using hemostats, break the needle tips from new 30-gauge &#189; inch needles and insert their blunt ends into the 30-gauge tubing.&#160;Sterility or disinfection of the needle tips will be necessary for Step 8.3.</li>
	<li>Using tape, arrange the 30-gauge tubes with needles such that the tubes connected to the inner ports are positioned on top of the tubes connected to the outer ports. This arrangement will prevent tangling of tubing during anterior chamber cannulation.</li>
	<li>Turn on the 0.1% heparin sodium flow to the five-valve manifold and to each individual port.
	<ol>
		<li>Ensure that the 0.1% heparin sodium is flowing strongly for each port. If a port is flowing weakly, replace the port or clear it with air from a sterile syringe.</li>
		<li>Allow the 0.1% heparin sodium to flow for 2 - 3 min to remove air bubbles from the 30-gauge tubes and 5-valve manifold.</li>
	</ol>
	</li>
	<li>Turn off all the ports on the 5-valve manifold.</li>
</ol>
7. Prepare the Mice for Surgery
<ol>
	<li>Bring the mice to the surgical suite. Supply water bottles for each cage in order to prevent animal dehydration during surgery.</li>
	<li>Record the weight of each mouse.</li>
	<li>Inject each mouse intraperitoneally&#160;with 0.02 ml cocktail per gram bodyweight.</li>
	<li>Tag and record the number of each mouse.</li>
	<li>Place all the mice into the empty container on the surgery table. Allow 5-10 min for all the mice to achieve deep anesthetization as confirmed by absence of the pedal withdrawal response to toe pinch.</li>
	<li>For each mouse, administer one drop of Tropicamide into each eye for pupil dilation and short-term lubrication.</li>
	<li>For each mouse, administer one drop of Proparacaine into each eye for local anesthesia and short-term lubrication.</li>
	<li>Arrange the mice in the order of anesthetization so that the first mouse to be anesthetized will be the first mouse to be cannulated.</li>
	<li>Allow approximately 2 min for the eye drops to take effect.</li>
	<li>Prepare straight 4-inch pieces of tape by pulling tightly on the tape. Set aside one piece of tape for each mouse.&#160;&#160;&#160;&#160; 
	NOTE: Failing to pull tightly on the tape will result in curling of the tape.</li>
</ol>
8. Cannulate the Anterior Chamber
<ol>
	<li>Arrange the first mouse under the surgical microscope, and focus the microscope on the preferred cornea.&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; 
	NOTE: Cannulation may be performed on either eye. Right-hand dominant surgeons may find it easiest to cannulate the left eye, while left-hand dominant surgeons may prefer the right.</li>
	<li>Turn on the first 0.1% heparin sodium port on the five-valve manifold.</li>
	<li>Under the surgical microscope, use a pair of forceps to gently proptose the eye. Insert the 30-gauge cannula needle into the anterior chamber approximately halfway between the zonule fibers and the apex of the cornea.
	<ol>
		<li>Take care to avoid scratching or puncturing the iris, lens, or inner corneal surface.</li>
		<li>Avoid penetrating the cornea a second time.</li>
		<li>Using a gentle twisting motion to overcome friction between the cannula and the cornea, insert the cannula deeply in the anterior chamber.</li>
	</ol>
	</li>
	<li>Use a strip of tape to secure the 30-gauge tubing to the table. To minimize movement of the inserted cannula, press the 30-gauge tubing against the tabletop while reaching for the tape.</li>
