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.

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			<td>Acetone</td>
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			<td>201</td>
			<td>Used for electron microscopy tissue preparation (step 5)</td>
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			<td>Active electrode for electroretinography</td>
			<td>Hansol Medical Co</td>
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			<td>Stainless steel needle 0.25 mm &#215; 15 mm</td>
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			<td>Anestalcon</td>
			<td>Novartis Bioci&#234;ncias S/A</td>
			<td>MS-1.0068.1087</td>
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			<td>Calcium chloride</td>
			<td>Vetec</td>
			<td>560</td>
			<td>Used for electron microscopy tissue preparation (step 5)</td>
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			<td>Cautery Low Temp Fine Tip 10/bx</td>
			<td>Bovie Medical Corporation</td>
			<td>AA00</td>
			<td>Low-temperature ophthalmic cautery</td>
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			<td>Cetamin</td>
			<td>Syntec do Brasil Ltda</td>
			<td>000200-3-000003</td>
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			<td>DAKO</td>
			<td>Dako North America</td>
			<td>S3023</td>
			<td>Antifade mounting medium</td>
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			<td>DAPI</td>
			<td>Thermo Fisher Scientific</td>
			<td>28718-90-3</td>
			<td>diamidino-2-phenylindole; blue fluorescent nuclear counterstain; emission at 452&#177;3 nm</td>
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		<tr>
			<td>Ecofilm</td>
			<td>Crist&#225;lia Produtos Qu&#237;micos Farmac&#234;uticos Ltda</td>
			<td>MS-1.0298.0487</td>
			<td>Carmellose sodium 0.5%</td>
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		<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>
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			<td>Glutaraldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>16110</td>
			<td>Used for electron microscopy tissue preparation (step 5)</td>
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		<tr>
			<td>Hyabak</td>
			<td>Uni&#227;o Qu&#237;mica Farmac&#234;utica Nacional S/A</td>
			<td>MS-8042140002</td>
			<td>Sodium hyaluronate 0.15%</td>
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			<td>Icare Tonolab</td>
			<td>Icare Finland Oy</td>
			<td>TV02 (model number)</td>
			<td>Rebound handheld tonometer</td>
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			<td>IgG donkey anti-mouse antibody + Alexa Fluor 555</td>
			<td>Thermo Fisher Scientific</td>
			<td>A31570</td>
			<td>Secondary antibody solution</td>
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			<td>LCD monitor 23 inches</td>
			<td>Samsung Electronics Co. Ltd.</td>
			<td>S23B550</td>
			<td>Model LS23B550, for electroretinogram recording</td>
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			<td>LSM 510 Meta</td>
			<td>Carl Zeiss</td>
			<td>-</td>
			<td>Confocal epifluorescence microscope</td>
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			<td>Maxiflox</td>
			<td>Crist&#225;lia Produtos Qu&#237;micos Farmac&#234;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>
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		<tr>
			<td>monoclonal IgG1 mouse anti-Brn3a</td>
			<td>MilliporeSigma</td>
			<td>MAB-1585</td>
			<td>Brn3a primary antibody solution</td>
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		<tr>
			<td>Neuropack Manager v08.33</td>
			<td>Nihon Kohden Corporation</td>
			<td>-</td>
			<td>Software for electroretinogram signal processing</td>
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			<td>Optomotry</td>
			<td>CerebralMechanics</td>
			<td>-</td>
			<td>System for optomotor response analysis</td>
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			<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 &#215; 37 mm</td>
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		<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&#227;o Qu&#237;mica Farmac&#234;utica Nacional S/A</td>
			<td>MS-1.0497.1287</td>
			<td>Prednisolone acetate 0.12%</td>
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		<tr>
			<td>Terolac</td>
			<td>Crist&#225;lia Produtos Qu&#237;micos Farmac&#234;uticos Ltda</td>
			<td>MS-1.0497.1286</td>
			<td>Ketorolac trometamol 0.5%</td>
		</tr>
		<tr>
			<td>Terramicina</td>
			<td>Laborat&#243;rios Pfizer Ltda</td>
			<td>MS-1.0216.0024</td>
			<td>Oxytetracycline hydrochloride 30 mg/g + polymyxin B 10,000 U/g</td>
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			<td>Tono-Pen XL</td>
			<td>Reichert Technologies</td>
			<td>230635</td>
			<td>Digital applanation handheld tonometer</td>
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		<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>
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			<td>Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>9036-19-5</td>
			<td>Non-ionic surfactant</td>
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			<td>Uranyl acetate</td>
			<td>Electron Microscopy Sciences</td>
			<td>22400</td>
			<td>Used for electron microscopy tissue preparation (step 5)</td>
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		<tr>
			<td>Xilazin</td>
			<td>Syntec do Brasil Ltda</td>
			<td>7899</td>
			<td>Xylazine hydrochloride 2%</td>
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		<tr>
			<td></td>
			<td>Carl Zeiss</td>
			<td>-</td>
			<td>Stereo microscope for surgery and retinal dissection</td>
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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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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

