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

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

1. 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® CR-1, Quimtia S/A, Brazil) and water (triple filter, with non-toxic activated charcoal) available ad libitum.
2. Prepare a stock anesthetic cocktail mixture composed of three parts of 10% ketamine hydrochloride and one part of 2% xylazine hydrochloride (diluent: sterile water).
3. Gently restrain the animal by holding the dorsal skin and induce anesthesia by intraperitoneal administration of 1 µL/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.
4. 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.
5. 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.
6. 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.
   1. 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 indicating that the equipment is ready to measure.
   2. 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'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.
7. 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.
8. With the aid of curved forceps, gently push forward the experimental eyeball so as to expose the vasculature that surrounds 360° of the limbus¹³. With the other hand, gently cauterize the vessels all around the cornea with a low-temperature ophthalmic cautery (1,300 °F; Bovie Medical, USA) (Figure 1B-E).
   NOTE: The mentioned cautery has a round tip that should touch limbal vasculature longitudinally.
9. 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.
10. 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 similar^13,24 and the cautery used has a gentle small tip, this protocol can work for the mouse eye as well without any adaptation.
11. After surgery, check the immediate post-operative IOP in both eyes such as described in step 1.5.
12. 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.
13. 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.
14. 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.
    1. 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.
    2. 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.
    3. 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.

2. Optomotor response (OMR) analysis

NOTE: For this procedure, a specific system was used²⁵.

1. Arrange four computer monitors in a quadrangle, that delimit an arena with a platform in the middle. On the monitors', 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%).
2. Position a video camera above the platform, allowing the experimenter to watch the animal's movements. Perform the test in photopic conditions and set the software preferentially on manual/separate mode in order to evaluate each eye separately.
3. 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.
4. Observe the OMR, consisting of a reflexive tracking of the animal'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.
   1. 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.
   2. If the animal eventually drops off the platform during the exam, immediately return it to the platform and resume the test.

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.

1. 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.
2. Topically anesthetize the cornea with a drop of proxymetacaine hydrochloride 0.5%, and maintain the fellow eye moistened with ophthalmic lubricants.
3. Carefully insert the active electrode (stainless steel needle 0.25 mm × 15 mm) at the temporal periphery of the cornea. Additionally, insert the reference and ground electrodes (stainless steel needle 0.4 mm × 37 mm) into the subcutaneous tissue of the ipsilateral temporal canthus and in one of the hind limbs, respectively (Figure 1I).
4. 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.
5. Position the animal at 20 cm from the stimulus screen (LCD monitor 0.58 m; Figure 1I), monitor signal baseline, and start PERG acquisition by pressing Analysis button on the acquisition system. 
6. 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.

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).

1. Subject experimental animals to cervical dislocation euthanasia, preceded by carbon dioxide inhalation to induce unconsciousness.
2. Immediately after euthanasia, dissect both eyes under a stereo microscope, using toothed forceps and curved scissors.
3. 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.
4. 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.
5. Separate the retina from the rest of the ocular tissues.
   1. 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.
   2. 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.
   3. Separate the cornea from the rest of the globe through a longitudinal 360° 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).
   4. Carefully detach the retinal tissue from the choroid and sclera, avoiding both random lacerations throughout the tissue and eventual topographic loss of orientation.
   5. 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.
6. 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.
7. 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.
8. 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 °C.
9. After 60 min of tissue blocking, incubate retinas in 0.2 mL of primary antibody solution at 4 °C for 72 h with gentle shaking.
10. 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.
11. 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.
12. 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.
13. 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.
14. 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.

5. Examination of the optic nerve

1. 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
2. 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 °C (0.1-0.2 mL).
3. 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 °C (0.1-0.2 mL). Wash fragments 3x with cold distilled water.
4. 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).
5. 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.
6. Remove samples from the specimen carrier, transfer them to embedding molds, and let it polymerize for 48 h at 60 °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.
7. 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.