We acknowledge our laboratory technicians Jos&#233;; Nilson dos Santos, Daianne Mandarino Torres, Jos&#233; Francisco Tib&#250;rcio, Gildo Brito de Souza, and Luciano Cavalcante Ferreira. This research was funded by FAPERJ, CNPq, and CAPES.

All procedures were performed in compliance with the Statement for the Use of Animals in Ophthalmic and Visual Research from the Association for Research in Vision and Ophthalmology (ARVO) and approved by the Ethics Committee on the Use of Animals in Scientific Experimentation from the Health Sciences Center, Federal University of Rio de Janeiro (protocol 083/17). In the present work, Lister Hooded rats of both genders were used, aged 2-3 months and weighing 180-320 g. However, the procedure can be adapted in different rat strains of various age ranges.
1. Ocular hypertension surgery and clinical follow-up
<ol>
	<li>House animals in a controlled temperature environment and 12 h light/dark cycle (6 am: light on/ 6 pm: light off) with standard pelletized gamma-irradiated food (Nuvilab&#174; CR-1, Quimtia S/A, Brazil) and water (triple filter, with non-toxic activated charcoal) available ad libitum.</li>
	<li>Prepare a stock anesthetic cocktail mixture composed of three parts of 10% ketamine hydrochloride and one part of 2% xylazine hydrochloride (diluent: sterile water).</li>
	<li>Gently restrain the animal by holding the dorsal skin and induce anesthesia by&#160;intraperitoneal administration of 1 &#181;L/g of body weight of the mixture (ketamine: 75 mg/kg; xylazine: 5 mg/kg). Check for proper anesthetization after approximately 5 min by performing a toe pinch response.</li>
	<li>Topically anesthetize both eye surfaces by instilling proxymetacaine hydrochloride 0.5% eyedrop. Wait for 30-60 s and remove the remaining solution from the anterior aspect of the globe, by gently touching the nasal or lateral bulbar conjunctiva with a small sterile cotton swab.</li>
	<li>Measure baseline IOP of both experimental and contralateral control eyes by placing the animal in a ventral decubitus position on the benchtop such that the corneal surface is easily accessible to the tip of the tonometer.</li>
	<li>Use either applanation or rebound handheld tonometer (Figure 1A). Position the tonometer tip such that it slightly touches the central corneal zone perpendicularly. Acquire and average 3-5 reliable machine-generated averages. Each mean is automatically calculated by the device after six successful individual measurements.
	<ol>
		<li>Load the rebound tonometer with a probe and press the measurement button once to turn it on, while placing the tip of the device upwards, in order to avoid the probe to fall. After turning on, the display will show 00&#160;indicating that the equipment is ready to measure.</li>
		<li>Position the device with the tip of the probe at a distance of 1-4 mm from the cornea and acquire measurements by rapidly and carefully pressing the measurement button, without moving the equipment. Each successful measure is identified by a short beep and after six times, the mean is displayed on the device&#39;s screen. 
		NOTE: Despite a long experience with applanation tonometer, to optimize the entire procedure the authors recommend the use of a rebound tonometer, which provides easier acquisition of reliable IOP measurement.</li>
	</ol>
	</li>
	<li>Place the animal on a slight lateral decubitus position under a stereo microscope, and carefully plan the surgery by inspecting the experimental eye at 40x magnification. Do not forget to maintain the contralateral control eye lubricated during the procedure by instilling a drop of carmellose sodium or sodium hyaluronate.</li>
	<li>With the aid of curved forceps, gently push forward the experimental eyeball so as to expose the vasculature that surrounds 360&#176; of the limbus13. With the other hand, gently cauterize the vessels all around the cornea with a low-temperature ophthalmic cautery (1,300 &#176;F; Bovie Medical, USA) (Figure 1B-E). 
	NOTE: The mentioned cautery has a round tip that should touch limbal vasculature longitudinally.</li>
	<li>Be careful not to cauterize the corneal periphery, as this may result in post-operative corneal opacification, which precludes in vivo assessment of retinal function.</li>
	<li>Observe the emergence of small circular marks of cauterization on the scleral limbus, the obliteration of limbal vasculature, and pupil dilation in the operated eye, which are signs of a successful surgical procedure. 
	NOTE: As the limbal vasculature of rats and mice is anatomically similar13,24&#160;and the cautery used has a gentle small tip, this protocol can work for the mouse eye as well without any adaptation.</li>
	<li>After surgery, check the immediate post-operative IOP in both eyes such as described in step 1.5.</li>
	<li>Apply a drop of ophthalmic prednisolone acetate (1.2 mg/mL) and maintain this in contact with the anterior surface of the experimental eye for around 40 s, then replace it with an ophthalmic ointment of antibiotics (oxytetracycline hydrochloride 30 mg/g plus polymyxin B 10,000 U/g; or ciprofloxacin 3.5 mg/g). Perform intramuscular injection of tramadol hydrochloride (single dose; 2 mg/kg) to prevent pain after the procedure.</li>
	<li>Follow closely the animal recovery from anesthesia, preferentially in a warm environment such as a heating pad or inside its housing cage with proper bedding. Pay attention to the respiratory pattern.</li>
	<li>Perform a daily clinical follow-up of the experimental eye with topical medications composed of a non-steroidal anti-inflammatory drug (NSAID, e.g., ketorolac trometamol 0.5% eye drop) and antibiotic ointment (preferably the same one used immediately after surgery). 
	NOTE: The use of NSAID rather than steroidal anti-inflammatory eye drops is recommended because the latter may induce OHT and is regularly applied for glucocorticoid-induced glaucoma models in rodents.
	<ol>
		<li>Under slight sedation (ketamine: 18.75 mg/kg; xylazine: 1.25 mg/kg), measure IOP preferentially at the same period of the day, to avoid bias due to physiological circadian IOP fluctuation.</li>