Preparing Undercut Model of Posttraumatic Epileptogenesis in Rodents

Partially isolated cortex ("undercut") is an animal model of posttraumatic epileptogenesis. The surgical procedure involves cutting through the sensorimotor cortex and the underneath white matter (undercut) so that a specific region of the cerebral cortex is largely isolated from the neighboring cortex and subcortical regions1-3. After a latency of two or more weeks following the surgery, epileptiform discharges can be recorded in brain slices from rodents1; and electrical or behavior seizures can be observed in vivo from other species such as cat and monkey4-6. This well established animal model is efficient to generate and mimics several important characteristics of traumatic brain injury. However, it is technically challenging attempting to make precise cortical lesions in the small rodent brain with a free hand. Based on the procedure initially established in Dr. David Prince's lab at the Stanford University1, here we present an improved technique to perform a surgery for the preparation of this model in mice and rats. We demonstrate how to make a simple surgical device and use it to gain a better control of cutting depth and angle to generate more precise and consistent results. The device is easy to make, and the procedure is quick to learn. The generation of this animal model provides an efficient system for study on the mechanisms of posttraumatic epileptogenesis.

Partially isolated cortex (&#x201C;undercut&#x201D;) is an efficient animal model of posttraumatic epileptogenesis. Here we demonstrate how to make a novel surgical device and use it to make more precise and consistent lesions to generate this model.

Partially isolated cortex ("undercut") is an animal model of posttraumatic epileptogenesis. The surgical procedure involves cutting through the ...

Whole Animal Perfusion Fixation for Rodents

The goal of fixation is to rapidly and uniformly preserve tissue in a life-like state. While placing tissue directly in fixative works well for small pieces of tissue, larger specimens like the intact brain pose a problem for immersion fixation because the fixative does not reach all regions of the tissue at the same rate 5,7. Often, changes in response to hypoxia begin before the tissue can be preserved 12. The advantage of directly perfusing fixative through the circulatory system is that the chemical can quickly reach every corner of the organism using the natural vascular network. In order to utilize the circulatory system most effectively, care must be taken to match physiological pressures 3. It is important to note that physiological pressures are dependent on the species used. Techniques for perfusion fixation vary depending on the tissue to be fixed and how the tissue will be processed following fixation. In this video, we describe a low-cost, rapid, controlled and uniform fixation procedure using 4% paraformaldehyde perfused via the vascular system: through the heart of the rat to obtain the best possible preservation of the brain for immunohistochemistry. The main advantage of this technique (vs. gravity-fed systems) is that the circulatory system is utilized most effectively.

Here we describe a low-cost, rapid, controlled and uniform fixation procedure using 4% paraformaldehyde perfused via the vascular system: through the heart of the rat to obtain the best possible preservation of the brain.

The goal of fixation is to rapidly and uniformly preserve tissue in a life-like state. While placing tissue directly in fixative works well for small ...