	<li>Record the start time of the surgery.</li>
	<li>Using the microscope, verify that no leakage is apparent. If leakage is present, the movement of fluid will be visible near the eye.</li>
	<li>Visually confirm ocular distention by observing that the I/R eye is larger than the contralateral eye. Together, the absence of leakage and the presence of ocular distention demonstrate a successful elevation of intraocular pressure.</li>
	<li>Apply hypromellose to both eyes. Hypromellose serves to lubricate the cornea and seal microleaks. Reapply hypromellose as needed (approximately every 30 min) for sustained lubrication.</li>
	<li>Repeat Step 8 until all animals have been cannulated.</li>
</ol>
9. Monitor Anesthesia
<ol>
	<li>Use the toe pinch, visual observation of the whiskers, or visual observation of the tail to verify that each mouse remains anesthetized. Should a mouse demonstrate a pedal withdrawal response, whisker twitching, or tail movement, proceed immediately to Step 9.2.</li>
	<li>If a mouse begins to awaken during surgery, lift the tail to inject 0.05 ml booster intraperitoneally into the lower abdomen from behind the mouse. Allow 1 - 2 min for the booster to take effect.</li>
	<li>Should a mouse&#160;require additional sedation, repeat Step 9 as necessary.</li>
</ol>
10. Remove the Cannula&#160;from the Anterior Chamber
<ol>
	<li>For each animal, after 90 min have elapsed, gently pull the cannula from the anterior chamber.</li>
	<li>Untape the mouse from the surgical table, taking care not to disturb the cannula tubing of adjacent animals.</li>
	<li>Lubricate both eyes with lubricating jelly.</li>
	<li>As subsequent cannulae are removed, arrange the mice in the empty container on the surgical table to recover from surgery. Do not turn off the heat of the surgical table.</li>
	<li>Allow, at minimum, 2 - 3 hr for recovery on the heated surgical table.&#160;Observe the mice frequently until they have fully recovered from the anesthesia.</li>
</ol>
11. Clean the Equipment
<ol>
	<li>Disinfect&#160;the cannulae using alcohol wipes.&#160; 
	NOTE: Other methods for disinfection or sterilization, such as autoclaving, may be substituted.</li>
	<li>Expel 0.1% heparin sodium from the 5-valve manifold, 30-gauge needles, 30-gauge tubing, and cannulae using a 60 ml syringe filled with air.</li>
	<li>Rinse the 5-valve manifold, 30-gauge needles, 30-gauge tubing, and cannulae using a 60 ml syringe filled with distilled water.</li>
	<li>Expel distilled water from the 5-port manifold, 30-gauge needles, 30-gauge tubing, and cannulae using a 60 ml syringe filled with air.</li>
	<li>After disinfecting&#160;the cannulae and rinsing the tubing apparatus, store this equipment for reuse.</li>
	<li>Store, discard, or turn off all other equipment. Leave the heat of the surgical table on.</li>
</ol>
12. Return All Mice to Their Home Cages
<ol>
	<li>After the mice awaken from surgery (after 2 - 3 hr), return each animal to its home cage. Provide gel food for each cage.&#160;Return all cages to their designated rooms.</li>
	<li>Turn off the heat of the surgical table. Discard all waste, and wipe down the surgical suite.</li>
</ol>
13. Perform Retinal Assessment
<ol>
	<li>As appropriate, collect retinas for histological analysis or dark adapt mice for electroretinogram recording.</li>
</ol>