		<li>During the ocular examination, note rare clinical intercurrences, such as hyphema, corneal fibrosis, or scleral thinning associated with uveal prolapse. Corneal edema, chemosis, and conjunctival hyperemia are common but temporary.</li>
		<li>Notice full clinical recovery at around postoperative day 7. Limbal revascularization usually begins in the first two days after surgery, in line with the expected gradual IOP return to baseline.</li>
	</ol>
	</li>
</ol>
2. Optomotor response (OMR) analysis
NOTE: For this procedure, a specific system was used25.
<ol>
	<li>Arrange four computer monitors in a quadrangle, that delimit an arena with a platform in the middle. On the monitors&#39;, display the image of a virtual cylinder with vertically oriented sine-wave gratings (alternate black and white stripes), rotating around the platform at a fixed speed (12 degree/s) and contrast (100%).</li>
	<li>Position a video camera above the platform, allowing the experimenter to watch the animal&#39;s movements. Perform the test in photopic conditions and set the software preferentially on manual/separate mode in order to evaluate each eye separately.</li>
	<li>Allow animals to habituate for approximately 2 min on the platform. Maintain the red crosshair cursor on the video frame between the eyes of the freely moving animal, as it indicates the center of the virtual cylinder (Figure 1G)25.</li>
	<li>Observe the OMR, consisting of a reflexive tracking of the animal&#39;s head and neck elicited by the rotating gratings. Test the left and the right visual pathways by rotating the directions of the sine-wave gratings clockwise and counterclockwise, respectively.
	<ol>
		<li>Initially present a stimulus with low spatial frequency (0.042 cycles/degree). Then, increment the frequency progressively until tracking movement is not noticed anymore. The highest spatial frequency in which a clear OMR is elicited corresponds to the threshold spatial frequency of the evaluated eye.</li>
		<li>If the animal eventually drops off the platform during the exam, immediately return it to the platform and resume the test.</li>
	</ol>
	</li>
</ol>
3. Recording of pattern-electroretinogram (PERG)
NOTE: The electroretinogram was recorded using a specific system for signal processing and related software for storage and analysis of the waveforms.
<ol>
	<li>Deeply anesthetize animals by intra-muscular injection of ketamine hydrochloride and xylazine hydrochloride (75 mg/kg and 5 mg/kg, respectively). Deep anesthesia (surgical plane) decreases the chances of artifacts by involuntary muscle movements or alternative sources of noise during the exam. Check for proper anesthetization by performing a toe pinch response. 
	NOTE: The mentioned anesthetic agents do not affect the amplitude of the response26. As a small needle inserted into the cornea was used as the active electrode, intra-muscular anesthesia is preferred instead of intraperitoneal, as it is easier to reinject in case the rat needs a booster dose (1/2 of the initial dose), with lower chances to modify the electrode position and thus leading to more reproducible records during the whole experiment, which may last up to 60 min.</li>
	<li>Topically anesthetize the cornea with a drop of proxymetacaine hydrochloride 0.5%, and maintain the fellow eye moistened with ophthalmic lubricants.</li>
	<li>Carefully insert the active electrode (stainless steel needle 0.25 mm &#215; 15 mm) at the temporal periphery of the cornea. Additionally, insert the reference and ground electrodes (stainless steel needle 0.4 mm &#215; 37 mm) into the subcutaneous tissue of the ipsilateral temporal canthus and in one of the hind limbs, respectively (Figure 1I).</li>
	<li>For PERG, set the stimulus to a black and white reserving checkerboard, alternating at 15 reversals/s with constant average luminance (250 cd/m2). Set the band-pass filter to 1 Hz-100 Hz. 
	NOTE: The rapid reversing stimulus generates the steady-state PERG, a stable and reproducible sinusoid that gathers the wave component most likely associated with RGC bioelectrical response: NII deflection of the transient-state PERG.</li>
	<li>Position the animal at 20 cm from the stimulus screen (LCD monitor 0.58 m;&#160;Figure 1I), monitor signal baseline, and start PERG acquisition by pressing&#160;Analysis&#160;button on the acquisition system.&#160;</li>
	<li>During the procedure, keep the animals adapted to environmental light (white light of ~140 lux). Here, six distinct spatial frequencies (in cycles per degree: 0.018, 0.037, 0.073, 0.146, 0.292, 0.585) were presented in random sequence. 
	NOTE: The software used for signal processing automatically performs averaging. The average of 200-300 individual waves was considered adequate to stand out from the noise and analyze wave amplitude.</li>
</ol>
4. Quantification of retinal ganglion cells somas
NOTE: The following procedure is for quantification of RGC somas, based on immunohistochemical staining of retinal flat-mounts with an antibody against the brain-specific homeobox/POU domain protein 3A (Brn3a).
<ol>
	<li>Subject experimental animals to cervical dislocation euthanasia, preceded by&#160;carbon dioxide inhalation to induce unconsciousness.</li>
	<li>Immediately after euthanasia, dissect both eyes under a stereo microscope, using toothed forceps and curved scissors.</li>
	<li>Perform careful dissection such that both the distal portion of the extraocular muscles and the caruncle remain attached to the globe, as they are important landmarks for topographic orientation of the retina (described in step 4.5). Try to save a stretch of the optic nerve attached to the globe as long as possible for future analysis.</li>
	<li>After enucleation, place the eyes in a 1 mL solution of 4% paraformaldehyde (4% PFA) in 0.1 M phosphate buffer (PBS) and keep it for 24 h for proper chemical fixation.</li>
	<li>Separate the retina from the rest of the ocular tissues.
	<ol>
		<li>Under a dissecting microscope place the eyeball in a Petri dish covered with 1x PBS, and pay attention to important landmarks such as nasal caruncle, choroid fissures, and scleral imprints from vortex veins for proper topographic orientation27.</li>