Large-scale Recording of Neurons by Movable Silicon Probes in Behaving Rodents

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of neurons and circuits requires methods with the spatial selectivity and temporal resolution appropriate for mechanistic analysis of neural ensembles in the behaving animal, i.e. recording of representatively large samples of isolated single neurons. Ensemble monitoring of neuronal activity has progressed remarkably in the past decade in both small and large-brained animals, including human subjects1-11. Multiple-site recording with silicon-based devices are particularly effective because of their scalability, small volume and geometric design.
Here, we describe methods for recording multiple single neurons and local field potential in behaving rodents, using commercially available micro-machined silicon probes with custom-made accessory components. There are two basic options for interfacing silicon probes to preamplifiers: printed circuit boards and flexible cables. Probe supplying companies (<a href="http://www.neuronexustech.com/" >http://www.neuronexustech.com/</a>; <a href="http://www.sbmicrosystems.com/">http://www.sbmicrosystems.com/</a>; <a href="http://www.acreo.se/">http://www.acreo.se/</a>) usually provide the bonding service and deliver probes bonded to printed circuit boards or flexible cables. Here, we describe the implantation of a 4-shank, 32-site probe attached to flexible polyimide cable, and mounted on a movable microdrive. Each step of the probe preparation, microdrive construction and surgery is illustrated so that the end user can easily replicate the process.

We describe methods for large-scale recording of multiple single units and local field potential in behaving rodents with silicon probes. Drive fabrication, probe attachment to the drive and probe implantation processes are illustrated in sufficient details for easy replication.

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of ...

A Low Cost Setup for Behavioral Audiometry in Rodents

In auditory animal research it is crucial to have precise information about basic hearing parameters of the animal subjects that are involved in the experiments. Such parameters may be physiological response characteristics of the auditory pathway, e.g. via brainstem audiometry (BERA). But these methods allow only indirect and uncertain extrapolations about the auditory percept that corresponds to these physiological parameters. To assess the perceptual level of hearing, behavioral methods have to be used. A potential problem with the use of behavioral methods for the description of perception in animal models is the fact that most of these methods involve some kind of learning paradigm before the subjects can be behaviorally tested, e.g. animals may have to learn to press a lever in response to a sound. As these learning paradigms change perception itself 1,2 they consequently will influence any result about perception obtained with these methods and therefore have to be interpreted with caution. Exceptions are paradigms that make use of reflex responses, because here no learning paradigms have to be carried out prior to perceptual testing. One such reflex response is the acoustic startle response (ASR) that can highly reproducibly be elicited with unexpected loud sounds in na&iuml;ve animals. This ASR in turn can be influenced by preceding sounds depending on the perceptibility of this preceding stimulus: Sounds well above hearing threshold will completely inhibit the amplitude of the ASR; sounds close to threshold will only slightly inhibit the ASR. This phenomenon is called pre-pulse inhibition (PPI) 3,4, and the amount of PPI on the ASR gradually depends on the perceptibility of the pre-pulse. PPI of the ASR is therefore well suited to determine behavioral audiograms in na&iuml;ve, non-trained animals, to determine hearing impairments or even to detect possible subjective tinnitus percepts in these animals. In this paper we demonstrate the use of this method in a rodent model (cf. also ref. 5), the Mongolian gerbil (Meriones unguiculatus), which is a well know model species for startle response research within the normal human hearing range (e.g. 6).

A fast and inexpensive method for the behavioral determination of hearing parameters like hearing thresholds, hearing impairments or phantom perceptions (subjective tinnitus) is described. It uses pre-pulse inhibition of the acoustic startle response and can be easily implemented in a personal computer using a programmable AD/DA-converter and a piezo sensor.

In auditory animal research it is crucial to have precise information about basic hearing parameters of the animal subjects that are involved in the ...

Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

Deep Brain Stimulation with Simultaneous fMRI in Rodents

In order to visualize the global and downstream neuronal responses to deep brain stimulation (DBS) at various targets, we have developed a protocol for using blood oxygen level dependent (BOLD) functional magnetic resonance imaging (fMRI) to image rodents with simultaneous DBS. DBS fMRI presents a number of technical challenges, including accuracy of electrode implantation, MR artifacts created by the electrode, choice of anesthesia and paralytic to minimize any neuronal effects while simultaneously eliminating animal motion, and maintenance of physiological parameters, deviation from which can confound the BOLD signal.&#160;Our laboratory has developed a set of procedures that are capable of overcoming most of these possible issues. For electrical stimulation, a homemade tungsten bipolar microelectrode is used, inserted stereotactically at the stimulation site in the anesthetized subject. In preparation for imaging, rodents are fixed on a plastic headpiece and transferred to the magnet bore. For sedation and paralysis during scanning, a cocktail of dexmedetomidine and pancuronium is continuously infused, along with a minimal dose of isoflurane; this preparation minimizes the BOLD ceiling effect of volatile anesthetics. In this example experiment, stimulation of the subthalamic nucleus (STN) produces BOLD responses which are observed primarily in ipsilateral cortical regions, centered in motor cortex. Simultaneous DBS and fMRI allows the unambiguous modulation of neural circuits dependent on stimulation location and stimulation parameters, and permits observation of neuronal modulations free of regional bias. This technique may be used to explore the downstream effects of modulating neural circuitry at nearly any brain region, with implications for both experimental and clinical DBS.

This protocol describes a standard method for simultaneous functional magnetic resonance imaging and deep brain stimulation in the rodent. The combined use of these experimental tools allows for the exploration of global downstream activity in response to electrical stimulation at virtually any brain target.

In order to visualize the global and downstream neuronal responses to deep brain stimulation (DBS) at various targets, we have developed a protocol for ...

Limbal Approach-Subretinal Injection of Viral Vectors for Gene Therapy in Mice Retinal Pigment Epithelium

The eye is a small and enclosed organ which makes it an ideal target for gene therapy. Recently various strategies have been applied to gene therapy in retinopathies using non-viral and viral gene delivery to the retina and retinal pigment epithelium (RPE). Subretinal injection is the best approach to deliver viral vectors directly to RPE cells. Before the clinical trial of a gene therapy, it is inevitable to validate the efficacy of the therapy in animal models of various retinopathies. Thus, subretinal injection in mice becomes a fundamental technique for an ocular gene therapy. In this protocol, we provide the easy and replicable technique for subretinal injection of viral vectors to experimental mice. This technique is modified from the intravitreal injection, which is widely used technique in ophthalmology clinics. The representative results of RPE/choroid/scleral complex flat-mount will help to understand the efficacy of this technique and adjust the volume and titer of viral vectors for the extent of gene transduction.

Subretinal injection is a surgical technique for effective gene delivery to retinal pigment epithelium in the mouse eye. Here we describe an easy and replicable method for subretinal injection of viral vectors to retinal pigment epithelium in experimental mice.

The eye is a small and enclosed organ which makes it an ideal target for gene therapy. Recently various strategies have been applied to gene therapy in ...

Quantification of Cerebral Vascular Architecture using Two-photon Microscopy in a Mouse Model of HIV-induced Neuroinflammation

Human Immunodeficiency Virus 1 (HIV-1) infection frequently results in HIV-1 Associated Neurocognitive Disorders (HAND), and is characterized by a chronic neuroinflammatory state within the central nervous system (CNS), thought to be driven principally by virally-mediated activation of microglia and brain resident macrophages. HIV-1 infection is also accompanied by changes in cerebrovascular blood flow (CBF), raising the possibility that HIV-associated chronic neuroinflammation may lead to changes in CBF and/or in cerebral vascular architecture. To address this question, we have used a mouse model for HIV-induced neuroinflammation, and we have tested whether long-term exposure to this inflammatory environment may damage brain vasculature and result in rarefaction of capillary networks. In this paper we describe a method to quantify changes in cortical capillary density in a mouse model of neuroinflammatory disease (HIV-1 Tat transgenic mice). This generalizable approach employs in vivo two-photon imaging of cortical capillaries through a thin-skull cortical window, as well as ex vivo two-photon imaging of cortical capillaries in mouse brain sections. These procedures produce images and z-stack files of capillary networks, respectively, which can be then subjected to quantitative analysis in order to assess changes in cerebral vascular architecture.