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The authors have no disclosures.

Retinal I/R&#160;injury by elevated IOP has proven its utility in modeling cellular damage and dysfunction, particularly neurodegeneration, in the rodent retinal neurovascular unit. This procedure provides a robust control tissue and is easily accessible in terms of technical sophistication. It has been noted in this and other I/R injury models that increasing the pressure and duration of ischemia may increase injury severity24. For this reason, some practitioners have elected to use ischemic pressures and dur...

Retinal ischemia-reperfusion (I/R) characterizes many human retinal pathologies, including glaucoma, diabetic retinopathy, and retinal vascular occlusions1. In retinal I/R, reduced blood flow (ischemia) in the retinal vasculature creates a state of retinal hypersensitivity to oxygen and other nutrients, precipitating severe oxidative and inflammatory damage when circulation is subsequently reinstated (reperfusion)2. The neural retina appears particularly vulnerable to these changes, with retinal neurodegeneration being perhaps the most salient feature of I/R-induced damage. Presented here is a protocol for modeling retinal I/R injury in the mouse. This technique enables researchers to examine potential mechanisms and treatment strategies for human diseases of the retinal neurovascular unit.
Pioneered in 1952 by surgeons seeking to understand the neurodegenerative consequences of surgical anemia3, rodent retinal I/R by elevated intraocular pressure (IOP) was reestablished in 1991 for the purpose of standardizing neurodegenerative endpoints after ischemic insult4. Using the drip of a saline reservoir to raise IOP above systolic blood pressure, these studies demonstrated that pressurized ocular cannulation was sufficient to suspend the retinal circulation and thereby initiate neuronal degeneration. More recent efforts using retinal I/R by elevated IOP have begun to elaborate the mechanisms underlying I/R-induced retinal neurodegeneration5-12. Multiple groups have reported additional pathologic changes including inflammation13,14, vascular permeability15,16, and capillary degeneration14,17. Taken together, these studies have established retinal I/R injury by elevated IOP as a model of retinal neurovascular disease more generally.
Characterizing the mechanisms of I/R injury is essential for the study of vascular disease. Retinal I/R injury by elevated IOP is one of many hypoxia-induced injury models, including I/R injuries in lung18, heart19, brain20, liver21, kidney22, and intestine23. These models have been paramount in advancing our understanding of vascular illness and its clinical remedies. By extending the investigation of I/R processes to ocular tissues, retinal I/R injury by elevated IOP helps to paint a more comprehensive picture of these related conditions.
Corresponding closely with clinical neurodegenerative conditions in retina, retinal I/R injury by elevated IOP presents a valuable tool for researchers interested in exploring ischemic pathogenesis. The protocol described herein is targeted, tractable, and accessible. It is complemented well by endpoints in neuronal degeneration, such as quantification of retinal neurons, measurement of retinal thickness, and electrophysiological recording of retinal neuron function. This model has proven its utility in advancing neurovascular inquiry, and it shows promise in earning status as a foundational protocol in visual medicine research.

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a mouse model of retinal ischemia-reperfusion injury through elevation of intraocular pressure

Retinal ischemia-reperfusion (I/R) is a pathophysiological process contributing to cellular damage in multiple ocular conditions, including glaucoma, diabetic retinopathy, and retinal vascular occlusions. Rodent models of I/R injury are providing significant insights into mechanisms and treatment strategies for human I/R injury, especially with regard to neurodegenerative damage in the retinal neurovascular unit. Presented here is a protocol for inducing retinal I/R injury in mice through elevation of intraocular pressure (IOP). In this protocol, the ocular anterior chamber is cannulated with a needle, through which flows the drip of an elevated saline reservoir. Using this drip to raise IOP above systolic arterial blood pressure, a practitioner temporarily halts inner retinal blood flow (ischemia). When circulation is reinstated (reperfusion) by removal of the cannula, severe cellular damage ensues, resulting ultimately in retinal neurodegeneration. Recent studies demonstrate inflammation, vascular permeability, and capillary degeneration as additional elements of this model. Compared to alternative retinal I/R methodologies, such as retinal arterial ligation, retinal I/R injury by elevated IOP offers advantages in its&#160;anatomical specificity, experimental tractability, and technical accessibility, presenting itself as a valuable tool for examining neuronal pathogenesis and therapy in the retinal neurovascular unit.

The neurodegenerative effects of retinal I/R by elevated IOP are commonly evaluated using two standard approaches. NeuN immunolabeling of neuronal nuclei has revealed significant neuronal cell loss following I/R insult (Figure 1). Briefly, eyes enucleated 7 days after I/R were fixed in paraformaldehyde, labeled with the neuronal cell marker NeuN, and whole-mounted. Images were captured using confocal microscopy, and cells labeled with NeuN were quantified by counting...