		<li>Penetrate the anterior chamber from the central cornea by using toothed forceps and curved scissors (westcott). Make two radial cuts to the superior (dorsal) aspect of the sclera, toward the scleral foramen of the optic nerve, so as to delimit the dorsal quadrant of the eyeball.</li>
		<li>Separate the cornea from the rest of the globe through a longitudinal 360&#176; cut at the scleral limbus. Remove the lens and the iris and delimit the dorsal retinal quadrant using the same scleral radial demarcations described above (step 4.5.2).</li>
		<li>Carefully detach the retinal tissue from the choroid and sclera, avoiding both random lacerations throughout the tissue and eventual topographic loss of orientation.</li>
		<li>Separate the ciliary body from retinal ora serrata. Carefully remove the remaining vitreous body from the retinal cup by using both curved non-toothed forceps plus curved scissors (pull vitreous and dissect it close to the internal limiting membrane) and a small brush. 
		NOTE: Removing vitreous humor is an important step to obtain a strong and clean immunohistochemical signal.</li>
	</ol>
	</li>
	<li>Transfer isolated retinas into a 24-well culture plate (one retina per well) containing 1 mL of 1x PBS and keep the inner retina facing up.</li>
	<li>Permeabilize tissue by washing 3x for 10 min with a non-ionic surfactant 0.5% diluted in 1x PBS (0.3 mL). Then keep the tissue gently shaking in 5% bovine serum albumin (BSA) in non-ionic surfactant 2% and 1x PBS (blocking solution; 0.25 mL) for 60 min at room temperature.</li>
	<li>During step 4.7, prepare the Brn3a primary antibody solution by diluting 1:200 in non-ionic surfactant 0.5% and 1x PBS plus 5% BSA, and store it at 4 &#176;C.</li>
	<li>After 60 min of tissue blocking, incubate retinas in 0.2 mL of primary antibody solution at 4 &#176;C for 72 h with gentle shaking.</li>
	<li>Wash the tissue 3x for 10 min with 1x PBS, then incubate the tissue for 2 h at room temperature in 0.2 mL of the secondary antibody solution diluted 1:750 in 1x PBS plus 5% BSA.</li>
	<li>Further, incubate the tissue for 10 min in nuclear counterstain solution for fluorescent nuclei staining. Conclude immunohistochemistry with a final washing step with 1x PBS (0.3 mL) repeated 3x.</li>
	<li>Transfer retinas with the aid of two small brushes onto glass microscope slides, maintaining the vitreous side up. Position the dorsal retinal quadrant up on the microscope slides (previously delimited in step 4.5.3). Make two more radial cuts towards the optic nerve head to delimit the other 3 quadrants (nasal, ventral, and temporal). 
	NOTE: The cut dimensions are not fixed. They should not be too short that impairs an efficient flattening of the retina on the slide and should not be too long so that it reaches the optic nerve foramen and completely separates the delimited retinal quadrant from the rest of the tissue.</li>
	<li>Finally, apply 0.2 mL of antifade mounting medium on a glass coverslip and place this onto the flat-mounted retina for tissue microscopic analysis. To estimate RGCs density, examine the flat mounts under a confocal epifluorescence microscope using a 40x/1.3 objective.</li>
	<li>For each quadrant of the retina, take eight photos: two from the central retina (~0.9 mm from optic disc), three from mid-retina (~2.0 mm from optic disc), and three from the peripheral retina (~3.7 mm from optic disc), totaling to 32 photos per retina. Use the FIJI software to count Brn3a-positive cells and estimate the mean cell density.</li>
</ol>
5. Examination of the optic nerve
<ol>
	<li>After euthanasia and eyeball enucleation (steps 4.1-4.3), remove the proximal segment of the intraorbital portion of the optic nerve (1-2 mm), including part of the intraocular portion, and immediately place the samples into vials/tubes containing 0.2-0.3 mL of cold fixative (2.5% glutaraldehyde solution in 0.1 M sodium cacodylate buffer (pH 7.4)) for 2 h. 
	NOTE: The following steps for optic nerve processing are performed in the same vials/tubes from step 5.1</li>
	<li>Wash the material 3x for 5 min with cold 0.1 M sodium cacodylate buffer. Post fixate tissue for 1 h under gentle shaking in a solution of 1.0% osmium tetroxide in 0.8% potassium ferrocyanide and 5 nM calcium chloride diluted in 0.1 M sodium cacodylate buffer at 4 &#176;C (0.1-0.2 mL).</li>
	<li>Wash optic nerve fragments 3x for 5 min with cold 0.1 M sodium cacodylate buffer and subsequently with cold distilled water, 3x for1 min. Keep material overnight under gentle shaking in a solution of 1.0% uranyl acetate in distilled water for staining at 4 &#176;C (0.1-0.2 mL). Wash fragments 3x with cold distilled water.</li>
	<li>Progressively dehydrate the tissue with a graded acetone series (0.5 mL each), with subsequent replacements of the following dilutions in distilled water: 2x 7 min incubation in 15% ice-cold acetone; 2x 7 min incubation in 30% ice-cold acetone; 2x 7 min incubation in 50% ice-cold acetone; 2x 7 min incubation in 70% ice-cold acetone; 2x 7 min incubation in 80% ice-cold acetone; 2x 7 min incubation in 90% ice-cold acetone; 2x 15 min incubation in 100% acetone at room temperature (RT).</li>
	<li>Perform 3 infiltration/ embedding steps, with subsequent replacement of the following solutions: 1 part epoxy resin: 2 parts acetone (total volume: 0.5 mL), at RT for 12 h; 1 part epoxy resin: 1 part acetone (total volume: 0.5 mL), at RT for 12 h; 2 parts epoxy resin: 1 part acetone (total volume: 0.5 mL), at RT for 12 h. Finally, infiltrate tissue in pure epoxy resin at RT for 24 h.</li>
	<li>Remove samples from the specimen carrier, transfer them to embedding molds, and let it polymerize for 48 h at 60 &#176;C. Cut transversal semi-thin sections (300-400 nm) of the optic nerve fragments using an ultramicrotome, collect and transfer them onto a microscope glass slide.Stain sections with toluidine blue and image the using optic microscope at 100x magnification.</li>
	<li>For ultrastructural analysis, perform ultrathin cross-sections (70 nm), collect them on copper grids, and stain them with uranyl acetate and lead citrate. Examine the sections in a transmission electron microscope.</li>
</ol>