This paper describes a method by which the vascular architecture in the brain can be quantified using in vivo and ex vivo two-photon microscopy.

Human Immunodeficiency Virus 1 (HIV-1) infection frequently results in HIV-1 Associated Neurocognitive Disorders (HAND), and is characterized by a chronic ...

Cannula Implantation into the Cisterna Magna of Rodents

Cisterna magna cannulation (CMc) is a straightforward procedure that enables direct access to the cerebrospinal fluid (CSF) without operative damage to the skull or the brain parenchyma. In anesthetized rodents, the exposure of the dura mater by blunt dissection of the neck muscles allows the insertion of a cannula into the cisterna magna (CM). The cannula, composed either by a fine beveled needle or borosilicate capillary, is attached via a polyethylene (PE) tube to a syringe. Using a syringe pump, molecules can then be injected at controlled rates directly into the CM, which is continuous with the subarachnoid space. From the subarachnoid space, we can trace CSF fluxes by convective flow into the perivascular space around penetrating arterioles, where solute exchange with the interstitial fluid (ISF) occurs. CMc can be performed for acute injections immediately following the surgery, or for chronic implantation, with later injection in anesthetized or awake, freely moving rodents. Quantitation of tracer distribution in the brain parenchyma can be performed by epifluorescence, 2-photon microscopy, and magnetic resonance imaging (MRI), depending on the physico-chemical properties of the injected molecules. Thus, CMc in conjunction with various imaging techniques offers a powerful tool for assessment of the glymphatic system and CSF dynamics and function. Furthermore, CMc can be utilized as a conduit for fast, brain-wide delivery of signaling molecules and metabolic substrates that could not otherwise cross the blood brain barrier (BBB).

Here we describe a protocol to perform cisterna magna cannulation (CMc), a minimally invasive way to deliver tracers, substrates and signaling molecules into the cerebrospinal fluid (CSF). Combined with different imaging modalities, CMc enables glymphatic system and CSF dynamics assessment, as well as brain-wide delivery of various compounds.

Cisterna magna cannulation (CMc) is a straightforward procedure that enables direct access to the cerebrospinal fluid (CSF) without operative damage to ...

The Power of Interstimulus Interval for the Assessment of Temporal Processing in Rodents

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in neurocognitive disorders. Despite the popularization of prepulse inhibition (PPI) in recent years, many current protocols promote using a percent of control measure, thereby precluding the assessment of temporal processing. The present study used cross-modal PPI and gap prepulse inhibition (gap-PPI) to demonstrate the benefits of employing a range of interstimulus intervals (ISIs) to delineate effects of sensory modality, psychostimulant exposure, and age. Assessment of sensory modality, psychostimulant exposure, and age reveals the utility of an approach varying the interstimulus interval (ISI) to establish the shape of the ISI function, including increases (sharper curve inflections) or decreases (flattening of the response amplitude curve) in startle amplitude. Additionally, shifts in peak response inhibition, suggestive of a differential sensitivity to the manipulation of ISI, are often revealed. Thus, the systematic manipulation of ISI affords a critical opportunity to evaluate temporal processing, which may reveal the underlying neural mechanisms involved in neurocognitive disorders.

Temporal processing, a preattentive process, may underlie deficits in higher-level cognitive processes, including attention, commonly observed in neurocognitive disorders. Using prepulse inhibition as an exemplar paradigm, we present a protocol for manipulating interstimulus interval (ISI) to establish the shape of the ISI function to provide an assessment of temporal processing.