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A Mouse Model of Retinal Ischemia-Reperfusion Injury Through Elevation of Intraocular Pressure

This article describes a procedure for inducing retinal ischemia-reperfusion injury by elevated intraocular pressure in mice. Retinal ischemia-reperfusion injury by elevated intraocular pressure serves to model human pathologies characterized by compromised oxygen and nutrient delivery in the retina, enabling researchers to examine potential cellular mechanisms and treatments for human diseases of the retinal neurovascular unit.

Retinal ischemia-reperfusion (I/R) is a pathophysiological process contributing to cellular damage in multiple ocular conditions, including glaucoma, ...

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Universidad de Cartagena UDEC

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JoVE Journal

Medicine

14.8K Views.  Johns Hopkins University. This article describes a procedure for inducing retinal ischemia-reperfusion injury by elevated intraocular pressure in mice. Retinal ischemia-reperfusion injury by elevated intraocular pressure serves to model human pathologies characterized by compromised oxygen and nutrient delivery in the retina, enabling researchers to examine potential cellular mechanisms and treatments for human diseases of the retinal neurovascular unit.

Video: A Mouse Model of Retinal Ischemia-Reperfusion Injury Through Elevation of Intraocular Pressure

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 ...

Glaucoma-inducing Procedure in an In Vivo Rat Model and Whole-mount Retina Preparation

Glaucoma is a disease of the central nervous system affecting retinal ganglion cells (RGCs). RGC axons making up the optic nerve carry visual input to the brain for visual perception. Damage to RGCs and their axons leads to vision loss and/or blindness. Although the specific cause of glaucoma is unknown, the primary risk factor for the disease is an elevated intraocular pressure. Glaucoma-inducing procedures in animal models are a valuable tool to researchers studying the mechanism of RGC death. Such information can lead to the development of effective neuroprotective treatments that could aid in the prevention of vision loss. The protocol in this paper describes a method of inducing glaucoma - like conditions in an in vivo rat model where 50 &#181;l of 2 M hypertonic saline is injected into the episcleral venous plexus. Blanching of the vessels indicates successful injection. This procedure causes loss of RGCs to simulate glaucoma. One month following injection, animals are sacrificed and eyes are removed. Next, the cornea, lens, and vitreous are removed to make an eyecup. The retina is then peeled from the back of the eye and pinned onto sylgard dishes using cactus needles. At this point, neurons in the retina can be stained for analysis. Results from this lab show that approximately 25% of RGCs are lost within one month of the procedure when compared to internal controls. This procedure allows for quantitative analysis of retinal ganglion cell death in an in vivo rat glaucoma model.

Glaucoma-inducing Procedure in an In Vivo Rat Model and Whole-mount Retina Preparation

Glaucoma is characterized by damage to retinal ganglion cells. Inducing glaucoma in animal models can provide insight into the study of this disease. Here, we outline a procedure that induces loss of RGCs in an in vivo rat model and demonstrates the preparation of whole-mount retinas for analysis.

Glaucoma is a disease of the central nervous system affecting retinal ganglion cells (RGCs). RGC axons making up the optic nerve carry visual input to 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 ...

A Reversible Silicon Oil-Induced Ocular Hypertension Model in Mice

Elevated intraocular pressure (IOP) is a well-documented risk factor for glaucoma. Here we describe a novel, effective method for consistently inducing stable IOP elevation in mice that mimics the post-operative complication of using silicone oil (SO) as a tamponade agent in human vitreoretinal surgery. In this protocol, SO is injected into the anterior chamber of the mouse eye to block the pupil and prevent inflow of aqueous humor. The posterior chamber accumulates aqueous humor and this in turn increases the IOP of the posterior segment. A single SO injection produces reliable, sufficient, and stable IOP elevation, which induces significant glaucomatous neurodegeneration. This model is a true replicate of secondary glaucoma in the eye clinic. To further mimic the clinical setting, SO can be removed from the anterior chamber to reopen the drainage pathway and allow inflow of aqueous humor, which is drained through the trabecular meshwork (TM) at the angle of the anterior chamber. Because IOP quickly returns to normal, the model can be used to test the effect of lowering IOP on glaucomatous retinal ganglion cells. This method is straightforward, does not require special equipment or repeat procedures, closely simulates clinical situations, and may be applicable to diverse animal species. However, minor modifications may be required.