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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+three-dimensional+organisation+of+the+post-trabecular+aqueous+outflow+pathway+and+limbal+vasculature+in+the+mouse.">The three-dimensional organisation of the post-trabecular aqueous outflow pathway and limbal vasculature in the mouse.</a> Experimental Eye Research. 125, 226-235 (2014).</li><li>Prusky, G. T., Alam, N. M., Beekman, S., Douglas, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+quantification+of+adult+and+developing+mouse+spatial+vision+using+a+virtual+optomotor+system.">Rapid quantification of adult and developing mouse spatial vision using a virtual optomotor system.</a> Investigative Ophthalmology &amp; Visual Science. 45 (12), 4611-4616 (2004).</li><li>Sasovetz, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ketamine+hydrochloride:+an+effective+general+anesthetic+for+use+in+electroretinography.">Ketamine hydrochloride: an effective general anesthetic for use in electroretinography.</a> Annals of Ophthalmology. 10 (11), 1510-1514 (1978).</li><li>Stabio, M. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+map+of+the+mouse+eye+for+orienting+retinal+topography+in+anatomical+space.">A novel map of the mouse eye for orienting retinal topography in anatomical space.</a> The Journal of Comparative Neurology. 526 (11), 1749-1759 (2018).</li><li>Blanco, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Chronic+Ocular-Hypertensive+Rat+Model+induced+by+Injection+of+the+Sclerosant+Agent+Polidocanol+in+the+Aqueous+Humor+Outflow+Pathway.">A Chronic Ocular-Hypertensive Rat Model induced by Injection of the Sclerosant Agent Polidocanol in the Aqueous Humor Outflow Pathway.</a> International Journal of Molecular Sciences. 20 (13), 3209 (2019).</li><li>Paranhos, A., Prata, J. A., de Mello, P. A., da Silva, F. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Post-Trabecular+Glaucomas+with+Elevated+Episcleral+Venous+Pressure.">Post-Trabecular Glaucomas with Elevated Episcleral Venous Pressure.</a> Mechanisms of the Glaucomas. , 139-157 (2008).</li><li>Ou, Y., Jo, R. E., Ullian, E. M., Wong, R. O. L., Della Santina, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Selective+Vulnerability+of+Specific+Retinal+Ganglion+Cell+Types+and+Synapses+after+Transient+Ocular+Hypertension.">Selective Vulnerability of Specific Retinal Ganglion Cell Types and Synapses after Transient Ocular Hypertension.</a> The Journal of Neuroscience: The Official Journal of The Society for Neuroscience. 36 (35), 9240-9252 (2016).</li></ol>