Here, we present a protocol to induce ocular hypertension and glaucomatous neurodegeneration in mouse eyes by intracameral injection of silicone oil and the procedure for silicone oil removal from the anterior chamber to return elevated intraocular pressure to normal.

Elevated intraocular pressure (IOP) is a well-documented risk factor for glaucoma. Here we describe a novel, effective method for consistently inducing ...

Oxygen-Induced Retinopathy Model for Ischemic Retinal Diseases in Rodents

One of the commonly used models for ischemic retinopathies is the oxygen-induced retinopathy (OIR) model. Here we describe detailed protocols for the OIR model induction and its readouts in both mice and rats. Retinal neovascularization is induced in OIR by exposing rodent pups either to hyperoxia (mice) or alternating levels of hyperoxia and hypoxia (rats). The primary readouts of these models are the size of neovascular (NV) and avascular (AVA) areas in the retina. This preclinical in vivo model can be used to evaluate the efficacy of potential anti-angiogenic drugs or to address the role of specific genes in the retinal angiogenesis by using genetically manipulated animals. The model has some strain and vendor specific variation in the OIR induction which should be taken into consideration when designing the experiments.

Oxygen-induced retinopathy (OIR) can be used to model ischemic retinal diseases such as retinopathy of prematurity and proliferative diabetic retinopathy and to serve as a model for proof-of-concept studies in evaluating antiangiogenic drugs for neovascular diseases. OIR induces robust and reproducible neovascularization in the retina that can be quantified.

One of the commonly used models for ischemic retinopathies is the oxygen-induced retinopathy (OIR) model. Here we describe detailed protocols for the OIR ...

Full-Circle Cauterization of Limbal Vascular Plexus for Surgically Induced Glaucoma in Rodents

Glaucoma, the second leading cause of blindness worldwide, is a heterogeneous group of ocular disorders characterized by structural damage to the optic nerve and retinal ganglion cell (RGC) degeneration, resulting in visual dysfunction by interrupting the transmission of visual information from the eye to the brain. Elevated intraocular pressure is the most important risk factor; thus, several models of ocular hypertension have been developed in rodents by either genetic or experimental approaches to investigate the causes and effects of the disease. Among those, some limitations have been reported such as surgical invasiveness, inadequate functional assessment, requirement of extensive training, and highly variable extension of retinal damage. The present work characterizes a simple, low-cost, and efficient method to induce ocular hypertension in rodents, based on low-temperature, full-circle cauterization of the limbal vascular plexus, a major component of aqueous humor drainage. The new model provides a technically easy, noninvasive, and reproducible subacute ocular hypertension, associated with progressive RGC and optic nerve degeneration, and a unique post-operative clinical recovery rate that allows in vivo functional studies by both electrophysiological and behavioral methods.

The goal of this protocol is to characterize a novel model of glaucomatous neurodegeneration based on 360&#176; thermic cauterization of limbal vascular plexus, inducing subacute ocular hypertension.

Glaucoma, the second leading cause of blindness worldwide, is a heterogeneous group of ocular disorders characterized by structural damage to the optic ...