The authors have nothing to disclose.

Limbal plexus cauterization (LPC) is a novel post-trabecular model with the advantage that it targets easily accessible vascular structures not requiring conjunctival or tenon dissection17,28. Differently from the vortex veins cauterization model, a renowned OHT model based on the surgical impairment to choroid venous drainage, venous congestion is not expected to influence IOP rise in the LPC model, as limbal veins are situated upstream in aqueous humor outflow....

Medical literature understands glaucoma as a heterogeneous group of optic neuropathies characterized by progressive degeneration of retinal ganglion cells (RGCs), dendrites, soma, and axons, resulting in structural cupping (excavation) of the optic disc and functional deterioration of the optic nerve, leading to amaurosis in uncontrolled cases by interrupting the transmission of visual information from the eye to the brain1. Glaucoma is currently the most common cause of irreversible blindness worldwide, predicted to reach approximately 111.8 million people in 20402, thus deeply affecting patients&#39; quality of life (QoL) and leading to significant socioeconomic concerns3.
Elevated intraocular pressure (IOP) is one of the most important and the only modifiable risk factor for the development and progression of glaucoma. Among the multiple types of glaucoma, all, except for normal tension glaucoma (NTG), are associated with elevated IOP at some time in the clinical history of the disease. Despite remarkable clinical and surgical advances to target IOP and slow down or stop disease progression, patients still lose sight due to glaucoma4,5. Therefore, a thorough understanding of the complex and multifactorial pathophysiology of this disease is imperative for the development of more effective treatments, especially to provide neuroprotection to RGCs.
Among a variety of experimental approaches for the understanding of disease mechanisms, animal models based on ocular hypertension (OHT) most closely resemble human glaucoma. Rodent models are particularly useful as they are&#160;low-cost, are easy to handle,&#160;can be genetically manipulated, have a short lifespan, and present ocular anatomical and physiological features comparable to humans, such as aqueous humor production and drainage6,7,8,9,10,11,12,13. Currently used models include sclerosis of the trabecular meshwork following injection of hypertonic saline into episcleral veins14, intracameral injection of microbeads15&#160;or viscoelastic substances16, cauterization of vortex veins17, photocoagulation of the trabecular meshwork with argon laser18, circumlimbal suture19, and use of a transgenic model of age-related OHT (DBA/2J mice)8. However, invasiveness, post-operative opacification of the cornea, anterior segment disruption, extensive learning curves, expensive equipment, and highly variable postoperative IOPs, are among few of the reported pitfalls associated with the current models, making the development of an alternative model of OHT a demand to overcome these problems20,21,22.
The present protocol formalizes a novel surgical procedure to induce OHT as a proxy to glaucoma, based on limbal plexus cauterization (LPC) in rodents23. This is an easy, reproducible, accessible, and non-invasive model that provides high efficiency and low variability of IOP elevation, associated with a uniquely high rate of full clinical recovery, therefore providing in vivo functional evaluation in a reduced number of animals used in each experiment. The surgery technique induces subacute OHT with a gradual return to baseline levels in a few days, which models the hypertensive attack seen in acute angle-closure glaucoma. Moreover, the IOP recovery in the model is followed by continuous glaucomatous neurodegeneration, which is useful for future mechanistic studies of the secondary degeneration of RGCs, which occurs in several cases of human glaucoma despite adequate control of IOP.

<table><tbody><tr><td>Acetone</td><td>Isofar</td><td>201</td><td>Used for electron microscopy tissue preparation (step 5)</td></tr><tr><td>Active electrode for electroretinography</td><td>Hansol Medical Co</td><td>-</td><td>Stainless steel needle 0.25 mm &amp;times; 15 mm</td></tr><tr><td>Anestalcon</td><td>Novartis Bioci&amp;ecirc;ncias S/A</td><td>MS-1.0068.1087</td><td>Proxymetacaine hydrochloride 0.5%</td></tr><tr><td>Calcium chloride</td><td>Vetec</td><td>560</td><td>Used for electron microscopy tissue preparation (step 5)</td></tr><tr><td>Cautery Low Temp Fine Tip 10/bx</td><td>Bovie Medical Corporation</td><td>AA00</td><td>Low-temperature ophthalmic cautery</td></tr><tr><td>Cetamin</td><td>Syntec do Brasil Ltda</td><td>000200-3-000003</td><td>Ketamine hydrochloride 10%</td></tr><tr><td>DAKO</td><td>Dako North America</td><td>S3023</td><td>Antifade mounting medium</td></tr><tr><td>DAPI</td><td>Thermo Fisher Scientific</td><td>28718-90-3</td><td>diamidino-2-phenylindole; blue fluorescent nuclear counterstain; emission at 452&amp;plusmn;3 nm</td></tr><tr><td>Ecofilm</td><td>Crist&amp;aacute;lia Produtos Qu&amp;iacute;micos Farmac&amp;ecirc;uticos Ltda</td><td>MS-1.0298.0487</td><td>Carmellose sodium 0.5%</td></tr><tr><td>EPON Resin</td><td>Polysciences, Inc.</td><td>-</td><td>Epoxy resin used for electron microscopy, composed of a mixture of four reagents: Poly/Bed 812 Resin (CAT#08791); DDSA - Dodecenylsuccinic Anhydride (CAT#00563); NMA - Nadic Methyl Anhydride (CAT#00886); DMP-30 - 2,4,6-tris(dimethylaminomethyl)phenol (CAT#00553)</td></tr><tr><td>Glutaraldehyde</td><td>Electron Microscopy Sciences</td><td>16110</td><td>Used for electron microscopy tissue preparation (step 5)</td></tr><tr><td>Hyabak</td><td>Uni&amp;atilde;o Qu&amp;iacute;mica Farmac&amp;ecirc;utica Nacional S/A</td><td>MS-8042140002</td><td>Sodium hyaluronate 0.15%</td></tr><tr><td>Icare Tonolab</td><td>Icare Finland Oy</td><td>TV02 (model number)</td><td>Rebound handheld tonometer</td></tr><tr><td>IgG donkey anti-mouse antibody + Alexa Fluor 555</td><td>Thermo Fisher Scientific</td><td>A31570</td><td>Secondary antibody solution</td></tr><tr><td>LCD monitor 23 inches</td><td>Samsung Electronics Co. Ltd.</td><td>S23B550</td><td>Model LS23B550, for electroretinogram recording</td></tr><tr><td>LSM 510 Meta</td><td>Carl Zeiss</td><td>-</td><td>Confocal epifluorescence microscope</td></tr><tr><td>Maxiflox</td><td>Crist&amp;aacute;lia Produtos Qu&amp;iacute;micos Farmac&amp;ecirc;uticos Ltda</td><td>MS-1.0298.0489</td><td>Ciprofloxacin 3.5 mg/g</td></tr><tr><td>MEB-9400K</td><td>Nihon Kohden Corporation</td><td>-</td><td>System for electroretinogram recording</td></tr><tr><td>monoclonal IgG1 mouse anti-Brn3a</td><td>MilliporeSigma</td><td>MAB-1585</td><td>Brn3a primary antibody solution</td></tr><tr><td>Neuropack Manager v08.33</td><td>Nihon Kohden Corporation</td><td>-</td><td>Software for electroretinogram signal processing</td></tr><tr><td>Optomotry</td><td>CerebralMechanics</td><td>-</td><td>System for optomotor response analysis</td></tr><tr><td>Osmium tetroxide</td><td>Electron Microscopy Sciences</td><td>19100</td><td>Used for electron microscopy tissue preparation (step 5)</td></tr><tr><td>Potassium ferrocyanide</td><td>Electron Microscopy Sciences</td><td>20150</td><td>Used for electron microscopy tissue preparation (step 5)</td></tr><tr><td>Reference and ground electrodes for electroretinography</td><td>Chalgren Enterprises</td><td>110-63</td><td>Stainless steel needles 0.4 mm &amp;times; 37 mm</td></tr><tr><td>Sodium cacodylate buffer</td><td>Electron Microscopy Sciences</td><td>12300</td><td>Used for electron microscopy tissue preparation (step 5)</td></tr><tr><td>Ster MD</td><td>Uni&amp;atilde;o Qu&amp;iacute;mica Farmac&amp;ecirc;utica Nacional S/A</td><td>MS-1.0497.1287</td><td>Prednisolone acetate 0.12%</td></tr><tr><td>Terolac</td><td>Crist&amp;aacute;lia Produtos Qu&amp;iacute;micos Farmac&amp;ecirc;uticos Ltda</td><td>MS-1.0497.1286</td><td>Ketorolac trometamol 0.5%</td></tr><tr><td>Terramicina</td><td>Laborat&amp;oacute;rios Pfizer Ltda</td><td>MS-1.0216.0024</td><td>Oxytetracycline hydrochloride 30 mg/g + polymyxin B 10,000 U/g</td></tr><tr><td>Tono-Pen XL</td><td>Reichert Technologies</td><td>230635</td><td>Digital applanation handheld tonometer</td></tr><tr><td>TO-PRO-3</td><td>Thermo Fisher Scientific</td><td>T3605</td><td>Far red-fluorescent nuclear counterstain; emission at 661 nm</td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>9036-19-5</td><td>Non-ionic surfactant</td></tr><tr><td>Uranyl acetate</td><td>Electron Microscopy Sciences</td><td>22400</td><td>Used for electron microscopy tissue preparation (step 5)</td></tr><tr><td>Xilazin</td><td>Syntec do Brasil Ltda</td><td>7899</td><td>Xylazine hydrochloride 2%</td></tr><tr><td></td><td>Carl Zeiss</td><td>-</td><td>Stereo microscope for surgery and retinal dissection</td></tr></tbody></table>

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 quantitative variables are expressed as mean &#177; standard error of the mean (SEM). Except for the comparison of IOP dynamics between OHT and control groups (Figure 1F), statistical analysis was performed using two-way ANOVA followed by Sidak&#39;s multiple comparisons test. A p-value &#60; 0.05 was considered statistically significant.
Figure 1 illustrates surgical steps of the full-circle limbal plexus cauterization (LPC) mode...