Neuroscience

A Murine Model of Ischemic Retinal Injury Induced by Transient Bilateral Common Carotid Artery Occlusion

Diverse vascular diseases such as diabetic retinopathy, occlusion of retinal veins or arteries and ocular ischemic syndrome can lead to retinal ischemia. To investigate pathological mechanisms of retinal ischemia, relevant experimental models need to be developed. Anatomically, a main retinal blood supplying vessel is the ophthalmic artery (OpA) and OpA originates from the internal carotid artery of the common carotid artery (CCA). Thus, disruption of CCA could effectively cause retinal ischemia. Here, we established a mouse model of retinal ischemia by transient bilateral common carotid artery occlusion (tBCCAO) to tie the right CCA with 6-0 silk sutures and to occlude the left CCA transiently for 2 seconds via a clamp, and showed that tBCCAO could induce acute retinal ischemia leading to retinal dysfunction. The current method reduces reliance on surgical instruments by only using surgical needles and a clamp, shortens occlusion time to minimize unexpected animal death, which is often seen in mouse models of middle cerebral artery occlusion, and maintains reproducibility of common retinal ischemic findings. The model can be utilized to investigate the pathophysiology of ischemic retinopathies in mice and further can be used for in vivo drug screening.

Here, we describe a mouse model of retinal ischemia by transient bilateral common carotid artery occlusion using simple sutures and a clamp. This model can be useful for understanding the pathological mechanisms of retinal ischemia caused by cardiovascular abnormalities.

Diverse vascular diseases such as diabetic retinopathy, occlusion of retinal veins or arteries and ocular ischemic syndrome can lead to retinal ischemia. ...

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), ...

Mouse Model for Retinal Ischemia-Reperfusion Injury to Study Retinal Pathologies

Induction of Retinal Ischemia-Reperfusion Injury in a Mouse Eye Model

Ischemia-reperfusion injuries are known to cause a range of retinal pathologies, including diabetic retinopathy, glaucoma, retinal vascular occlusions, and other vaso-occlusive conditions. This manuscript presents a method for inducing ischemia-reperfusion injury in a mouse model. The method utilized anterior chamber cannulation attached to a saline reservoir, generating hydrostatic pressure to raise the intraocular pressure to 90-100 mmHg. This method effectively caused constriction of retinal capillaries to induce retinal ischemia. At the end of the ischemic period (60 min), the intraocular pressure was normalized (&#8804;20 mmHg) before removing the cannula from the anterior chamber to initiate reperfusion. Days after the ischemia/reperfusion procedure, the eyes were collected and sectioned for histological staining. The histopathology of the retinal sections was scored by evaluating eight parameters of retinal injury: folds, hemorrhage, deformation, cell loss in the ganglion cell, inner nuclear, outer nuclear, and photoreceptor layers, and damage to retinal pigment epithelial cells. This method provided a reproducible model to study the mechanisms and pathology of retinal ischemia/reperfusion injury. In addition, this model can facilitate the discovery of potential therapeutic targets to treat retinal ischemia/reperfusion injury, advancing the study of retinal pathologies and improving patient outcomes.

This protocol models retinal ischemia-reperfusion injury in a mouse eye by inducing retinal ischemia via anterior chamber cannulation and intraocular pressure elevation, followed by intraocular pressure normalization to initiate the reperfusion.

Ischemia-reperfusion injuries are known to cause a range of retinal pathologies, including diabetic retinopathy, glaucoma, retinal vascular occlusions, ...

A Mouse Model of Retinal Ischemia-Reperfusion Injury Through Elevation of Intraocular Pressure

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Laser-Induced Chronic Ocular Hypertension Model on SD Rats

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Glaucoma-inducing Procedure in an In Vivo Rat Model and Whole-mount Retina Preparation

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

A Reversible Silicon Oil-Induced Ocular Hypertension Model in Mice

Oxygen-Induced Retinopathy Model for Ischemic Retinal Diseases in Rodents

Full-Circle Cauterization of Limbal Vascular Plexus for Surgically Induced Glaucoma in Rodents

A Murine Model of Ischemic Retinal Injury Induced by Transient Bilateral Common Carotid Artery Occlusion

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

Induction of Retinal Ischemia-Reperfusion Injury in a Mouse Eye Model