Watch this Scientific Journal Video about Full-Circle Cauterization of Limbal Vascular Plexus for Surgically Induced Glaucoma in Rodents at JoVE.com

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

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 ...

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Research

JoVE Journal

Neuroscience

1.3K Views.  Universidade Federal do Rio de Janeiro. The goal of this protocol is to characterize a novel model of glaucomatous neurodegeneration based on 360° thermic cauterization of limbal vascular plexus, inducing subacute ocular hypertension.

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

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

Ultrasound Cyclo Plasty in Eyes with Glaucoma

Glaucoma is a chronic disease caused by the progressive degeneration of the optical nerve fibers, resulting in decreased visual field that can lead to severe visual impairment, and eventually blindness. This manuscript describes a simple, surgeon-friendly, non-incisional technique, named Ultrasound Cyclo Plasty (UCP), for reducing intraocular pressure (IOP) in glaucoma patients. The technique determines a selective coagulation necrosis of the ciliary body; in addition, the stimulation of supra-choroidal and trans-scleral portions of the uveo-scleral outflow pathway has been recently proposed. UCP shows several technical improvements in ultrasound technology compared to previous techniques, providing more precise focusing on the target zone. The procedure is performed in the operating room under peribulbar anesthesia. Briefly, the coupling cone is put in contact with the eye and the ring probe, that contains six piezoelectric transducers which produce the ultrasound beams, is inserted inside it. Their proper centering over the ocular surface represents a crucial step for the correct targeting of the ciliary body. Sterile balanced salt solution is used to fill the empty spaces to ensure ultrasound acoustic propagation. Surgical treatment consists in the sequential automatic activation of each of the six transducers, for a total duration of less than 3 min. The patient leaves the hospital 1 h after the procedure with the treated eye patched. In the present study, 10 patients with open-angle glaucoma were followed-up during at least 12 months after the procedure. IOP was reduced at each interval compared to pre-operative, as well as the number of hypotensive medications. Twenty percent of patients did not respond to the treatment, and needed subsequent surgery to better control IOP. Treatment tolerability was good, with no cases of hypotony or phthisis. The UCP procedure is simpler, faster, safer, and less invasive than traditional cyclodestructive procedures with similar results in reducing IOP.

Glaucoma is a chronic disease with progressive degeneration of optic nerve fibers resulting in decreased visual field. Elevated intraocular pressure is considered the most important and the only treatable risk factor. This manuscript describes a simple, surgeon-friendly, non-incisional technique, named Ultrasound Cyclo Plasty, for reducing intraocular pressure in glaucoma patients.

Glaucoma is a chronic disease caused by the progressive degeneration of the optical nerve fibers, resulting in decreased visual field that can lead to ...

Behavior

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

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 Simple Mechanical Procedure to Create Limbal Stem Cell Deficiency in Mouse

Limbal stem cell deficiency (LSCD) is a state of malfunction or loss of limbal epithelial stem cells, after which the corneal epithelium is replaced with conjunctiva. Patients suffer from recurrent corneal defects, pain, inflammation, and loss of vision.
Previously, a murine model of LSCD was described and compared to two other models. The goal was to produce a consistent mouse model of LSCD that both mimics the phenotype in humans and lasts long enough to make it possible to study the disease pathophysiology and to evaluate new treatments. Here, the technique is described in more detail.
A motorized tool with a rotating burr has been designed to remove the rust rings from the corneal surface or to smooth the pterygium bed in patients. It is a suitable device to create the desired LSCD model. It is a readily available, easy-to-use tool with a fine tip that makes it appropriate for working on small eyes, as in mice. Its application prevents unnecessary trauma to the eye and it does not result in unwanted injuries, as often is the case with chemical injury models. As opposed to a blunt scraper, it removes the epithelium with the basement membrane. In this protocol, the limbal area was abraded two times, and then the whole corneal epithelium was shaved from limbus to limbus. To avoid stroma injury, care was taken not to brush the corneal surface once the epithelium was already removed.

The following article provides an easy, reproducible technique to effectively create a sustainable mouse model of limbal stem cell deficiency (LSCD). This animal model is useful in testing and comparing the efficacy of treatments for limbal stem cell diseases.

Limbal stem cell deficiency (LSCD) is a state of malfunction or loss of limbal epithelial stem cells, after which the corneal epithelium is replaced with ...

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

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

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

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

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 ...

Retinal Pigment Epithelium Transplantation in a Non-human Primate Model for Degenerative Retinal Diseases

Retinal pigment epithelial (RPE) transplantation holds great promise for the treatment of inherited and acquired retinal degenerative diseases. These conditions include retinitis pigmentosa (RP) and advanced forms of age-related macular degeneration (AMD), such as geographic atrophy (GA). Together, these disorders represent a significant proportion of currently untreatable blindness globally. These unmet medical needs have generated heightened academic interest in developing methods of RPE replacement. Among the animal models commonly utilized for preclinical testing of therapeutics, the non-human primate (NHP) is the only animal model that has a macula. As it shares this anatomical similarity with the human eye, the NHP eye is an important and appropriate preclinical animal model for the development of advanced therapy medicinal products (ATMPs) such as RPE cell therapy.
This manuscript describes a method for the submacular transplantation of an RPE monolayer, cultured on a polyethylene terephthalate (PET) cell carrier, underneath the macula onto a surgically created RPE wound in immunosuppressed NHPs. The fovea-the central avascular portion of the macula-is the site of the greatest mechanical weakness during the transplantation. Foveal trauma will occur if the initial subretinal fluid injection generates an excessive force on the retina. Hence, slow injection under perfluorocarbon liquid (PFCL) vitreous tamponade is recommended with a dual-bore subretinal injection cannula at low intraocular pressure (IOP) settings to create a retinal bleb. 
Pretreatment with an intravitreal plasminogen injection to release parafoveal RPE-photoreceptor adhesions is also advised. These combined strategies can reduce the likelihood of foveal tears when compared to conventional techniques. The NHP is a key animal model in the preclinical phase of RPE cell therapy development. This protocol addresses the technical challenges associated with the delivery of RPE cellular therapy in the NHP eye.

The non-human primate (NHP) is an ideal model for studying human retinal cellular therapeutics due to the anatomical and genetic similarities. This manuscript describes a method for submacular transplantation of retinal pigment epithelial cells in the NHP eye and strategies to prevent intraoperative complications associated with macular manipulation.

Retinal pigment epithelial (RPE) transplantation holds great promise for the treatment of inherited and acquired retinal degenerative diseases. These ...

A Modified Biopsy Punch Method for Precise Alkali Burns in Mice

Corneal and Limbal Alkali Injury Induction Using a Punch-Trephine Technique in a Mouse Model

The cornea is critical for vision, and corneal healing after trauma is fundamental in maintaining its transparency and function. Through the study of corneal injury models, researchers aim to enhance their understanding of how the cornea heals and develop strategies to prevent and manage corneal opacities. Chemical injury is one of the most popular injury models that has extensively been studied on mice. Most previous investigators have used a flat paper soaked in sodium hydroxide to induce corneal injury. However, inducing corneal and limbal injury using flat filter paper is unreliable, since the mouse cornea is highly curved. Here, we present a new instrument, a modified biopsy punch, that enables the researchers to create a well-circumscribed, localized, and evenly distributed alkali injury to the murine cornea and limbus. This punch-trephine method enables researchers to induce an accurate and reproducible chemical burn to the entire murine cornea and limbus while leaving other structures, such as the eyelids, unaffected by the chemical. Moreover, this study introduces an enucleation technique that preserves the medial caruncle as a landmark for identifying the nasal side of the globe. The bulbar and palpebral conjunctiva, and lacrimal gland are also kept intact using this technique. Ophthalmologic examinations were performed via slit lamp biomicroscope and fluorescein staining on days 0, 1, 2, 6, 8, and 14 post-injury. Clinical, histological, and immunohistochemical findings confirmed limbal stem cell deficiency and ocular surface regeneration failure in all experimental mice. The presented alkali corneal injury model is ideal for studying limbal stem cell deficiency, corneal inflammation, and fibrosis. This method is also suitable for investigating pre-clinical and clinical efficacies of topical ophthalmologic medications on the murine corneal surface.

Author Spotlight: Advancing Research in Corneal Opacity Treatment and Regeneration 

This protocol describes a method to induce an accurate and reproducible corneal and limbal alkali injury in a mouse model. The protocol is advantageous as it allows for an evenly distributed injury to the highly curved mouse cornea and limbus.

The cornea is critical for vision, and corneal healing after trauma is fundamental in maintaining its transparency and function. Through the study of ...

 Induction of Elevated Intraocular Pressure in a Rat Model

<a href='https://app.jove.com/v/63442/full-circle-cauterization-limbal-vascular-plexus-for-surgically'>Source: Lani-Louzada, R., et al., Full-Circle Cauterization of Limbal Vascular Plexus for Surgically Induced Glaucoma in Rodents. J. Vis. Exp. (2022)</a>This video demonstrates the induction and monitoring of elevated intraocular pressure (IOP) in a rat model. The process involves cauterizing the limbal blood vessels, resulting in vessel occlusion and elevated IOP. The rat is then allowed to recover, and IOP is regularly monitored in both the experimental and control eyes for comparison.

Source: Lani-Louzada, R., et al., Full-Circle Cauterization of Limbal Vascular Plexus for Surgically Induced Glaucoma in Rodents. J. Vis. Exp. (2022)This ...

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

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Corneal and Limbal Alkali Injury Induction Using a Punch-Trephine Technique in a Mouse Model

Induction of Elevated Intraocular Pressure in a Rat Model